Phytoremediation of nitrate and bisphenol a by scindapsus auresus

Chapter 1. Introduction Phytoremediation is a technology that utilizes plants and associated rhizopheric microorganisms to remove pollutants and ‘clean’ contaminated soils and water (Dietz and Schnoor, 2001; Singh et al., 2003). This technology has gained the interest of government agencies and industries in recent years (Pilon-Smits, 2005), due to a number of reasons. With plants playing the major role, most of their processes are sun-dependent, making phytoremediation very much cheaper than the various engineering methods used at present to treat contaminated areas (Salt et al., 1998; Pilon-Smits, 2005). Also, phytoremediation can be carried out at the site of contamination, thus reducing exposure of humans and wildlife to toxins and contaminants (Pilon-Smits, 2005). Lastly, the use of plants helps to project an environmentally-friendly image as opposed to chemical plants (Pilon-Smits, 2005) and, depending on the species used, the plants may also be aesthetically pleasing (Dietz and Schnoor, 2001), hence gaining popularity with the general public. Due to the many advantages of phytoremediation, ongoing research is taking place to better develop this technology. These studies not only focus on finding plant species capable of phytoremediating the contaminated sites, but also on identifying the exact mechanisms, genes, enzymes and physiological processes involved. Nitrate (NO3-) pollution of surface water has been a major problem in several countries. NO3- is one of the two forms of inorganic N introduced excessively into the environment via anthropogenic sources such as agricultural run-offs and point-source discharges of sewage effluents and industrial wastewater (Camargo and Alonso, 2006; Maier et al., 2009; O’Shea and Wade, 2009). High levels of NO3- pollution of surface water are hazardous to the environment, causing eutrophication of freshwater and 1 marine ecosystems (Camargo and Alonso, 2006; Maier et al., 2009), and contamination of groundwater (Rivett et al., 2008; Umezawa et al., 2008). NO3- at high concentrations can easily be converted into nitrite (NO2-), causing health problems such as methaemoglobinaemia and gastro-intestinal cancers (Camargo and Alonso, 2006; Hsu et al., 2009). Therefore, there is a pressing need to come up with new methods and technologies to remove and/or control NO3- levels in surface water. Another pollutant that is a major problem in several developed countries is the organic chemical, bisphenol A (BPA). BPA (CAS Registry No. 80-05-7), also known as 4,4’-isopropylidenediphenol, is commonly used in the production of polycarbonate plastic and as a plasticizer (Staples et al., 1998; Sajiki and Yonekubo, 2003). The exposure of BPA-containing plastic to heat, acidic or alkaline conditions as well as the duration of the exposure to these conditions can lead to the release of BPA into the environment, risking contact of humans and wildlife to it (López-Cervantes et al., 2003; Kang et al., 2006a; Richter et al., 2007). High BPA levels have been detected in wastewater from sewage treatment plants and manufacturing factories, as well as leachates from water landfills (Staples et al., 1998; Yamamoto et al., 2001; Fromme et al., 2002; Kang and Kondo, 2006). These effluents result in BPA contamination of the aquatic environment. Several studies have shown that aquatic organisms can absorb and accumulate BPA (Staples et al., 1998; Heinonen et al., 2002; Li et al., 2009b). The bioaccumulated BPA can then make its way up the food chain (Muñoz et al., 1996; Li et al., 2009b) and into humans via aquatic food sources (Basheer et al., 2004). BPA has been shown to act as an endocrine disrupting chemical by mimicking the effects of estrogen (Kang et al., 2006a; Richter et al., 2007). Studies have shown that exposure of mice to BPA can lead to advanced puberty (Howdeshell et al., 1999; 2 Ryan and Vandenbergh, 2006), increased aggression and anxiety (Kawai et al., 2003; Ryan and Vandenbergh, 2006), as well as upregulated expression of several hormone receptors (Ramos et al., 2003). Therefore, there is also an urgent need to remove and/or degrade BPA from the environment, in order to decrease the potential harm of BPA to the aquatic ecosystems and even to humans. Although, at present, there are several existing engineering and chemical methods available to remove NO3- and BPA from water bodies, these methods involve the construction of elaborate infrastructures, usage of excessive chemicals and transportation of the polluted waters away from the source, resulting in high monetary costs and the risk of public exposure to the polluted waters (Pilon Smits, 2005). As compared to these existing methods, phytoremediation is much more desirable because the use of plants allows for on-site bioremediation of the contaminated waters without incurring high costs (Pilon Smits, 2005). This project focused on the use of Scindapsus aureus (Lindl. & André) Engl. (Golden Pothos) in the phytoremediation of NO3- and BPA contaminated waters. Characteristics of S. aureus that make it a potential study candidate for phytoremediation of water bodies include its tolerance to submergence of roots in water, high growth rate under submerged conditions and easy propagation via shoot cuttings. The ability of S. aureus to grow in water allows for phytoremediation in situ. High growth rate and easy propagation are both indicators of efficient phytoremediating plant species (Brix and Schierup, 1989; Forni et al., 2001). Furthermore, S. aureus is sold in the market as an ornamental house-plant, indicating that it has both aesthetic and commercial values. Hence, the use of S. aureus in phytoremediation may bring along additional aesthetic and commercial values. 3 Aside from determining the phytoremediation potential of S. aureus, it is also necessary to study the physiological impact of high NO3- and BPA levels on the plant to ensure that this species can be used for long-term phytoremediation of these pollutants without causing detrimental effects to the ecosystem or the plant itself. Therefore, the objectives of this study included the determination of the phytoremediation efficiency of S. aureus in removing NO3- and BPA from water as well as the physiological responses of S. aureus to high NO3- and BPA levels, in terms of plant growth and metabolism, photosynthetic capacity, oxidative stress levels and the activities of antioxidant enzymes, including ascorbate peroxidase, glutathione reductase, superoxide dismutase and guaiacol peroxidase. 4 Chapter 2. Literature Review 2.1 Phytoremediation Phytoremediation refers to the use of plants and their associated microbes for the cleanup of environmental pollution (Pilon-Smits, 2005). This technology has been under extensive research in recent years due to a number of benefits that it brings. As mentioned in the introduction chapter, phytoremediation has captured the attention of several government agencies and industries because if carried out on a full scale, it can be more cost-effective and is potentially safer than existing physical and chemical methods (Salt et al., 1998; Pilon-Smits, 2005). Arthur et al. (2005) reported that current price estimates for remediation of full-scale commercial sites could begin at US$ 200,000, with an additional US$40 – 70 per cubic yard of soil. In contrast, the use of plants is solar-dependant, which makes the process of decontamination much cheaper. Also, the use of plants is environmentally-friendly and aesthetically pleasing, which makes phytoremediation a publically acceptable way to remove environmental pollutants (Arthur et al., 2005; Pilon-Smits, 2005). Phytoremediation can be used to treat soils and sediments, water bodies and even the surrounding atmosphere (Pilon-Smits, 2005). A large number of inorganic pollutants, such as plant macronutrients (nitrates and phosphates, for instance), heavy metals and radioactive isotopes, as well as organic pollutants, such as polyaromatic hydrocarbons (PAH), explosives (TNT), petrochemicals, polychlorinated biphenyls (PCBs) and bisphenol A (BPA) can be phytoremediated (Arthur et al., 2005; PilonSmits, 2005). Also, through research, a large number of botanical families with phytoremediating capabilities have been identified (Gawronski and Gawronska, 2007) and several databases such as PHYTOPET and PHYTOREM have compiled lists of 5 plants that can specifically remediate petroleum hydrocarbons and heavy metals, respectively (Pilon-Smits, 2005). Plants can treat pollutants in a number of ways (Fig. 1). First, pollutants can be extracted by the plants and accumulated in the tissues, after which the plants can be removed together with the pollutants – a process termed phytoextraction (Arthur et al., 2005; Pilon-Smits, 2005). For example, Murakami and Ae (2009) reported the ability of Oryza sativa (rice) and Zea mays (corn) to phytoextract copper, and the ability of Glycine max (soybean) to phytoextract zinc. Second, plants can also take up the contaminants and degrade them into harmless compounds via enzymatic activities within their tissues, a process known as phytodegradation (Arthur et al., 2005; PilonSmits, 2005). Reed wetlands have been reported to be able to phytodegrade extra heavy oil hydrocarbons without much impact on the long-term reed yield of the wetlands (Ji et al., 2004). Third, contaminants absorbed into plant tissues can be converted to volatile forms and eventually leave the plant through a process called phytovolatilization (Arthur et al., 2005; Pilon-Smits, 2005). Pilon-Smits et al. (1998) reported the volatilization and assimilation of selenium by Populus tremula x alba). Fourth, the roots of certain plant species can convert contaminants to less bioavailable forms in order to prevent leaching or runoff in a process termed phytostabilization (Arthur et al., 2005; Pilon-Smits, 2005). For instance, Quercus ilex (Holm oak) was not only tolerant to high levels of cadmium, but the fine roots of the plants were also able to retain the metal within the soil without translocation into the leaves (Domínguez et al., 2009). Lastly, plants can also stimulate microbial growth and activity at the rhizospheric region to induce degradation of contaminants by these microbes. This process is termed as phytostimulation (Arthur et al., 2005; Pilon-Smits, 6 Fig. 1. Potential fates of pollutants (black circles) during phytoremediation. Pollutants can be stabilized or degraded in the rhizosphere, accumulated or degraded within plant tissues, or volatilized. (Figure modified from Pilon-Smits, 2005). 7 2005). Toyama et al., (2006) reported that degradation of organic chemicals (phenol, aniline and 2,4-dichlorophenol) by Spirodela polyrrhiza (giant duckweed) was accelerated by the selective accumulation of certain bacterial strains in its rhizosphere. Similarly for Phragmites australis, the degradation of bisphenols was accelerated by the interactions between the plants and bisphenol degrading bacteria, Novosphingobium sp. strain TYA-a and Sphingobium yanoikuyae strain TYF-1 Toyama et al., (2009). Studies on phytoremediation are mainly targeted at the potential of different plants to remove the target pollutant from the substrate and on the practicality of applying this technology out in the field (Dushenkov et al., 1995; Newman et al., 1999; Huang et al., 2000; Sooknah and Wilkie, 2004; Novak and Chan., 2008). Yet, the ability to fully utilize the potential of plants for phytoremediation is restricted by limited understanding of plant metabolism and tolerance mechanisms (Doran, 2009). Therefore, in recent years, more research is extended towards comprehending the biological pathways behind phytoremediation potentials of the plants (Schrӧder and Collins, 2002; Singer et al., 2003; Singh et al., 2003). The increased use of plant tissue culture techniques in the study of phytoremediation has assisted in the understanding of the responses and metabolic pathways of plants towards various pollutants up to the cellular level. Plant tissue culture involves the growth of plant cells and tissues in vitro under aseptic conditions and because cells and tissues are grown under a microbe-free environment, the physiological responses and metabolic capabilities of plant cells can be clearly distinguished from those of microorganisms commonly present in plant tissues or rhizospheric regons (Doran, 2009). The form of tissue culture most commonly utilized in phytoremediation studies are cell 8 suspensions and hairy roots (Doran, 2009), but whole plants have also been employed (Imai et al., 2007). Aside from physiological and metabolic studies, a lot of focus has also been placed on molecular responses towards plant uptake of pollutants. Plant molecular ecotoxicology investigates the ecological implications of genetic and molecular responses to contaminants and helps in the understanding of relationships between plant genotypes and ecological phenotypes, the regulatory networks formed by secondary messenger, as well as transcriptional and post-transcriptional events induced by exposure to certain pollutants (Sandermann Jr., 2004). Also, several transgenic plants with enhanced abilities to degrade xenobiotics (Eapen et al., 2007) and sequester or detoxify heavy metals (Yang et al., 2005) have been developed using genetic engineering. 2.2 Phytoremediation of Nitrates 2.2.1 Nitrate Pollution Nitrate (NO3-) is one of the main forms of inorganic nitrogen (N) introduced excessively into the environment via anthropogenic sources. It has been reported that current human activities have severely altered the global nitrogen cycle, and the rate of N input is still increasing (Camargo and Alonso, 2006). The most widely known cause of NO3- pollution is agricultural run-offs, of which farmers use large amounts of NO3- fertilizers to boost their crop growth and the excess NO3- leach into the surrounding water bodies, contaminating them (Kurosawa et al., 2004; Camargo and Alonso, 2006; Cherry et al., 2008; Maier et al., 2009; O’Shea and Wade, 2009). In 9 urban regions, NO3- pollution of surface waters is also prevalent. Point source discharges of sewage effluents and industrial wastewater contribute to high NO3levels detected in the waterways of urbanized cities (Forni et al., 2001; Camargo and Alonso, 2006; Li et al., 2007; Maier et al., 2009; O’Shea and Wade, 2009). The risks that NO3- pollutant cause to human health and the environment have resulted in an array of legislations set by governments and organizations all over the world to control NO3- levels in surface water bodies (O’Shea and Wade, 2009). In the UK, the Drinking Water Directive set a maximum limit of 11.3 mg-N/L in drinking water (O’Shea and Wade, 2009). The World Health Organization and Europeon Union also set a rule to maintain water NO3- levels below 50 mg/L, with a limit cap at 100 mg/L (World Health Organization, 2006; Umezawa et al., 2008). In Japan and the United States, federal standards are set at 10 mg-N/L (Umezawa et al., 2008). In Singapore, the Public Utilities Board (PUB) has also set a regulatory guideline to maintain NO3- levels in drinking water to be below 45 mg/L (World Health Organization, 2006). High levels of NO3- pollution of surface water bring adverse effects to the environment. Eutrophication of water bodies is the result of the release of surplus NO3- into surface waters. Highly eutrophic waters lead to undesirable algal blooms, which may contain toxic species. Eutrophication can drastically affect the structure and function of freshwater, marine and terrestrial ecosystems through poisoning, loss of keystone aquatic species, or even the loss of light and oxygen supply in water (Camargo and Alonso, 2006; Maier et al., 2009; O’Shea and Wade, 2009). NO3- from surface waters can even enter groundwater, degrading the quality of human drinking water (Rivett et al., 2008; Umezawa et al., 2008). Furthermore, NO3- at high 10 concentrations can easily be converted into nitrite (NO2-). The ingestion of NO2--rich water can cause methaemoglobinaemia or ‘blue-blood syndrome’ in infants (Camargo and Alonso, 2006). This condition can be fatal if not detected early and treated. Ingested NO2- may be converted into N-nitroso compounds, which are potent carcinogens suspected to cause gastro-intestinal cancers (Camargo and Alonso, 2006; Hsu et al., 2009). 2.2.2 Methods for Phytoremediation of Nitrate from Water Due to the hazardous effects that NO3- can bring, several technologies have been developed in order to remove it from drinking water and various water bodies. These techniques include the use of activated carbon to remove NO3- (Demiral and Gündüzoğlu, 2010), reverse osmosis (Schoeman and Steyn, 2003), ion exchange (Samatya et al., 2006), electrodialysis and chemical adsorption (Menkouchi Sahli et al., 2008) and even microbial treatment (Ayyasamy et al., 2007). These methods may be effective in the removal of NO3-, but they are expensive to set up on a big scale. (Kapoor and Viraraghavan, 1997; Ayyasamy et al., 2009). Also, microbial treatments require external supplies of organic carbon and nutrients, which may not be fully consumed and can become secondary pollutants, hence requiring extensive post treatments to remove (Ayyasamy et al., 2009). Taking all these limitations into consideration, phytoremediation appears as a more preferred alternative method for NO3- removal. There are several different approaches for the phytoremediation of NO3-. One approach is the use of floating beds made up of aquatic macrophytes for NO3- phytoremediation. Several floating aquatic plant species such as Azolla filiculoides (Forni et al., 2001), Eichhornia crassipes, Pistia stratiotes (Sooknah and Wilkie, 2004; Ayyasamy et al., 2009), Hydrocotyle 11 umbellate (Sooknah and Wilkie, 2004), Salvinia molesta (Ayyasamy et al., 2009), Salvinia minima and Spirodela polyrrhiza (Olguín and Sánchez-Galván, 2007) have been assessed for their abilities to remove NO3- from organic wastewater and groundwater. These plants were able to phytoremediate NO3- from concentrations up to 500 mg/L, although the efficiency of NO3- removal decreased during exposure to 500 mg/L NO3- concentrations (Ayyasamy et al., 2009). However, several of the aquatic plants studied (e.g. Eichhornia crassipes) are considered weeds in freshwater bodies and have been known to rapidly colonize freshwater bodies, resulting in the collapse of aquatic eco-populations (Li et al., 2007). The large-scale use of these plants for NO3- phytoremediation may not be beneficial in the long term. The use of the root systems of terrestrial plants to create a rhizofitration system of floating beds for the removal of NO3- from water has also been studied. Monnet et al. (2002) has reported the use of horticultural roses to treat domestic wastewater and observed that the system was efficient in the removal of inorganic N and phosphate (P). Li et al. (2007; 2009) also used Ipomoea aquatica to remove NO3from eutrophic water and even utilized low energy N+ ion implantation as a mutagen to increase the growth and nitrogen-removal efficiency of these plants. However, these methods require high cost to set up in a big scale as extensive infrastructure and equipment are required. Another approach is the use of constructed wetlands, made up of emergent plants such as Phragmites australis (Huett et al., 2005), Scirpus cyperinus and Typha latifolia (Huang et al., 2000). The roots of these emergent plants were found to be efficient absorbers of NO3- (Huang et al., 2000; Huett et al., 2005). In China, phytoremediation projects involving constructed wetlands have already been 12 established and one such project in the Hangzhou Botanical Garden has indeed greatly improved the quality of the groundwater resource (Yang et al., 2008). Despite the effectiveness of the various phytoremediation methods developed for the removal of NO3-, they still have their limitations with respect to fieldapplications. The efficiency of NO3- removal is highly dependent upon genetic and environmental factors and these are attributed to the choice of plant species and the way in which they are utilized (Li et al., 2007). Therefore, further studies to identify new plant species and improve on the setup are required to elevate the efficiency of NO3- removal and promote field-applications. 2.2.3 Nitrate Absorption and Assimilation In order to study the phytoremediation mechanisms of plants, it is essential to first understand how plants absorb and assimilate NO3-. As NO3- is considered a ratelimiting nutrient for plant growth (Redinbaugh and Campbell, 1991; Leleu et al., 2000), the concentration of NO3- available to plants can lead to several physiological, biochemical and even molecular changes in the plants. Deficiencies in NO3--N can lead to decrease in leaf growth (Lawlor et al., 2001), increase in root: shoot biomass ratio (Ingestad and Ågren, 1991; Améziane et al., 1997; Touraine et al., 2001; Hermans et al., 2006), decreased photosynthetic capacity (Khamis et al., 1990; Skillman and Osmond, 1998) and accumulation of carbohydrates in leaves (Scheible et al., 1997; Hermans et al., 2006). NO3- Metabolism NO3- influx into roots is an active process, driven by the H+ gradient (Forde, 2002), which explains the alkalization of the surrounding medium in which the plant 13 grows in (Tischner and Kaiser, 2007). NO3- uptake involved two or more H+ cotransported with every NO3- (Crawford, 1995). Kinetic experiments on net NO3uptake have provided evidence of two distinct transport components to the NO3uptake mechanism, namely the high-affinity transport systems (HATS) and lowaffinity transport systems (LATS) (Grignon et al., 2001; Touraine et al., 2001). Both LATS and HATS are strongly dependent on the NO3- concentration available and the plant internal N demand. HATS are capable of efficiently transporting NO3- even when its concentration in the medium is extremely low, in the range of just 10 µM (Grignon et al., 2001). There are two different HATS systems – a low capacity constitutive high-affinity transport system (cHATS) and a high capacity inducible high-affinity transport system (iHATS) (Touraine et al., 2001). cHATS plays an important role in allowing the cytoplasmic concentrations of NO3- to rise to levels enough to induce iHATS (Touraine et al., 2001). iHATS is a multicomponent system, partly encoded by genes of the NRT2 family (Forde, 2000). In the absence of NO3-, NRT2 genes are expressed at very low levels in roots but they are induced within minutes upon NO3- reapplication (Forde, 2002). However, at a much higher NO3concentration (mM range), HATS activity plateaus off and LATS become induced (Grignon et al., 2001; Touraine et al., 2001). Independent of NO3-, LATS appears to be constitutively induced (Forde, 2000), but is subjected to other forms of regulation, such as the internal plant N demand (Touraine et al., 2001). NO3- uptake, in general, can be regulated by glutamine as well as by NO3- itself (Tischner and Kaiser, 2007). If plants are exposed to high NO3- concentrations over long periods of time, NO3- uptake can be inhibited as uptake rates match the growth rate of the plant and storage pools become overloaded (Tischner and Kaiser, 2007). 14 Upon uptake into the plant cells, NO3- is reduced to NO2- by nitrate reductase (NR) in the cytosol (Sechley et al., 1992; Tischner and Kaiser, 2007). NO2- then enters the plastid (chloroplasts in shoots) to be reduced to NH4+ by nitrite reductase (NiR) (Sechley et al., 1992; Tischner and Kaiser, 2007). NH4+-N is then assimilated via the GS-GOGAT pathway into glutamine/glutamate, which are substrates for transamination reactions to produce other amino acids and proteins (Tischner and Kaiser, 2007). The assimilation of NO3- is summarized in the diagram as shown in Fig. 2. Nitrate Reductase (NR) NR catalyses the first step of NO3- assimilation, and is also a rate-limiting process in NO3- uptake in plants (Campbell, 1999). NR can be found in shoot mesophyll cells and root epidermal and cortical cells, but activity levels in either region differ depending on plant species (Sechley et al., 1992; Crawford, 1995). Higher plant NR catalyses the following reaction: NO3- + NAD(P)H + H+  NO2- + NAD(P)+ + H2O In higher plants, two forms of NR exist, the more common form, NADHspecific NR (EC 1.6.6.1) and a second isoform, NAD(P)H-bispecific NR (EC 1.6.6.2) (Ahmad and Abdin, 1999; Meyer and Stitt, 2001). The NR protein contains three domains, namely FAD, a haem cofactor and a molybdenum cofactor, all involved in the transfer of electrons from NAD(P)H to NO3- (Campbell, 1999; Meyer and Stitt, 2001). It is well-known that NR activity increases with increasing NO3- supply (Hageman and Flesher, 1960; Ivashikina and Sokolov, 1997; Chen et al., 2004; Taghavi and Babalar, 2007). The increase in NR activity upon exposure to NO3- is 15 Fig. 2. Schematic diagram of NO3- uptake and assimilation pathway. NO3- is transported through the plasma membrane via ATPase proton pumps (P), NO3transporters (T) and channels. NO3- is then reduced into NO2- by nitrate reductase (NR) in the cytosol, after which NO2- is transported into the plastid (chloroplast in leaves) and further reduced into NH4+ by nitrite reductase (NiR). NH4+ is then converted into glutamine via the GS-GOGAT pathway. NO3- in the cytosol is also either translocated to other plant organs via the xylem or accumulated within the vacuoles of the cell. (Diagram redrawn from Grignon et al., 2001). 16 due to an increase in mRNA levels (Crawford et al., 1986; Faure et al., 2001). Induction of NR expression by NO3- can occur within minutes of NO3- exposure, and only requires NO3- concentrations of less than 10 µM (Meyer and Stitt, 2001). When faced with NO3- starvation, both NR activity and protein levels decrease rapidly, but NR mRNA levels can still remain constant for more than 10 days – a result of posttranslational control of protein degradation by proteases or a decrease in mRNA translation (Faure et al., 2001). Aside from NO3- concentrations, several other plant and environmental factors can affect NR activity such as age of the leaves (Kenis et al., 1992), cytokinins (Lu et al., 1992; Suty et al., 1993) , light intensity (Hageman and Flescher, 1960; Hoff et al., 1994; Crawford, 1995; Chow et al., 2004), CO2 availability (Kaiser and Huber, 2001), plant photosynthetic rate (Kaiser and Huber, 2001; Meier and Stitt, 2001), carbohydrate content (Hoff et al., 1994; Crawford, 1995), anoxia (Botrel et al., 1996; Morad et al., 2004) and even salt stress (Surabhi et al., 2008). 2.3 Phytoremediation of Bisphenol A (BPA) 2.3.1 BPA Pollution Bisphenol A (BPA) is a monomer used in the preparation of epoxide resins, polycarbonate plastics, and as a stabilizer in polyvinylchloride – plastics commonly used to make packaging materials (Staples et al., 1998; Sajiki and Yonekubo, 2003). Prolonged exposure of these plastics to heat, acidic or alkaline conditions can lead to the release of BPA into the environment, resulting in exposure of humans and wildlife 17 to this contaminant (López-Cervantes et al., 2003; Kang et al., 2006a; Richter et al., 2007). BPA pollution of water is a major problem in several countries, particularly in the urbanized regions. Sources of BPA contamination come from wastewater released by sewage treatment plants and manufacturing factories, as well as leachates from water landfills (Staples et al., 1998; Yamamoto et al., 2001; Fromme et al., 2002; Kang et al., 2006a). From these sources, BPA can then make its way into freshwater and marine bodies, and even into groundwater. In Japan, BPA pollution is widespread, as reported by the Environmental Agency of Japan (Yamamoto et al. 2001). Levels of BPA detected in freshwater and seawater samples collected throughout Japan ranged from 0.010 to 0.268 µg/ L, whereas in leachates of hazardous waste landfills, BPA levels can range from 1.3 – 17, 200 µg/L (Yamamoto et al. 2001). BPA pollution is also a problem in Europe. Fromme et al. (2002) collected 116 surface-water samples and 39 sewage effluent samples throughout Germany for analysis and reported that BPA levels in surface water ranged from 0.0005 – 0.41 µg/L, whereas BPA levels in sewage effluents ranged from 0.018 – 0.702 µg/L. Kolpin et al. (2002) also reported that BPA concentrations in river waters of USA ranged from 0.14 – 12.00 µg/L. Even within Singapore, Basheer et al. (2004) reported the detection of BPA (1 – 2 ng/L) in the surface seawater. BPA present in the environment is disastrous because organisms can absorb and bioaccumulate BPA, thereby introducing it into the food chain (Muñoz et al., 1996; Staples et al., 1998; Heinonen et al., 2002; Li et al., 2009). Basheer et al. (2002) analysed several aquatic food sources available in Singapore supermarkets and detected BPA [13.3 – 213.1 ng/g (w/w)] in prawns, crabs, blood cockle, white clam, 18 squid and fish. Shao et al. (2007) also detected BPA [0.27 – 1.01 µg/kg (w/w)] in pork, mutton, beef and fish sold in Beijing supermarkets. The consumption of food sources contaminated with BPA and the ability to bioaccumulate BPA puts human health at risk of BPA toxicity. Realising the potential toxicity of BPA to humans, the United States Environmental Protection Agency (EPA) set a maximum acceptable dose for BPA at 0.05 mg/kg body weight/day, and the European Union (EU) has also established a temporary tolerable daily intake (TDI) for BPA of 10 µg/kg body weight (Kang et al., 2006a). Endocrine Disrupting Properties of BPA BPA has been shown to act as an endocrine disrupting chemical (EDC) by mimicking the effects of estrogen (Kang et al., 2006a; Richter et al., 2007). Daily exposure of BPA (even in low doses of 50 mg/kg/day) to animals can cause adverse physiological, hormonal, reproductive and even behavioural changes (Richter et al., 2007). In male rats, daily exposure to low levels of BPA can cause oxidative stress in the liver (Bindhumol et al., 2003), kidneys (Kabuto et al., 2003), striatum (Obata and Kubota, 2000) and epididymal sperm (Chitra et al., 2003), permanent alterations in the male prostate and hypothalamic-pituitary-gonadal axis (Ramos et al., 2003), as well as increased aggression and anxiety (Kawai et al., 2003; Ryan and Vandenbergh, 2006). In female rats, daily exposure to BPA can disrupt the development of nervous systems, sexual dimorphic behaviours and increased anxiety (Ryan and Vandenbergh, 2006) as well as advanced puberty (Howdeshell et al., 1999). ). It has also been demonstrated in pregnant rats that BPA absorbed into the body could be passed on to fetuses through the placenta (Takahashi and Oishi, 2000). This problem is particularly crucial as fetuses have low endogenous concentrations of UDP19 glucuronosyltransferase (UGT), the enzyme responsible for detoxification of BPA in the liver (Kang et al., 2006a). Although there is a lack of studies done on the endocrine disrupting properties of BPA on humans, the risk of BPA toxicity to humans is still present as BPA exposure is prevalent. There is hence still a need for further research on the detoxification and removal of BPA from the environment. 2.3.2 Methods for Phytoremediation of BPA from Water Abiotic degradation of BPA via photolysis and free radical oxidation are possible (Staples et al., 1998; Sajiki and Yonekubo, 2003) but the degradation rates have been shown to be much lower compared to biotic degradation (Imai et al., 2007). Espinoza et al. (2007) attempted to enhance solar UV-degradation of BPA via the addition of NO3-, Fe (III) and HCO3-. However, the addition of these ions to the water may cause other negative effects to the water body under remediation, such as eutrophication. Microbial degradation of BPA has been detected in river water (Kang and Kondo 2002a; 2002b) and wastewater treatment plants (Spivak et al., 1994), and BPA degradation rates were relatively high (Kang et al., 2006b). However, not all bacterial strains have high BPA biodegradability rates. Kang and Kondo (2002a) isolated 10 bacterial strains with BPA biodegradability from river waters, but only a Pseudomonas sp. and a Pseudomonas putida strain were able to biodegrade >90% of BPA within 10 days. Furthermore, BPA degradation by microbes are affected by temperature, bacterial counts (Kang and Kondo, 2002b), as well as aerobic and anaerobic conditions (Kang and Kondo, 2002a). 20 In comparison to the above methods, phytoremediation of BPA appears to be more desirable. As a result, in recent years, research on phytoremediation of BPA has been gaining popularity, with biodegradation studies involving plant enzymes, plant cell suspension cultures, rhizospheric degradation and even direct uptake into whole plants. Several different plant enzymes can degrade BPA. Even crude enzyme extracts were reported to be able to degrade BPA (Xuan et al., 2002; Imanaka et al., 2005; Kang and Kondo, 2006). Among them all, the most commonly studied enzyme is peroxidase. Peroxidases extracted from Glycine max (soybean) (Caza et al., 1999), Momordica charantia (bitter gourd) (Karim and Husain, 2009), and Cochlearia armoracia (horseradish) (Sakuyama et al., 2003) have shown to be able to oxidize BPA. Yoshida et al. (2002) also reported that polyphenol oxidases extracted from vegetables (e.g. potato, mushroom, eggplants, edible burdock and yacon) were able to oxygenate BPA to quinones. Fungal laccases extracted from Trametes villosa (Fukuda et al., 2001) also could polymerise and degrade BPA from liquid media. As mentioned above, tissue culture techniques have aided tremendously in the understanding of mechanisms behind phytoremediation. The exact pathways behind BPA biodegradation by plants are still relatively unknown. However, the use of plant cell suspension cultures has revealed some insights on plant BPA degradation. Nakajima et al. (2002, 2004) intensively utilized suspension cultures of Nicotiana tabacum BY-2 cells to uncover the mechanism behind plant processing of BPA. They discovered that BPA was glycosylated by the plant cells into several secondary complex glycosides, which increased the solubility of BPA – an integral process of detoxification – and removed the endocrine disrupting property of the original 21 chemical (Nakajima et al., 2002, 2004). Cell suspension cultures of Eucalyptus perriniana (Hamada et al., 2002), Glycine max, Triticum aestivum, Digitalis purpurea and Datura stramonium (Schmidt and Schuphan, 2002) were also reported to be able to glycosylate and hydroxylate BPA into complex glycosides, hence removing it from the liquid growth media. Suspension cultures of freshwater microalgal species (Nakajima et al., 2007) and marine microalgae species (Li et al., 2009b) were also reported to be able to remove BPA from water bodies. Despite the knowledge obtained from the use of enzymes and cell suspension cultures, field-based application of this technology is extremely difficult and expensive. In contrast, the use of whole plants to phytoremediate BPA from water bodies is much more feasible. Therefore, research to identify potential plant species capable of phytoremediating BPA from water is intensively carried out. Several aquatic macrophytes have been identified for the phytoremediation of BPA from water. Eichhornia crassipes (Kang and Kondo, 2006) and Ipomoea aquatica (Noureddin et al., 2004) were able to remove BPA from water. Toyama et al. (2009) found out that Phragmites australis was able to phytoremediate BPA from river sediments via phytostimulation of Novoshingobiam sp. strain TYA-1 at the rhizosphere. Ferrara et al. (2006) reported that various hydroponic seedlings (Vicia faba, Lycopersicon esculentum, Triticum durum and Lactuca sativa) were able to remove BPA from hydroponic growth solutions. Imai et al. (2007) went one step further and grew Portulaca oleracea plants under aseptic conditions to show that the plant was able to phytoremediate BPA, independent of microbial influence. P. oleracea is also an ornamental plant, which provided an additional benefit of being aesthetically pleasing, if it was ever utilized in field-based applications. 22 Phytoremediation of BPA is relatively less researched on, compared to other organic pollutants such as pesticides and other polyaromatic hydrocarbons (PAHs). Further research will be required not only to identify new plant species that are more efficient in the phytoremediation of this pollutant, but also to better understand the processes behind the biodegradation of BPA. 2.4 Plant Stress Responses Stress is defined as an abiotic and biotic factor/environment modified in a manner (excess or deficit) in which it has the capability of causing injury, disease or aberrant physiology to the organism (Gaspar et al., 2002). As plants are generally restricted to the place where they grow, it is difficult for them to avoid unfavourable changes in their environment, such as nutrient deficiency and environmental pollution. Therefore, they have developed several mechanism/ strategies to defend themselves against these biotic and abiotic stresses, and these mechanisms/ strategies are often combined with an alteration of growth and developmental patterns in plants (Gaspar et al., 2002). A stress response is usually initiated when the plant recognises a stress factor at the cellular level (Gaspar et al., 2002). Several physiological changes within the plant can take place and the degree of change is dependent upon the duration, severity and rate at which the stress is imposed. Some of the changes that are known to occur include alterations in photosynthetic capacity (Smillie and Hetherington, 1983; Ralph, 2000; Rutherford and Krieger-Liszkay, 2001), carbon (sugar) balance (Ehness et al., 1997; Sheen et al., 1999; Saladin et al., 2003) and soluble protein concentrations (Saladin et al., 2003; Ashraf and Harris, 2004). With respect to phytoremediation, the study of plant stress responses can give a better understanding of the degree of stress 23 induced in the plants upon exposure to various pollutants and this will help in determining whether the plant under study is able to build tolerance towards the pollutant to allow for long-term phytoremediation. One of the responses towards environmental stress is the production of reactive oxygen species (ROS) (Gaspar et al., 2002). ROS comprises of a group of highly reactive molecules, which include superoxide radical (O2-.), hydrogen peroxide (H2O2), hydroxyl radical (.OH) and singlet oxygen (1O2) (Mach and Greenberg, 2004). A build-up of ROS concentrations within the plant can lead to oxidative stress. ROS are formed during certain redox reactions, during photorespiration and glyoxylation within peroxisomes, as well as during the incomplete reduction of oxygen or oxidation of water by the mitochondrial or chloroplast electron transport chains (Gaspar et al., 2002; Mach and Greenberg, 2004). The reactions that produce ROS are summarized in Fig. 3. High levels of ROS can cause protein oxidation, DNA damage and lipid peroxidation (Mach and Greenberg, 2004). Protein oxidation can disrupt the protein structure through side chain alterations and backbone cleavages, leading to denaturation, aggregation and degradation (Dean et al., 1997; Mach and Greenberg, 2004). Lipid peroxidation can cause deterioration of biomembranes and leakage of solutes (Pauls and Thomson, 1984). To prevent the hazardous effects of ROS build-up, plants scavenge and remove these molecules via several antioxidant defence systems. These defence systems involve both non-enzymatic and enzymatic reactions that are not distributed uniformly to ensure that the defence systems vary among different sub-cellular compartments (Gaspar et al., 2002). 24 Fig. 3. Reactions producing various Reactive Oxygen Species (ROS) within plant cells. (Figure modified from Gaspar et al., 2002). 25 Fig. 4. Diagram showing detoxification of Reactive Oxygen Species (ROS). Superoxide anion (O2-.) is converted into hydrogen peroxide (H2O2) by superoxide dismutase. H2O2 is either converted into H2O and O2 by catalase or enters the Halliwell-Asada pathway and is converted to H2O. (Diagram redrawn from Mach and Greenberg, 2004). 26 A variety of antioxidant enzymes are involved in the enzymatic detoxification of ROS (Fig. 4). Peroxidases and catalases act specifically to scavenge for H2O2 (Gaspar et al., 2002), but differ in that peroxidases require an additional substrate for catalysis whereas catalases do not (Bowler et al., 1992). There are two big groups of peroxidases in plant cells – (1) peroxidases of which oxidation products of the electron donors have physiological roles, and (2) peroxidises, of which function involves the scavenging of H2O2 (Asada, 1992). Guaiacol peroxidases (POD) belong to the first group and catalyse reactions that generate free radicals, leading to the cross-linking of cell wall proteins, polyphenols and cutin, in order to form walls of greater resistance towards environmental and/or pathogenic attacks (Asada, 1992; Gaspar et al., 2002). PODs are generally localized in the cell wall (Bowler et al., 1992). Ascorbate peroxidase (APX), on the other hand, belongs to the second group of peroxidases and plays a more important role in preventing oxidative stress (Asada, 1992; Mach and Greenberg, 2004). APXs are localized in the chloroplasts and together with glutathione reductase (GR), act specifically to scavenge for H2O2 and maintain a high GSH/GSSG ratio in the Halliwell-Asada pathway (Asada, 1992; Foyer and Noctor, 2005). Unlike APX, GR is not confined to the chloroplasts, but can also be found in the mitochondria and cytoplasm, and can even work together with another antioxidant enzyme, superoxide dismutase (SOD), to remove O2-. (Bowler et al., 1992) SOD is an antioxidant enzyme that is present in all subcellular compartments where oxidative stress is likely to occur and acts to reduce O2-. levels (Bowler et al., 1992). 27 2.5 Scindapsus aureus (Lindl. & André) Engl. Scindapsus aureus (Fig. 5), also known as Epipremnum aureum, is a terrestrial monocot, native to Southeast Asia and the Solomon Island (Zhang et al., 2005). It belongs to the family Araceae, and the shoots possess insoluble calcium oxalate crystals, a characteristic common to members of Araceae family (Mayo et al., 1997). Common names for this plant include golden porthos, devil’s ivy and money plant. S. aureus is a shade plant and is highly tolerant to drought (Kamel et al., 2007). However, it is easily propagated via shoot cuttings and is able to grow quickly in water (Kamel et al., 2007). The leaves of S. aureus are evergreen and heart-shaped and in certain cultivars, prolonged exposure to sunlight can result in yellow- to whitecoloured varigation. S. aureus produces trailing stems when growing along the ground, but is also able to climb trees and walls via adhesive aerial roots (Kamel et al., 2007). Because of its low maintenance and attractive foliage, S. aureus is a very popular indoor ornamental plant. Recently, there has been a growing interest in researching about the phytoremediation capabilities of S. aureus. Kamel et al. (2007) reported that S. aureus roots were not only able to tolerate high concentrations of cobalt-60 and cesium-137, but could also remove them from soil and accumulate them in the roots. S. aureus could also take up nicotine as a xenobiotic and accumulate it in the mesophyll cells (Weidner et al., 2005). Research has shown that S. aureus is not only able to remove pollutants from soil and water, but also from the atmosphere. Tani et al. (2007) observed that S. aureus leaves were able to remove methyl isobutyl ketone (a fumigation volatile organic compound) from the surrounding air. Sawada and Oyabu 28 B A Fig. 5. Scindapsus aureus plants kept as ornamental plants. Fig. 5A shows trailing stems of S. aureus along the ground. Fig. 5B shows S. aureus as an indoor ornamental plant. 29 (2008) also reported that S. aureus plants could purify indoor air by removing formaldehyde, toluene and xylene from the atmosphere. Since S. aureus have such potential in phytoremediation, it would be an ideal candidate to research on for the phytoremediation of NO3- and BPA. 30 Chapter 3. Materials and Methods 3.1 Phytoremediation of Nitrates by S. aureus 3.1.1 Plant Materials and Growth Conditions Shoot cuttings of Scindapsus aureus were grown partially submerged in water in the greenhouse of the National University of Singapore (daily PPFD ranged from 70 – 300 µmol m-2 s-1 and daily temperature ranged from 25°C to 35°C). Cuttings, weighing ca. 3 g each, with two leaves and two nodes and all roots removed were allowed to acclimatize in 100 times diluted Hoagland’s solution (100X diluted HS) (Hoagland and Arnon, 1950) for 10 days before the start of the experiment. The HS used was modified to remove all nitrogen-containing compounds: 5 mM KCl, 5 mM CaCl2, 2 mM MgSO4.7H2O, 1 mM KH2PO4, 0.5% (w/v) Fe.EDTA, 0.28% (w/v) H3BO4, 0.022% (w/v) ZnSO4.7 H2O, 0.0025% (w/v) Na2MoO4, 0.181% (w/v) MnCl2.4 H2O, 0.008% (w/v) CuSO4.5 H2O. Subsequently, various concentrations of KNO3 (0 mg/L, 20 mg/L, 50 mg/L, 100 mg/L and 200 mg/L) were added into the 100X diluted HS, constituting the ‘polluted’ waters for this study. Eight shoot cuttings were grown in plastic tanks (30 X 23 X 20 cm) containing 4L of polluted water for 8 weeks. Black polyvinyl chloride holders (15 X 9 X 5 cm) were used to anchor the cuttings. The total plant fresh weight per water setup was standardized at 20 g post-acclimatization. The ‘polluted’ water in each tank was topped-up to 4 L every week with 100X diluted HS. Water setups without any plant material were used as controls. All experiments were replicated four times. 31 3.1.2 Water Samples Water samples were collected weekly, syringe-filtered through 0.2 µm filters (Satorius, USA) and kept frozen at 0 °C until NO3- quantification. Levels of NO3- in the samples were analyzed via an ion chromatograph system (ICS-1000, Dionex, Sunnyvale, CA, USA) equipped with an isocratic pump. Weekly measurements of the conductivity and pH of the various polluted water were also made. Water conductivity was measured with a conductivity meter (Cond 315i/SET, WTW, Germany) at 29 °C – 31 °C. pH readings were recorded at 25 °C using a pH meter (pH meter 145, Corning, USA). 3.1.3 Plant Samples Plant samples were harvested every two weeks for the analyses of various physiological parameters. All harvested plant materials were washed with distilled water and dried with paper towels before analysis. 3.1.4 Determination of Algal Growth Algal growth was determined based on chlorophyll a and pheophytin a concentrations according to Yentsch and Menzel (1963) with some modifications. Water samples were collected by the end of 8 weeks to determine the amount of algal growth. Samples were first filtered through glass fibre filter paper (GF/C, Whatman, USA), 47 mm in diameter, and then extracted with 80% (v/v) methanol. The suspension was collected in a 50 ml centrifuge tube and placed in a water bath (65 – 80°C) for 5 min, after which the tubes were immediately cooled in an ice bath. The suspension was then centrifuged at 1,500 g and 20°C for 5 min and the supernatant removed. Three ml of each supernatant were transferred to a cuvette and absorbances 32 were recorded at 665 nm and 750 nm. Twelve µl of 0.5 M HCl were subsequently added to the extract in the cuvette and mixed. Ninety seconds later, absorbances of the acidified extract were taken at 665 nm and 750. Chlorophyll a and pheophytin a concentrations were calculated via the following formulae (Yentsch and Menzel, 1963): [Chlorophyll a] (µg/ L) [Pheophytin a] (µg/ L) [26.7 X (OD 665 – OD 665 acidified ) X Vol extract] = Vol filtered 26.7 X [(1.7 X OD 665 ) – OD665 acidified ] X Vol extract] = Vol filtered 3.1.5 Plant Growth Parameters The harvested cuttings were divided into shoots and roots, after which the fresh weight (FW) of each individual part was determined. Each part was then wrapped in aluminum foil and placed in an 80 °C oven for two weeks before determining the dry weight (DW). Readings for the total leaf area were obtained by tracing the leaf outlines on black paper and feeding the cut-outs through an area-meter (LI-3100 Area-meter, LiCor BioSciences, USA). Total leaf area per leaf was determined by dividing the total leaf area by the number of leaves produced per plant. 3.1.6 Determination of Photosynthetic Pigment Concentrations Photosynthetic pigment concentrations (chlorophyll a, chlorophyll b, carotenoids) were quantified according to Arnon (1949) and Embry and Nothnagel (1988). Photosynthetic pigments were extracted by grinding 0.03 g of fresh leaf 33 samples in 5 ml of absolute acetone. Concentrations of the various photosynthetic pigments were calculated according to Arnon (1949) and Embry and Nothnagel (1988) and expressed as mg photosynthetic pigment per g FW and per g DW. [Chlorophyll a] (mg/L) = [OD 663 X 12.7] – [OD 645 X 2.69] [Chlorophyll b] (mg/L) = [OD 645 X 22.9] – [OD 663 X 4.68] [Total chlorophylls] (mg/L) = [OD 645 X 20.2] + [OD 663 X 8.02] [Carotenoids] (mg/L) = [OD 460 X 5] – [OD 645 X 14.87] + [OD 663 X 2.84] 3.1.7 Chlorophyll Fluorescence Chlorophyll fluorescence was determined using a versatile pulse amplitude fluorometer (FMS II, Hanstech Instruments Ltd, Norfolk, United Kingdom). Chlorophyll fluorescence emission from the upper surface of attached leaves was determined. Each leaf sample was dark-adapted for 30 min using a light-tight clip (Hanstech Instruments Ltd, Norfolk, United Kingdom) before analysis. Minimal fluorescence level, Fo, was determined at a light level lower than 0.01 µmol m-2 s-1. The maximal fluorescence yield of the dark-adapted leaf, Fm, was determined by applying a saturating light pulse of 10, 000 µmol m-2 s-1. The maximal fluorescence yield of the light-adapted leaf, Fm’, was then determined by the application of a saturating light pulse of 10, 000 µmol m-2 s-1 for 1 s duration, at intervals of 60 s for 30 min. At the end of 30 min, the saturating light pulse was turned off and a far-red light of 735 nm, 600 µmol m-2 s-1, was switched on. The value of the minimal fluorescence yield of the light-adapted leaf, Fo’, was subsequently determined. The values of Fv / Fm, Fv’ / Fm’, qP, qNP, NPQ and ФPSII were calculated using the Modfluor software V. 2.00 (Hanstech Instruments Ltd, Norfolk, United Kingdom). 34 3.1.8 Total Nitrogen Analysis in Plant Tissues The level of total nitrogen (TN) in the oven-dried plant tissues was determined by the Kjeldahl digestion method with some modifications. Four ml of concentrated H2SO4 and 1 piece of Kjeltab (1.5 g K2SO4, 1.5 g Se) were added to each plant sample in a glass digestion tube. These samples were then digested in a Kjeldahl Digestion Stand (Digestion System 12, 1009 Digester, Tecator) for 2 h at 350 °C. Digested samples were left overnight in a fume hood to cool. The total nitrogen level in each of the digested samples was determined using the Tecator Kjeltec Auto 1030 Analyzer (Sweden). Total nitrogen level of the plant samples was expressed according to Emmert (1935): Total Nitrogen Level (mg N/ g DW) = (reading) X (Normality sulfuric acid X Mol Wt Nitrogen ) DW of sample (g) Normality sulfuric acid = 1/35 Mol Wt Nitrogen = 14.007 g/mol 3.1.9 Determination of Total Soluble Sugar (TSS) Concentration The concentration of TSS was determined using the phenol-sulphuric acid assay (Dubois et al., 1956) with some modifications. Fresh leaf samples (0.1 g) were suspended in 10 ml of distilled water and boiled in a water bath for 20 min. One ml of the extract was added to 1 ml of 5% (v/v) phenol, followed by 5 ml of concentrated sulphuric acid. The reaction mixture was thoroughly mixed and left to stand for 20 min at room temperature (25 °C) for color development. The absorbance of the solution was then read at 490 nm. Glucose was used as the calibration standard. The 35 concentration of TSS was expressed as mg glucose equivalent per g FW and per g DW. 3.1.10 Determination of Total Soluble Protein (TSP) Concentration Leaf materials (0.1 g FW) were ground in 1 ml of 0.1 M phosphate buffer (pH 7.5) at 4°C to extract TSP. The extract was then centrifuged at 13, 000 g and 4 °C for 10 min. The supernatant was removed to determine TSP concentrations. The concentration of TSP was determined using the Bradford method (Bradford, 1976) with the Bio-Rad Protein Assay Kit (Bio-Rad, Hercules, CA, USA) according to manufacturer’s instructions. Bovine serum albumin (Sigma) was used as the standard. The concentration of TSP was expressed as mg proteins per g FW and per g DW. 3.1.11 Determination of Nitrate Reductase (NR) Activity (EC 1.6.6.1) The activity of NR was determined according to Hageman and Hucklesby (1971), Ahmad and Abdin (1999) and Taghavi and Babalar (2007) with some modifications. Leaf and root materials (0.1 g FW each) were ground in 1 ml of extraction buffer containing 50 mM Tris buffer (pH 8.8), 1 mM EDTA, 10 mM cysteine, 0.3% (w/v) PVP (MW = 360,000) and 2 mM mercaptoethanol at 4°C. The mixture was then centrifuged at 13,000 g and 4°C for 15 min. The reaction mixture containing 50 mM phosphate buffer (pH 7.5), 5 mM KNO3, 1 mM NADH and 490 µl of enzyme extract, topped up to 2 ml with distilled water, was incubated for 15 min at 30 °C. At the end of the incubation period, the reaction was stopped by the addition of 1 ml 1% (w/v) sulfanilamide dissolved in 1.5 M HCl and 1 ml 0.02 % NED. The entire mixture was left to stand for 30 min to 36 allow for colour development. Enzymatic activity was determined by recording the absorbance at 540 nm and NR activity was expressed in µM NO2-/ g FW/ h and µM NO2-/ mg protein/ h. 3.1.12 Determination of Lipid Peroxidation Levels Lipid peroxidation levels were determined by recording the amount of malondialdehyde (MDA) formed upon reaction with thiobarbituric acid (TBA). The TBA reactivity (TBARs) was determined according to Heath and Packer (1968), and Khan et al. (2009) with some modifications. Fresh plant materials (leaves, each 0.3 g FW) were ground in 3 ml of 0.1% (w/v) trichloroacetic acid (TCA) and centrifuged at 2,500 g for 5 min. One ml of the supernatant was added to 4 ml of reaction mixture [0.5% (w/v) TBA; 20% (w/v) TCA] and heated at 95ºC for 30 min. The mixture was then left to cool for 5 min and centrifuged at 2,500 g for 5 min. The absorbance of the supernatant was determined at 532 nm and corrected for unspecific turbidity after subtraction from the absorbance obtained at 600 nm. TBARs was expressed as µM MDA formed per g FW and per g DW using the extinction coefficient of 155 mM-1 cm-1(Heath and Packer, 1968). 3.1.13 Determination of Ascorbate Peroxidase (APX) Activity (EC 1.11.1.11) The activity of APX was determined according to Nakano and Asada (1981) with some modifications. Leaf materials (0.1 g FW) were ground in 1 ml of extraction buffer containing 50 mM phosphate buffer (pH 7.8), 0.1mM EDTA, 5 mM ascorbic acid, 0.5% (w/v) PVP (MW = 360,000), 0.1% (v/v) Triton X-100 and 0.05% (v/v) mercaptoethanol at 4°C. The mixture was then centrifuged at 13,000 g and 4°C for 15 min. 37 The reaction mixture contained 50 mM phosphate buffer (pH 7.0), 0.1 mM EDTA, 0.5 mM ascorbic acid, 0.3 mM H2O2 and 50 µl of enzyme extract, topped up to 3ml with distilled water. The enzymatic activity was determined by recording the decrease in OD at 290 nm for 1 min. The activity was expressed as unit of APX per mg protein and per g FW using an extinction coefficient of 2.8 mM-1 cm-1 (Nakano and Asada, 1981). 3.1.14 Determination of Glutathione Reductase (GR) Activity (EC 1.6.4.2) The activity of GR was determined according to Gupta et al. (1993) with some modifications. Leaf materials (0.1 g FW) were ground in 1 ml of extraction buffer containing 200 mM phosphate buffer (pH 7.0), 0.1 mM EDTA and 1% (w/v) PVP (MW = 360,000) at 4°C. The mixture was then centrifuged at 13,000 g and 4°C for 15 min. The reaction mixture contained 100 mM phosphate buffer (pH 7.8), 2 mM EDTA, 0.1 mM NADPH, 0.5 mM GSSG and 80 µl of enzyme extract, topped up to 3ml with distilled water. The enzymatic activity was determined by recording the decrease in OD at 340 nm for 1 min. The activity was expressed as unit of GR per mg protein and per g FW using an extinction coefficient of 6.2 mM-1 cm-1 (Gupta et al., 1993). 3.1.15 Determination of Superoxide Dismutase (SOD) Activity (EC 1.15.1.1) The activity of SOD was determined according to Beauchamp and Fridovich (1971), and Giannopolitis and Ries (1977) with some modifications. Leaf materials (0.1 g FW) were ground in 1 ml of extraction buffer containing 50 mM phosphate 38 buffer (pH 7.8) and 0.1 mM EDTA at 4°C. The mixture was then centrifuged at 13,000 g and 4°C for 15 min. The reaction mixture contained 50 mM phosphate buffer (pH 7.8), 0.1 mM EDTA, 0.02 mM NaCN, 0.0117 mM riboflavin, 13 mM methionine, 5.6 X 10-2 mM NBT and 20 µl of enzyme extract, topped up to 3ml with distilled water. The reaction was initiated by illuminating the mixture with 120 µmol quanta m-2s-1 at 28°C for 15 min, after which the absorbance at 560 nm was determined. One unit of SOD was defined as the amount that caused a 50% inhibition on the photoreduction of NBT. The activity was expressed as unit of SOD per mg protein and per g FW. 3.1.16 Determination of Guaicol Peroxidase (POD) Activity (EC 1.11.1.7) The activity of POD was determined according to Civello et al. (1995) with some modifications. Leaf materials (0.1 g FW) were ground in 1 ml of extraction buffer containing 50 mM phosphate buffer (pH 7.0), 0.1 mM EDTA, 1% (w/v) PVP (MW = 360,000), 0.1% (v/v) Triton X-100 and 1M NaCl at 4°C. The mixture was then centrifuged at 13,000 g and 4°C for 15 min. The reaction mixture contained 50 mM phosphate buffer (pH 6.0), 20 mM guaiacol, 4 mM H2O2 and 50 µl of enzyme extract, topped up to 3ml with distilled water. The enzymatic activity was determined by recording the increase in OD at 470 nm for 1 min. The activity was expressed as unit of POD per mg protein and per g FW using an extinction coefficient of 26.6 mM-1 cm-1 (Civello et al. 1995). 39 3.2 Phytoremediation of BPA by S. aureus 3.2.1 Culture of Plant Materials One cm-long terminal shoot tips were cut from whole S. aureus plants and aseptically cultured in 90 mm (diameter) Petri dishes containing 20 ml of halfstrength Murashige-Skoog medium (including vitamins) supplemented with 2% (w/v) sucrose and 0.3% (w/v) Gelrite. All media were adjusted to pH 5.5 before autoclaving at 121°C for 20 min. Five shoot cuttings were cultivated in each Petri dish. After two weeks, the shoot cuttings were transferred to GA-7 containers (Magenta Corp., Chicago, USA) containing 50 ml of the same shoot cultivation medium. Four shoot cuttings were put into each of these GA-7 containers. All cultures were kept at 25 ± 3°C under a 16 h photoperiod with 40 µmol m-2 s-2 provided by daylight fluorescent lamps. These cultures were allowed to grow for 4 months before they were treated with various BPA levels for different durations. 3.2.2 BPA Treatment Set-up and Conditions Four months old plants were thoroughly washed with sterile distilled water and transferred under aseptic conditions into GA-7 containers containing liquid growth media – distilled water spiked with various concentrations of BPA (0, 50, 100, 250 µM). These plants were approximately the same size on Day 0 (D0), at the start of the experiments. Intact plants (including roots), each with three expanded leaves, were used throughout the entire experiment. The liquid growth media were prepared by diluting a 100 mM stock solution of BPA (dissolved in ethanol) with autoclaved distilled water. All plants were held in place using a stainless steel grate, with roots fully submerged in the liquid growth medium. The lower half of each GA-7 container 40 was wrapped in black paper to reduce the effect(s) of light on BPA degradation. Similar set-ups of GA-7 containers with BPA-spiked water, but without plant materials, were used as the experimental controls. All GA-7 containers (with or without plants) were placed under a 16 h photoperiod with 40 µmol m-2 s-2 provided by daylight fluorescent lamps for 7 days at 25 ± 3°C. 3.2.3 Water Samples Water samples were collected at 0, 3, 6, 9, 24, 48, 72, 96, 168 h after exposure of the plants to BPA. BPA levels were quantified according to Imai et al. (2007) with some modifications. One hundred µl of the water sample were passed through a HPLC system, using a reverse phase column (XBridgeTM, C-18 column, 5µm, 150 X 4.6 mm; Waters, Massachusetts, USA), with a flow rate of 0.5 ml min-1. Elution was carried out with mobile phase B (100% methanol) and mobile phase A (15 mM formic acid, pH 4) at a ratio of 95/5 (v/v). BPA was detected at a wavelength of 280 nm. 3.2.4 Plant Samples Plants were harvested at 0, 1, 4 and 7 days (D0, D1, D4 and D7 respectively) after exposure to BPA for the analyses of various physiological parameters. All harvested plants were washed with distilled water and dried with paper towels before analysis. 3.2.5 Plant Growth Parameters The harvested plants were divided into leaves and roots, after which the fresh weight (FW) and dry weight (DW) of each individual part were determined. 41 Determination of DW was done using the same method as mentioned for phytoremediation of nitrates (Section 3.1.5). 3.2.6 Determination of Photosynthetic Pigment Concentrations Photosynthetic pigment concentrations were determined using the same method as mentioned for phytoremediation of nitrates (Section 3.1.6). 3.2.7 Chlorophyll Fluorescence Chlorophyll fluorescence was determined using the same method as mentioned for phytoremediation of nitrates (Section 3.1.7). 3.2.8 Determination of Total Soluble Sugar (TSS) Concentration The concentration of TSS was determined using the same method as mentioned for phytoremediation of nitrates (Section 3.1.9). 3.2.9 Determination of Total Soluble Protein (TSP) Concentration The concentration of TSP was determined using the same method as mentioned for phytoremediation of nitrates (Section 3.1.10). 3.2.10 Determination of Lipid Peroxidation Levels Lipid peroxidation levels (TBARs) were determined using the same method as mentioned for phytoremediation of nitrates (Section 3.1.12) except both leaf and root materials were used. 42 3.2.11 Determination of Ascorbate Peroxidase (APX) Activity (EC 1.11.1.11) The activity of APX was determined using the same method as mentioned for phytoremediation of nitrates (Section 3.1.13). 3.2.12 Determination of Glutathione Reductase (GR) Activity (EC 1.6.4.2) The activity of GR was determined using the same method as mentioned for phytoremediation of nitrates (Section 3.1.14). 3.2.13 Determination of Superoxide Dismutase (SOD) Activity (EC 1.15.1.1) The activity of SOD was determined using the same method as mentioned for phytoremediation of nitrates (Section 3.1.15). 3.2.14 Determination of Guaiacol Peroxidase (POD) Activity (EC 1.11.1.7) The activity of POD was determined using the same method as mentioned for phytoremediation of nitrates (Section 3.1.16). 3.3 Statistical Analysis All results are presented as mean ± standard error and analysed for statistical significance by multifactor ANOVA. Where ANOVA detected differences, Fisher’s least significant difference (LSD) test was used at a 5% level of significance. Data with different alphabets indicated that the mean values were significantly different from one another. Correlation coefficients determined for scatterplots were calculated using Pearson’s product-moment correlation coefficient. 43 Chapter 4. Results 4.1 Preliminary Screening of Plants A preliminary screen was carried out using five different plant species to identify one species capable of tolerating partially submergence conditions and high NO3- concentrations, and at the same time, able to phytoremediate NO3- from water. The five species included Echinodorus palaefolius (Nées et Mart.) Macbr., Ipomoea aquatic Frosk?, Cryptocoryne ciliate (Roxburgh) Schott, Hydrocotyl sibthorpioides Lam. and S. aureus. 4.1.1 Calibration and Optimization of Aquarium Nitrate Test Kit Since the preliminary screen was to obtain a rough gauge of the ability of the plants to phytoremediate NO3-, a fast method was required to quantify NO3concentrations in water. A commercially available aquarium NO3- test kit (Aquarium Pharmaceuticals, Inc., PA, USA) was utilized for this purpose. This kit is normally used for the determination of NO3- levels in aquariums to ensure that the NO3- levels in the water do not exceed the critical toxic limits for aquarium organisms. NO3- concentrations were determined according to manufacturer’s instructions but with some modifications. According to manufacturer’s specifications, after mixing the indicator solutions with water samples containing NO3-, a visual comparison with a colour chart was required for determining NO3- levels. The colour of the indicator solution changed from yellow to orange to red with increasing NO3concentrations (Fig. 6). In order to increase the accuracy of the test, the kit was calibrated to a range of pre-determined NO3- concentrations (0 – 30 mg/L) and 44 B Fig. 6. Commercial NO3- aquarium kit calibration based on known NO3concentrations (0 – 30 mg/L). Colour changed from yellow to orange to red with increasing NO3- concentrations. The first cuvette shown (B) is the blank containing 0 mg/L NO3-. 45 absorbances were recorded using a UV-visible spectrometer (UV mini-1240, Shimadzu, Japan). Photometric spectrums were obtained for the various NO3- concentrations using wavelengths of 300 to 700 nm (includes the entire visible spectrum). Distinct peaks were only observed between wavelengths of 500 to 600 nm. To determine the ideal wavelength to use for this kit, OD readings obtained at wavelengths between 500 to 600 nm were used, and plotted against the various NO3- concentrations (Fig. 7A). Absorbance readings obtained were high at 540 nm and they were linearly related to the different NO3- concentrations. A coefficient of determination (R2) calculated for the best fit line was 0.993 (close to 1) (Fig. 7B), signifying a strong linear relationship between the two parameters. A concentration of 0 mg/L NO3(including indicator solution) was used as the blank for all assays. 4.1.2 Screening of Five Different Plant Species for Phytoremediation of Nitrate For the screening process, the plants were grown partially submerged in water. The set-up was standardized at 4.5 g of plant material for every 1 L of growth medium (NO3- spiked water). The plants were allowed to acclimatize in 100X diluted Hoagland’s solution (100X diluted HS), modified to remove all nitrogen containing compounds, for 10 days before the start of the experiment. Subsequently, various concentrations of KNO3 (0, 20, 50, 100 and 200 mg/L) were added into the 100X diluted HS, constituting the ‘polluted’ waters for this study. During the acclimatization phase, E. palaefolius and I. aquatica plants did not adapt well to the water set-up and the plants were extremely unhealthy by the end of the 10 days of acclimatization. E. palaefolius leaves turned brown and the plants 46 (A) (B) Fig. 7. Calibration graphs for aquarium NO3- test kit using NO3- concentrations ranging from 0 to 30 mg/L. Fig. 7A shows the relationship between NO3concentrations and absorbance between 500 to 600 nm. Fig. 7B shows the linear regression chart with absorbance readings determined at 540 nm. The coefficient of determination (R2) for the linear regression between both parameters was 0.993. Each value is represented as the mean ± SE (n = 4). 47 Fig. 8. Echinodorus palaefolium plants (A,B) before acclimatization and (C,D) after acclimatization (W0). Bar = 5 cm. All photographs are representative of 4 replicates. 48 Fig. 9. Ipomoea aquatica plants after acclimatization (W0). Fig. 9A shows plants in tanks. Bar = 20 cm. Fig. 9B and C show close-ups of chlorotic leaves. Bar = 2 cm. All photographs are representative of 4 replicates. 49 wilted (Fig. 8), whereas I. aquatica leaves underwent severe chlorosis and became bleached (Fig. 9). The experiment was, hence, discontinued for these two plant species. The remaining three plant species were grown in NO3--spiked water for four weeks, after which the NO3- concentrations remaining in the growth media were determined via the aquarium NO3- test kit (Table 1). The total fresh weights (FW) of the individual plants were also determined post-acclimatization (Week 0; W0) and at the end of the screening process (Week 4, W4). Percentage NO3- removal (Table 2) indicated that only H. sibthorpioides and S. aureus were able to remove NO3- from water. C. ciliata and H. sibthorpioides showed decreases in total FW on W4 compared to W0, particularly when they were exposed to high NO3- concentrations (200 mg/L) (Table 3). The shoots of both plant species also turned yellow and/or necrotic on W4 compared to W0 (Fig. 10, 11). Although H. sibthorpioides appeared to be a more efficient phytoremediator of NO3-, the plants died by the end of the experiment. S. aureus plants, however, showed more than 2-fold increase in total FW on W4, compared to W0, in all plants (Table 3). Photographs of the plants also showed extensive root growth and increased leaf number over the four weeks of screening (Fig. 12). 4.2 Phytoremediation of Nitrate by S. aureus 4.2.1 Removal of Nitrate by S. aureus Water Samples Observations from the screening process indicated that S. aureus was the best plant to use for further studies, as it has the highest percentage of NO3- removal 50 Fig. 10. Cryptocoryne ciliata plants (A) before acclimatization, (B) after acclimatization (W0) and after 4 weeks of exposure to (C) 0 mg/L, (D) 20 mg/L, (E) 50 mg/L, (F) 100 mg/L and (G) 200 mg/L NO3-. Holes observed in the leaves are a result of necrosis observed during NO3exposure. Bar = 2 cm. All photographs are representative of 4 replicates. 51 Fig. 11. Hydrocotyl sibthorpioides plants (A, B, C) after acclimatization (W0) and after 4 weeks of exposure to (D) 0 mg/L, (E) 20 mg/L, (F) 50 mg/L, (G) 100 mg/L and (H) 200 mg/L NO3-. Bar = 2 cm. All photographs are representative of 4 replicates. 52 Fig. 12. Scindapsus aureus plants (A) before acclimatization, (B) after acclimatization (W0) and after 4 weeks of exposure to (C) 0 mg/L, (D) 20 mg/L, (E) 50 mg/L, (F) 100 mg/L and (G) 200 mg/L NO3-. Bar = 2 cm. All photographs are representative of 4 replicates. 53 Table 1. Changes in NO3- concentration in the growth medium (NO3--spiked water) over a period of 4 weeks during preliminary screening of five different plant species (Echinodorus palaefolius, Ipomoea aquatica, Cryptocoryne ciliata, Hydrocotyl sibthorpioides and Scindapsus aureus). This preliminary screen was terminated by week 4 (W4) as several of the plant species died by W4 (Week 4) under experimental conditions. Values were absent for E. paleofolius and I. aquatica due to plant death at W0 (Week 0, start of the experiment). [NO3-] in Growth Medium (mg/L) [NO3-] E. palaefolius (mg/L) W0 W4 W0 W4 W0 W4 W0 W4 W0 W4 0 - - - - 6.34 0.91 3.76 0.00 5.24 1.54 20 - - - - 32.63 15.41 27.43 0.22 23.48 1.54 50 - - - - 59.82 146.53 60.84 13.27 55.56 1.54 100 - - - - 130.82 154.68 115.04 58.19 71.28 59.02 200 - - - - 279.76 347.13 196.02 158.85 184.49 132.14 I. aquatica C. ciliata H. sibthorpioides S. aureus Note: known [NO3-] concentrations were added to the growth media, the aquarium nitrate kit was not 100% accurate, hence resulting in the different levels of [NO3-] at W0. 54 Table 2. Percentage removal of NO3- from the growth medium (NO3--spiked water) by five different plant species (Echinodorus palaefolius, Ipomoea aquatica, Cryptocoryne ciliata, Hydrocotyl sibthorpioides and Scindapsus aureus) over a period of 4 weeks during preliminary screening. (+) indicates a percentage increase (instead of removal) in NO3- concentrations in the growth medium as compared to the initial concentrations in W0. This preliminary screen was terminated by week 4 (W4) as several of the plant species died by W4 (Week 4) under experimental conditions. Values were absent for E. paleofolius and I. aquatica due to plant death at W0 (Week 0, start of the experiment). - - NO3 Percentage Removal on W4 (%) [NO3 ] (mg/L) E. palaefolius I. aquatica C. ciliata H. sibthorpioides S. aureus 0 - - 86 100 71 20 - - 53 99 93 50 - - (+)145 78 97 100 - - (+) 18 50 17 200 - - (+) 24 19 28 55 Table 3. Percentage increase in fresh weight (FW) of five different plant species (Echinodorus palaefolius, Ipomoea aquatica, Cryptocoryne ciliata, Hydrocotyl sibthorpioides and Scindapsus aureus) over a period of 4 weeks during preliminary screening. Values were absent for E. paleofolius and I. aquatica due to plant death at W0. Negative values indicate a percentage decrease in FW as compared to the initial FW in W0. Each value is represented as the mean ± SE (n = 8). [NO3 -] Percentage Increase in FW from W0 to W4 (%) (mg/L) E. paleofolius I. aquatica 0 - - C. ciliata 8.3 ± 1.6 H. sibthorpioides S. aureus 12.1 ± 3.4 144.2 ± 11.9 (1.4-fold increase) 20 - - 11.0 ± 0.5 6.5 ± 2.5 201.7 ± 68.1 (2-fold increase) 50 - - 2.1 ± 1.6 -10.8 ± 6.3 195.1 ± 15.6 (2-fold increase) 100 - - -7.7 ± 2.6 13.0 ± 6.4 209.7 ± 15.5 (2-fold increase) 200 - - -7.5 ± 2.2 -7.0 ± 6.4 162.1 ± 13.4 (1.5-fold increase) 56 among the five different plant species. It also showed increased growth, despite the exposure to high NO3- concentrations. A more precise water set-up was then used for in-depth study of the NO3- phytoremediation potential of S. aureus (Section 3.1.1). S. aureus plants were grown in 100 X diluted HS spiked with different NO3concentrations. From Fig. 13 and Table 4, it could be seen that S. aureus was able to remove 96% of the NO3- from water containing 20 mg/L NO3- on W4, 89% of the NO3- from water containing 50 mg/L NO3- on W5 and 91% of the NO3- from water containing 100 mg/L NO3- on W6. S. aureus was also able to remove about 80% of the NO3- from water containing 200 mg/L NO3- by W8. In contrast, control set-ups (set-ups with no plants; algae growing in water) showed only 83% NO3- removal from water containing 20 mg/L NO3- on W4, 77% NO3- removal from water containing 50 mg/L NO3- on W5 and 79% NO3- removal from water containing 100 mg/L NO3- on W6. In water containing 200 mg/L NO3-, algae exhibited 91% NO3- removal by the end of W8. Algae Growth Algal growth increased with increasing NO3- content in the water, as observed in the control setups (Fig. 14 A – E). Algal growth determined by mass would be highly inaccurate. Hence, it was quantified using chlorophyll a concentration, and to account for any chlorophyll degradation, pheophytin a concentration of the same samples was also determined. Based on the data obtained on W8, concentrations of chlorophyll a (Fig. 15A) and pheophytin a (Fig. 15B) increased with increasing NO3concentrations in water set-ups with or without S. aureus. Also, in water set-ups with S. aureus, chlorophyll a and pheophytin a concentrations were much lower, compared 57 (A) (B) Fig. 13. Removal of NO3- from the growth medium (NO3--spiked water) by S. aureus over a period of eight weeks. Roots of the plants were submerged in water containing 0, 20, 50, 100 and 200 mg/L of NO3- (A). Experimental controls with the same NO3concentrations were also set up but without plants growing in them (B). Each value is represented as the mean ± SE (n = 4). 58 Table 4. Percentage removal of NO3- from the growth medium (NO3--spiked water) by S. aureus and algae (experimental controls) during eight weeks of exposure. (+) indicates a percentage increase (instead of removal) in NO3- concentrations in the growth medium as compared to the initial concentrations in W0. Each value is represented as the mean ± SE (n = 4). Nitrate Percentage Removal (%) Concentration (mg/L) 0 Control S. aureus 20 50 100 200 Week 1 Week 2 Week 3 Week 4 Week 5 Week 6 Week 7 Week 8 (+)13.3 ± 5.1 60.0 ± 17.3 90.6 ± 5.4 81.3 ± 7.0 85.3 ± 4.0 84.3 ± 0.3 88.4 ± 5.8 88.8 ± 4.0 57.7 ± 3.1 89.3 ± 2.3 83.4 ± 1.6 84.0 ± 1.5 77.8 ± 2.8 89.5 ± 5.26 85.0 ± 1.9 0.1 ± 10.8 Control (+)13.7 ± 7.6 8.9 ± 6.6 53.8 ± 17.0 81.8 ± 14.9 97.5 ± 0.1 96.9 ± 1.6 97.5 ± 0.9 97.9 ± 1.5 S. aureus (+) 5.5 ± 7.0 24.1 ± 1.3 70.6 ± 14.6 96.4 ± 0.6 97.6 ± 0.7 96.7 ± 0.5 97.9 ± 0.3 97.7 ± 0.5 Control (+) 5.2 ± 2.3 8.4 ± 4.3 25.8 ± 7.8 61.3 ± 37.4 76.1 ± 23.0 97.3 ± 1.7 99.1 ± 0.3 99.0 ± 0.3 S. aureus (+) 2.7 ± 6.4 11.6 ± 1.8 47.4 ± 7.0 76.0 ± 12.8 89.2 ± 6.8 98.7 ± 0.5 99.3 ± 0.3 99.3 ± 0.3 Control 4.2 ± 4.0 7.6 ± 0.5 18.6 ± 1.1 34.8 ± 5.0 58.7 ± 11.8 77.9 ± 12.5 93.6 ± 6.4 99.7 ± 0.1 S. aureus 8.0 ± 2.2 15.1 ± 9.0 34.1 ± 7.4 58.1 ± 11.9 80.8 ± 10.6 90.0 ± 6.6 97.0 ± 1.7 99.3 ± 0.15 Control (+) 2.6 ± 3.5 0.5 ± 0.4 6.4 ± 2.7 22.7 ± 8.2 46.5 ± 10.0 66.4 ± 7.1 78.9 ± 2.5 90.9 ± 2.3 S. aureus (+) 4.3 ± 6.0 3.2 ± 4.6 13.4 ± 5.9 32.3 ± 10.6 46.8 ± 12.4 62.5 ± 10.7 71.2 ± 12.2 78.4 ± 9.2 59 Fig. 14. Polluted water set-ups on W8 with controls (A) 0 mg/L, (B) 20 mg/L, (C) 50 mg/L, (D) 100 mg/L and (E) 200 mg/L NO3-, as well as S. aureus in water containing (F) 0 mg/L, (G) 20 mg/L, (H) 50 mg/L, (I) 100 mg/L and (J) 200 mg/L NO3-. Bar = 30 cm. 60 (A) (B) Fig. 15. Algal growth in polluted water set-ups with and without S. aureus on W8. Increased algal growth was correlated with increased concentrations of (A) chlorophyll a and (B) pheophytin a. Each value is represented as the mean ± SE (n = 4). 61 to control water set-ups without any S. aureus plants. For water containing 200 mg/L NO3-, the presence of S. aureus plants resulted in the reduction of chlorophyll a and pheophytin a concentrations to levels similar to that of control set-ups (without plants) with water containing 50 and 100 mg/L NO3-. Conductivity of Water Samples Conductivity of water generally increased with NO3- levels (W0, Fig. 16). In water set-ups containing S. aureus, conductivity increased from W0 to W6 for water containing 0 and 20 mg/L NO3-, but decreased for water containing 50 and 100 mg/L NO3-, during the same period (Fig. 16A). Conductivity of water containing 200 mg/L NO3- showed a continual decrease from W0 to W8. Experimental controls with algal growth displayed increases in conductivity from W0 to W6 for water containing 0 to 100 mg/L NO3- (Fig. 16B). In these controls, conductivity of water containing 200 mg/L NO3- slightly decreased from W0 to W8. pH of Water Samples pH of water (all concentrations of NO3-) containing S. aureus gradually increased from W0 until W5; it then started to decrease until W8 (Fig. 17A). On W5, water containing 200 mg/L NO3- showed the highest pH (8.31 ± 0.98) whereas water containing 0 mg/L NO3- had the lowest pH (6.64 ± 0.67). By W8, water containing 200 mg/L NO3- was still showing the highest pH (5.66 ± 0.80), but the lowest pH (4.16 ± 1.10) was detected in water containing100 mg/L NO3-. Experimental control set-ups with algal growth also showed similar trends for all concentrations of NO3- (Fig. 17B) with pH increasing from W0 to W5, followed by a decrease from W5 to W8. Water containing 200 mg/L NO3- showed the highest pH 62 (B) Fig.16. Conductivity of polluted water set-ups over a period of eight weeks with (A) and without (B) S. aureus. Each value is represented as the mean ± SE (n = 4). 63 (A) (B) Fig. 17. pH of polluted water set-ups over a period of eight weeks with (A) and without (B) S. aureus. Each value is represented as the mean ± SE (n = 4). 64 among all the NO3- concentrations throughout the 8 weeks. The lowest pH levels throughout the 8 weeks were detected in water that initially had no NO3- added to it. 4.2.2 Growth Parameters Fresh Weight and Dry Weight In general, FW and DW increased over the 8 weeks for all plants (Fig. 18). The largest increase in leaf FW from W0 to W8 was observed in plants exposed to 100 mg/L NO3- whereas the largest increase from W0 to W8 in root FW was observed in plants exposed to 200 mg/L NO3-. For DW, however, the largest increase in leaf DW and root DW were observed in plants exposed to 200 mg/L NO3-. The increase in total FW (Fig. 19A) from W0 to W8 was highest in plants exposed to 100 mg/L NO3-. However, the largest increase in total DW (Fig. 19B) from W0 to W8 was observed in plants treated with 200 mg/L NO3- . FW: DW ratio (Table 5) decreased in plants exposed to 0, 20 and 50 mg/L NO3-, but increased in plants exposed to 100 and 200 mg/L NO3-, by the end of the study. The largest decrease in FW: DW ratio (from W0 to W8) was observed in plants exposed to 50 mg/L NO3- whereas the largest increase (from W0 to W8) was observed in plants exposed to 100 mg/L NO3-. Not all plants developed roots by the end of the acclimatization period, which resulted in the root: shoot ratio on W0 to be 0. Root: shoot ratio (Table 6) increased in all plants from W0 to W8. The largest increase was observed in plants exposed to 0 mg/L NO3-, whereas the smallest increase was observed in plants exposed to 200 mg/L NO3-. 65 Fig. 18. Effects of various NO3- concentrations on fresh weight (FW) and dry weight (DW) of S. aureus over a period of eight weeks. Fig 6A, B, C, D, E represent FW, whereas Fig 6F, G, H, I, J represent DW of leaves and roots from plants exposed to 0, 20, 50, 100 and 200 mg/L NO3- respectively, during eight weeks of exposure. Each value is represented as the mean ± SE (n = 4). 66 (B) Fig .19. Effects of various NO3- concentrations on (A) total FW and (B) total DW of S. aureus during eight weeks of exposure. Each value is represented as the mean ± SE (n = 4). 67 Table 5. Effects of various NO3- concentrations on FW: DW ratio of S. aureus during eight weeks of exposure. Each value is represented as the mean ± SE (n = 4). Nitrate Concentration (mg/L) FW: DW Ratio Week 0 Week 2 Week 4 Week 6 Week 8 0 9.05 ± 0.12 cdefg 9.65 ± 0.45 efgh 7.56 ± 0.33 ab 7.48 ± 0.31 20 9.34 ± 0.73 defgh 8.72 ± 0.57 abcdefg 8.14 ± 0.49 abcd 8.95 ± 0.29 cdefg 50 9.21 ± 0.39 cdefg 8.58 ± 0.14 abcdef 8.84 ± 0.43 bcdefg 10.04 ± 0.68 ghi 7.99 ± 0.55 abc 100 8.32 ± 0.38 abcde 8.01 ± 0.31 abcd 9.57 ± 0.13 efgh 10.68 ± 0.33 hi 11.32 ± 0.36 200 9.80 ± 0.55 fgh 8.17 ± 0.24 abcd 8.45 ± 0.79 abcde a 8.66 ± 0.34 abcdef 8.15 ± 0.81 abcd 8.18 ± 0.20 abcd 11.37 ± 0.82 i i 68 Table 6. Effects of various NO3- concentrations on root: shoot ratio of S. aureus during eight weeks of exposure. Each value is represented as the mean ± SE (n = 4). Root: Shoot Ratio (X 10-2) Nitrate Concentration (mg/L) Week 0 Week 2 Week 4 Week 6 Week 8 0 0.04 ± 0.04 a 1.93 ± 1.03 ab 6.52 ± 1.71 bcd 10.23 ± 0.82 efgh 16.38 ± 0.50 j 20 0.08 ± 0.08 a 1.24 ± 0.54 a 6.55 ± 1.43 bcd 13.70 ± 1.12 hij 13.14 ± 1.42 hij 50 0.00 ± 0.00 a 2.92 ± 0.62 abc 9.46 ± 2.93 efgh 15.60 ± 2.56 ij 11.69 ± 3.87 ghi 100 0.07 ± 0.07 a 3.05 ± 2.07 abc 6.97 ± 1.34 cde 15.55 ± 0.84 ij 13.03 ± 1.61 hij 200 0.00 ± 0.00 a 2.42 ± 2.12 abc 6.96 ± 1.06 cde 7.91 ± 3.23 efg 11.44 ± 1.10 fghi 69 The scatterplot obtained from plotting total plant DW against NO3- removal showed a positive linear relationship between the two parameters (Fig. 20). The correlation coefficient (r) was 0.582 which indicated a positive but weak correlation between the two parameters. Leaf Growth Total leaf area increased over the 8 weeks for all plants (Fig. 22). The largest increase in total leaf area was observed in plants exposed to 100 mg/L NO3- (1.5-fold) compared to W0. However, the largest increase in area of newly expanded leaves after NO3- addition (NL) was observed in plants exposed to 200 mg/L NO3- (from 0 to 66.53 cm2) (Fig. 21E). The area of existing leaves before NO3- treatment (OL) decreased in plants exposed to 0, 20, 50 and 200 mg/L NO3- and the largest decrease (34.8%) was observed in plants treated with 20 mg/L NO3- (Fig. 21B, C, E). FW per cm2 total leaf area showed no significant differences in plants treated with different NO3- concentrations across 8 weeks (Table 7A). However, DW per cm2 total leaf area increased in plants exposed to low NO3- concentrations from W0 to W8, but decreased in plants exposed to high NO3- concentrations (Table 7B). The total number of leaves increased in all plants during the 8 weeks of study (Fig 23A). The largest number of leaves was found in plants exposed to 100 and 200 mg/L NO3-. However, the leaf area for each individual leaf (Fig 10B) decreased in all plants from W0 to W8. On W8, the areas of each individual leaf in plants exposed to 100 and 200 mg/L NO3- were significantly much larger than those of plants exposed to 0 mg/L NO3-. 70 Fig. 20. Scatterplot of total DW (g) against NO3- removal (mg/L) from growth medium (NO3- spiked water). 71 (A) (B) (C) (D) (E) Fig. 21. Effects of various NO3- concentrations on leaf area of existing leaves before NO3- exposure and newly formed leaves after NO3- exposure. Fig 6A, B, C, D, E represent S. aureus exposed to 0, 20, 50, 100 and 200 mg/L NO3- respectively, during eight weeks of exposure. Each value is represented as the mean ± SE (n = 4). 72 Fig. 22 . Effects of various NO3- concentrations on total leaf area of S. aureus during eight weeks of exposure. Each value is represented as the mean ± SE (n = 4). 73 Table 7. Effects of various NO3- concentrations on (A) g FW per cm2 total leaf area and (B) g DW per cm2 total leaf area of S. aureus during eight weeks of exposure. Each value is represented as the mean ± SE (n = 4). (A) 2 -2 2 FW/ cm Total Leaf Area (X 10 g FW/ cm ) Nitrate Concentration (mg/L) Week 0 Week 2 Week 4 Week 6 Week 8 0 2.82 ± 0.14 a 2.44 ± 0.03 a 2.69 ± 0.41 a 2.65 ± 0.06 a 2.63 ± 0.4 a 20 2.70 ± 0.10 50 2.54 ± 0.10 a 100 2.63 ± 0.09 200 2.52 ± 0.11 a a a a 2.61 ± 0.28 a 2.45 ± 0.05 2.85 ± 0.16 a 2.66 ± 0.12 a 2.74 ± 0.37 a a 2.68 ± 0.50 a 2.65 ± 0.12 2.47 ± 0.07 2.74 ± 0.08 2.58 ± 0.08 a 2.65 ± 0.11 a a 2.54 ± 0.08 a 2.69 ± 0.59 a a 2.70 ± 0.04 2.70 ± 0.44 a a 2.82 ± 0.06 a (B) DW/ cm2 Total Leaf Area (X 10-2 g DW/ cm2) Nitrate Concentration (mg/L) Week 0 Week 2 Week 4 Week 6 Week 8 0 0.315 ± 0.023 abcdef 0.253 ± 0.003 abc 0.357 ± 0.054 f 0.355 ± 0.007 ef 0.323 ± 0.05 bcdef 20 0.302 ± 0.018 abcdef 0.283 ± 0.008 50 0.283 ± 0.009 abcdef 0.333 ± 0.019 cdef 0.301 ± 0.014 abcdef 0.273 ± 0.036 abcd 0.337 ± 0.074 def 100 0.292 ± 0.003 abcdef 0.342 ± 0.010 def 0.280 ± 0.052 abcdef 0.248 ± 0.012 ab 0.238 ± 0.004 a 200 0.281 ± 0.015 abcdef 0.315 ± 0.010 abcdef 0.314 ± 0.013 abcdef 0.311 ± 0.051 abcdef 0.248 ± 0.005 ab abcdef 0.321 ± 0.034 bcdef 0.273 ± 0.005 abcde 0.311 ± 0.010 abcdef 74 Fig. 23. Effects of various NO3- concentrations on (A) number of leaves produced and (B) total leaf area per leaf of S. aureus during eight weeks of exposure. Each value is represented as the mean ± SE (n = 4). 75 The scatterplot obtained from plotting total leaf area per plant against NO3removal showed a positive linear relationship between the two parameters (Fig. 24). The correlation coefficient (r) was 0.674 which indicated a positive but weak correlation between the two parameters. 4.2.3 Photosynthetic Pigment Concentrations Chlorophyll a (Chl a) Concentration On a FW basis, chl a concentration in plants exposed to 0 mg/L NO3- peaked on W2, but showed a decreasing trend thereafter (Fig. 25A). Chl a concentrations in plants exposed to 20 and 50 mg/L NO3- peaked on W4, and they decreased from W4 to W8. In plants exposed to 100 mg/L NO3-, chl a level increased from W0 to W2 but dropped sharply on W4, after which they continued to increase from W4 to W8. Chl a concentration in plants exposed to 200 mg/L NO3- exhibited a small increase on W8 as compared to that of W0. On a DW basis, changes in chl a concentration were very similar to those expressed on a FW basis (Fig. 25B). However, chl a level in plants exposed to 200 mg/L NO3- showed a larger increase on W8 (42%) compared to that of W0. Chlorophyll b (Chl b) Concentration Chl b concentration (mg/ g FW) showed very similar changes as those of chl a (Fig. 26A). Chl b level in plants exposed to 0 mg/L NO3- increased from W0 to W2, but decreased thereafter. Chl b concentrations in plants exposed to 20 and 50 mg/L NO3- peaked on W4, but they decreased from W4 onwards. Chl b level in plants exposed to 100 mg/L NO3- increased from W0 to W2, decreased on W4 but continued 76 Fig. 24. Scatterplot of total leaf area per plant (cm2/ plant) against NO3- removal (mg/L) from growth medium (NO3- spiked water). 77 (A) (B) Fig. 25. Effects of various NO3- concentrations on the concentrations of chlorophyll a during eight weeks of exposure. Concentrations are expressed in (A) mg per g FW and (B) mg per g DW. Each value is represented as the mean ± SE (n = 4). 78 (A) (B) Fig. 26. Effects of various NO3- concentrations on the concentrations of chlorophyll b during eight weeks of exposure. Concentrations are expressed in (A) mg per g FW and (B) mg per g DW. Each value is represented as the mean ± SE (n = 4). 79 to increase from W4 to W8. Plants exposed to 200 mg/L NO3- exhibited a continual increase in chl b level from W0 to W8. On a DW basis, changes in chl b concentrations were highly similar to those expressed on a FW basis (Fig 26B). However, chl b level in plants exposed to 200 mg/L NO3- showed a much larger increase on W8 (35%) compared to that of W0. Total Chlorophyll (Total Chl) Concentration The changes in total chl concentrations (mg/ g FW) followed those of chl a and chl b (Fig. 27A). Total chl concentration in plants exposed to 0 mg/L NO3increased from W0 to W2, but decreased from W2 to W6, after which it recovered on W8. In plants exposed to 20 and 50 mg/L NO3-, total chl levels increased from W0 to W4 but decreased thereafter. Total chl concentration in plants exposed to 100 mg/L NO3- peaked once on W2, then decreased on W4 but increased again from W4 to W8. Total chl level in plants exposed to 200 mg/L NO3- gradually increased from W0 to W8 in all plants. On W4, total chl concentrations of plants exposed to 20 and 50 mg/L NO3- were 25% and 32% higher respectively than that of plants exposed to 0 mg/L NO3-, whereas total chl concentrations of plants exposed to 100 and 200 mg/L NO3were 18% and 1% lower respectively than that of plants exposed to 0 mg/L NO3-. However, on W8, the total chl concentrations of plants exposed to 20 mg/L and 50 mg/L NO3- changed to 9% lower and 13% higher respectively than that of plants exposed to 0 mg/L NO3-, whereas total chl concentrations of plants exposed to 100 and 200 mg/L NO3- changed to 53% and 34% higher respectively than that of plants exposed to 0 mg/L NO3-. 80 Fig. 27. Effects of various NO3- concentrations on the concentrations of total chlorophylls during eight weeks of exposure. Concentrations are expressed in (A) mg per g FW and (B) mg per g DW. Each value is represented as the mean ± SE (n = 4). 81 Even when they were expressed on a DW basis (Fig. 27B), total chl concentrations in all plants showed the same trends as those expressed on a FW basis. On W4, total chl concentrations of plants exposed to 20 and 50 mg/L NO3- were 35% and 55% higher respectively than that of plants exposed to 0 mg/L NO3-, whereas total chl concentrations of plants exposed to 100 and 200 mg/L NO3- were 4% and 11% higher respectively than that of plants exposed to 0 mg/L NO3-. However, on W8, the total chl concentrations of plants exposed to 20 mg/L and 50 mg/L NO3- changed to 9% lower and 11% higher than that of plants exposed to 0 mg/L NO3-, whereas total chl concentrations of plants exposed to 100 and 200 mg/L NO3- both changed to 2-fold higher than that of plants exposed to 0 mg/L NO3-. Carotenoid Concentration Changes in carotenoid concentrations (mg/ g FW) in plants exposed to 0, 20, 50 and 100 mg/L NO3-showed similar trends to those of chl a and chl b concentrations (Fig. 28A). Carotenoid concentration in plants exposed to 0 mg/L NO3- peaked on W2, but decreased thereafter. Carotenoid concentrations in plants exposed to 20 and 50 mg/L NO3- peaked on W4, but decreased from W6 onwards. In plants exposed to 100 mg/L NO3-, carotenoid level increased from W0 to W2, decreased on W4, but increased again from W4 to W8. In plants exposed to 200 mg/L NO3- however, carotenoid concentration increased from W0 to W4, but it subsequently decreased from W4 to W8. On a DW basis (Fig. 28B), changes in carotenoid concentrations were highly similar to those expressed on a FW basis for plants exposed to 0, 20 and 50 mg/L NO3-. However, in plants exposed to 100 mg/L NO3-, carotenoid level remained relatively constant from W0 to W4 (1.19 – 1.22 mg/g DW) but increased thereafter. 82 (A) (B) Fig. 28. Effects of various NO3- concentrations on the concentrations of carotenoids during eight weeks of exposure. Concentrations are expressed in (A) mg per g FW and (B) mg per g DW. Each value is represented as the mean ± SE (n = 4). 83 Carotenoid level in plants exposed to 200 mg/L also remained relatively constant from W0 to W6, and increased from W6 to W8. Chlorophyll a: b (Chl a: b) Ratio The lowest chl a: b ratio was observed in plants exposed to 0 mg/L NO3- after 6 weeks of treatment, but the value increased again on W8 (Table 8). Chl a: b ratio was the highest in plants exposed to 200 mg/L NO3- for 8 weeks. 4.2.4 Chlorophyll Fluorescence Fv/ Fm and Fv/ Fo Values of Fv/ Fm decreased over the 8 weeks in all plants (Fig. 29A). The largest decrease (11%) was observed in plants exposed to 200 mg/L NO3- on W8 compared to that on W0. Fv/Fm showed the smallest decrease in plants exposed to 20 and 100 mg/L NO3- (6% and 7% respectively) from W0 to W8. Values of Fv/ Fo also decreased from W0 to W8 in plants exposed to 0, 20, 50 and 100 mg/L NO3-, with the largest decrease in these plants observed from W0 to W2 (Fig. 29B). However, in plants exposed to 200 mg/L NO3-, Fv/Fo peaked on W4, but decreased thereafter. Fv’/ Fm’, ФPS II, qp and NPQ Values of Fv’/ Fm’ (Fig. 30A) generally decreased in all plants from W0 to W8. The largest decrease was observed in plants exposed to 200 mg/L NO3-, with Fv’/ Fm’ values decreasing by 22% from W0 to W8. Both ФPSII and qp values decreased in all plants from W0 to W8 (Fig. 30B and 31A). Plants exposed to 200 mg/L exhibited the largest fluctuations in both values, 84 Table 8. Effects of various NO3- concentrations on chlorophyll a: b ratio of S. aureus during eight weeks of exposure. Each value is represented as the mean ± SE (n = 4). Nitrate Concentration (mg/L) Chlorophyll a: b ratio Week 0 Week 2 Week 4 Week 6 Week 8 0 2.27 ± 0.12 ab 2.09 ± 0.11 ab 2.20 ± 0.21 ab 1.56 ± 0.56 a 2.10 ± 0.07 ab 20 2.27 ± 0.12 ab 2.19 ± 0.08 ab 2.37 ± 0.08 ab 2.16 ± 0.06 ab 2.07 ± 0.04 b 50 2.27 ± 0.12 ab 2.29 ± 0.06 ab 2.24 ± 0.06 ab 2.40 ± 0.02 ab 2.28 ± 0.07 ab 100 2.27 ± 0.12 ab 200 2.27 ± 0.12 ab 2.21 ± 0.02 ab 2.13 ± 0.04 ab 2.35 ± 0.08 ab 2.44 ± 0.08 ab 2.40 ± 0.04 ab ab 2.43 ± 0.03 ab 2.49 ± 0.10 2.15 ± 0.08 c 85 (A) (B) Fig. 29. Effects of various NO3- concentrations on (A) Fv/Fm and (B) Fv/Fo during eight weeks of exposure. Each value is represented as the mean ± SE (n = 4). 86 with both ФPSII and qp values on W4 similar to those of W0. In plants exposed to 20 mg/L NO3-, ФPSII and qp values decreased from W0 to W2, but subsequently increased on W4 and stayed relatively constant from W4 to W8. ФPSII and qp values for plants exposed to 100 mg/L NO3- stayed relatively constant from W0 to W2, but decreased from W2 to W4 and stayed relatively constant from W4 onwards. NPQ values (Fig. 31B) in all plants showed an increasing trend from W0 to W8, despite the large decrease observed from W0 to W2 in plants exposed to 50, 100 and 200 mg/L NO3- (21%, 35% and 23% respectively). 4.2.5 Total Nitrogen in Plant Tissues (Kjeldahl) Total nitrogen (TN) in plants (Fig. 33) treated with 0, 20 and 50 mg/L NO3increased from W0 to W2, but subsequently decreased from W2 onwards. TN in plants treated with 100 mg/L NO3- increased from W0 to W4, but decreased from W4 onwards. In plants treated with 200 mg/L NO3-, TN increased from W0 to W4, dropped on W6, but increased again on W8. On W8, TN for all plants was higher TN in leaves (Fig. 32) decreased from W0 to W8 in plants treated to 0, 20 and 50 mg/L NO3-, but increased from W0 to W8 in plants treated with 100 and 200 mg/L NO3-. TN in roots (Fig. 32) increased from W0 to W8 in all plants, with the largest increase observed in plants treated with 200 mg/L NO3-. Total root nitrogen: total shoot nitrogen ratio (TNR: TNS) of plants exposed to 20, 50, 100 and 200 mg/L NO3- peaked on W2 whereas it peaked on W4 for plants exposed to 0 mg/L NO3- (Table 9). On W4 and W6, TNR: TNS was highest for control plants (not exposed NO3-) as compared to plants exposed to NO3-. On W8, however, TNR: TNS was highest in plants exposed to 50 mg/L NO3- (1.04 ± 0.25). 87 (A) (B) Fig. 30. Effects of various NO3- concentrations on (A) Fv’/Fm’ and (B) ФPSII during eight weeks of exposure. Each value is represented as the mean ± SE (n = 4). 88 (A) (B) Fig. 31. Effects of various NO3- concentrations on (A) qp and (B) NPQ during eight weeks of exposure. Each value is represented as the mean ± SE (n = 4). 89 (A) (B) (C) (D) (E) Fig. 32. Effects of various NO3- concentrations on total nitrogen of leaves and roots. Fig. 32A, B, C, D, E represent S. aureus exposed to 0, 20, 50, 100 and 200 mg/L NO3respectively, during eight weeks of exposure. Each value is represented as the mean ± SE (n = 4). 90 Fig. 33. Effects of various NO3- concentrations on total nitrogen in whole plants of S. aureus during eight weeks of exposure. Each value is represented as the mean ± SE (n = 4). 91 Table 9. Effects of various NO3- concentrations on total root nitrogen: total shoot nitrogen ratio of S. aureus during eight weeks of exposure. Each value is represented as the mean ± SE (n = 4). Nitrate Total Root Nitrogen: Total Shoot Nitrogen Ratio Concentration (mg/ L) Week 2 Week 4 Week 6 Week 8 0 0.84 ± 0.49 ab 1.20 ± 0.16 bcde 1.06 ± 0.09 cd 0.83 ± 0.05 ab 20 1.90 ± 0.64 e 1.07 ± 0.09 abc 0.78 ± 0.03 ab 0.95 ± 0.06 abc 50 1.84 ± 0.15 de 1.09 ± 0.10 abcd 0.80 ± 0.06 ab 1.04 ± 0.25 abc 100 1.65 ± 0.64 cde 0.93 ± 0.04 abc 0.70 ± 0.05 ab 0.74 ± 0.01 ab 200 0.37 ± 0.37 a 1.08 ± 0.07 abc 0.76 ± 0.26 ab 0.73 ± 0.06 ab 92 The scatterplot obtained from plotting total leaf area per plant against TN of the whole plant showed that these two parameters were not correlated (Fig. 34). 4.2.6 Total Soluble Sugar (TSS) Concentration The concentration of total soluble sugars (TSS) (expressed on a FW basis) decreased from W0 to W8 in all plants (Fig. 35A). The lowest TSS levels were observed on W4 in plants exposed to 0, 20, 50 and 200 mg/L NO3-. In plants exposed to 100 mg/L NO3-, TSS decreased (37%) from W0 to W6, but increased from W6 to W8. A similar trend was observed for TSS levels expressed in terms of per g DW (Fig. 35B). The lowest TSS levels were also observed on W4 in plants exposed to 0, 20, 50 and 200 mg/L NO3-. In plants exposed to 100 mg/L NO3-, TSS decreased (40 %) from W0 to W2, but increased from W2 onwards. The scatterplots obtained from plotting TSS (mg/ g FW) against Fv/Fm, Fv/Fo, Fv’/Fm’, ФPSII, qp and NPQ showed that these parameters were not correlated (Fig. 36). 4.2.7 Total Soluble Protein (TSP) Concentration TSP concentrations were determined for leaves that were present on the plants even before NO3- addition (OL) and the first newly expanded leaf of each plant after NO3- addition (NL), in order to determine the change that took place in the various leaf types. 93 Fig. 34. Scatterplot of total leaf area per plant (cm2/ plant) against total nitrogen (mg N/ g DW) in whole plants of S. aureus. 94 (A) (B) Fig. 35. Effects of different NO3- concentrations on total soluble sugar concentration in leaves during eight weeks of exposure. All concentrations are expressed in (A) mg glucose equivalent per g FW and (B) mg glucose equivalent per g DW. Each value is represented as the mean ± SE (n = 4). 95 Fig. 36. Scatterplots of total soluble sugar concentration (mg/ g FW) against chlorophyll fluorescence parameters, (A) Fv/Fm, (B) Fv/Fo, (C) Fv’/Fm’, (D) ФPSII, (E) qp and (F) NPQ of leaves of S. aureus plants exposed to NO3- for 8 weeks. 96 TSP Concentration of existing leaves (OL) TSP levels (expressed on a FW basis) decreased in plants exposed to 0 and 20 mg/L NO3- from W0 to W8, but they increased from W0 to W8 in plants exposed to 50, 100 and 200 mg/L NO3- (Fig. 37). The same trend was observed for TSP levels expressed in terms of per g DW. TSP Concentration of first fully expanded leaves (NL) TSP concentrations could only be determined from W2 onwards when NLs were formed (Fig. 38). TSP concentrations decreased from W0 to W8 in plants exposed to 0, 20 and 50 mg/L NO3-, but they increased from W0 to W8 in plants exposed to 100 and 200 mg/L NO3-. The same trend was observed for TSP levels expressed in terms of per g DW. 4.2.8 Nitrate Reductase (NR) Activity The activity of NR was determined in leaves that were present on the plants even before NO3- addition (OL), the first newly expanded leaf of each plant after NO3addition (NL) and the roots, in order to determine the change that took place in the various organs. NR Activity in existing leaves (OL) of S. aureus The activity of NR (µM NO2-/ g FW/ h) in OL generally increased in all plants from W0 to W8 (Fig. 39A). Plants exposed to 0 mg/L NO3-showed a continual increase (3-fold) in NR activity from W0 to W8. NR activity in plants treated with 20 mg/L NO3- also increased by 4-fold from W0 to W8, with a slight decrease from W0 to W2. In plants exposed to 50 mg/L NO3-, NR activity increased from W0 to W2, but 97 (A) (B) Fig. 37. Effects of different NO3- concentrations on total soluble protein concentration in existing leaves before NO3- exposure, for a period of eight weeks. All concentrations are expressed in (A) mg BSA equivalent per g FW and (B) mg BSA equivalent per g DW. Each value is represented as the mean ± SE (n = 4). 98 (A) (B) Fig. 38. Effects of different NO3- concentrations on total soluble protein concentration in newly formed leaves after NO3- exposure, for a period of eight weeks. All concentrations are expressed in (A) mg BSA equivalent per g FW and (B) mg BSA equivalent per g DW. Each value is represented as the mean ± SE (n = 4). 99 decreased from W2 to W4, after which it increased again from W4 to W8. NR activity of plants exposed to 100 mg/L NO3- decreased slightly from W0 to W2 but eventually increased from W2 to W8. Plants treated with 200 mg/L NO3-, however, showed a sharp increase (3-fold) from W0 to W2 in NR activity, but it decreased from W2 to W6 and subsequently increased sharply again from W6 to W8. On a per mg protein basis (Fig. 39B), the activity of NR increased in all plants over 8 weeks. Plants exposed to 20 mg/L NO3- showed the largest increase (4.5-fold) from W0 to W8. Plants exposed to 0 mg/L NO3- also showed a large increase (6.5fold) in NR activity from W0 to W6, but it decreased from W6 onwards. NR Activity in first fully expanded leaves (NL) of S. aureus NLs were observed only from W2 onwards and NR activity was relatively low in all plants from W2 to W4 (Fig. 40A). NR activity (µM NO2-/ g FW/ h) in plants exposed to 0 mg/L NO3- increased from W2 to W8. NR activity in plants exposed to 20, 50 and 100 mg/L NO3- sharply decreased from W2 to W4, but increased from W4 to W8. Plants treated with 200 mg/L NO3- also showed a similar trend in NR activity from W4 to W6, but it dropped on W8. On a per mg protein basis, NR activity remained low from W2 to W4 in all plants, but it increased sharply on W6 (Fig 40B). Subsequently, NR activity increased in plants exposed to 20 mg/L NO3- on W8 but decreased in plants exposed to all other NO3- concentrations. NR Activity in roots of S. aureus New roots were only formed from W2 onwards. On a FW basis, the activity of 100 (A) (B) Fig. 39. Effects of different NO3- concentrations on nitrate reductase activity in existing leaves before NO3- exposure, for a period of eight weeks. All concentrations are expressed in (A) µM NO2-/ g FW/ h and (B) µM NO2-/ mg protein/ h. Each value is represented as the mean ± SE (n = 4). 101 NR in roots (µM NO2-/ g FW/ h) was relatively constant throughout the period of study (Fig. 41A), except in plants exposed to 0 mg/L NO3-. In these plants, NR activity increased significantly on W4. NR activity in plants exposed to 20 mg/L NO3also showed a steady increase from W2 to W8. In plants treated with 100 mg/L NO3-, NR activity also increased significantly on W6. On a per mg protein basis (Fig. 41B), the activity of NR in roots increased from W2 to W6 in all plants. NR activity continued to increase from W6 to W8 only in plants exposed to 0, 20 and 50 mg/L NO3-, but decreased in plants exposed to 100 and 200 mg/L NO3-. Plants exposed to 20 mg/L NO3- exhibited the largest increase (7fold) in root NR activity from W2 to W8. 4.2.9 Lipid Peroxidation Level (TBARs) TBARs (µMol MDA/ g FW) of plants exposed to 0 mg/L NO3- increased on W2, and then decreased and remained constant until W8 (Fig. 42A). In plants exposed to 20 mg/L NO3-, TBARs peaked on W4, but decreased and remained constant from W4 to W8. TBARs of plants exposed to 50 mg/L NO3- peaked on W2, decreased from W2 to W6, and then increased again on W8. In plants exposed to 100 mg/L NO3-, TBARs decreased from W0 to W4, but increased on W6, before reaching the same value as W0 on W8. In plants exposed to 200 mg/L NO3-, TBARs was highest on W2 and lowest on W4. On a per g DW basis (Fig. 42B), TBARs in plants exposed to 0 mg/L NO3increased on W2, and decreased thereafter. In plants exposed to 20 mg/L NO3-, TBARs also gradually decreased from W2 onwards. TBARS in plants exposed to 50 and 200 mg/L NO3- peaked on W2 but decreased from W2 to W4, it then increased 102 (A) (B) Fig. 40. Effects of different NO3- concentrations on nitrate reductase activity in newly formed leaves after NO3- exposure, for a period of eight weeks. All concentrations are expressed in (A) µM NO2-/ g FW/ h and (B) µM NO2-/ mg protein/ h. Each value is represented as the mean ± SE (n = 4). 103 (A) (B) Fig. 41. Effects of different NO3- concentrations on nitrate reductase activity in roots for a period of eight weeks. All concentrations are expressed in (A) µM NO2-/ g FW/ h and (B) µM NO2-/ mg protein/ h. Each value is represented as the mean ± SE (n = 4). 104 again on W6 but on W8, TBARs decreased in plants exposed to 50 mg/L NO3-, whereas it increased on W8 in plants exposed to 200 mg/L NO3-. 4.2.10 Leaf Antioxidant Enzymes Leaf Ascorbate Peroxidase (APX) Activity The specific activity of APX (U/ mg protein) displayed a general increase from W0 to W8 for all plants; it was lowest at the beginning of the study (Fig. 43A). In plants exposed to 0 mg/L NO3-, APX specific activity increased from W0 to W8. APX specific activity also increased from W0 to W6 in plants exposed to 20 mg/L NO3-, but it decreased slightly on W8. In plants exposed to 50 mg/L NO3-, APX specific activity increased from W0 to W8, but there was a sharp dip observed on W4. In plants exposed to 100 mg/L NO3-, APX activity exhibited a sharp increase from W2 to W4 and remained almost constant until W8. APX specific activity in plants exposed to 200 mg/L NO3- peaked on W2, but decreased on W4, after which it remained relatively constant about (0.08 ± 0.002) U/ mg protein. On a per g FW basis (Fig. 43B), APX activity increased from W0 to W8 in all plants, with plants exposed to 50 and 200 mg/L NO3- showing a drop in APX activity on W4. Leaf Glutathione Reductase (GR) Activity The specific activity of GR (U/ mg protein) decreased from W0 to W8 in all plants except those exposed to 50 mg/L NO3- (Fig. 44A). GR specific activity in plants exposed to 20 mg/L NO3- decreased (53%) from W0 to W2 and it decreased even further on W4. It increased from W4 to W6 but decreased again on W8. Plants exposed to 200 mg/L NO3- exhibited a sharp rise in GR activity on W4 but it decreased from W6 onwards. 105 (A) (B) Fig. 42. Effects of different NO3- concentrations on lipid peroxidation levels (TBARs) of leaves, expressed in (A) µM MDA/ g FW and (B) µM MDA/ g DW, during eight weeks of exposure. Each value is represented as the mean ± SE (n = 4). 106 (A) (B) Fig. 43. Effects of different NO3- concentrations on the activity of leaf ascorbate peroxidase, expressed in (A) U/ mg protein and (B) U/ g FW, during eight weeks of exposure. Each value is represented as the mean ± SE (n = 4). 107 On a per g FW basis (Fig. 44B), GR activity increased in plants exposed to 0 and 50 mg/L NO3- from W0 to W8, but it decreased from W0 to W8 in plants exposed to 20, 100 and 200 mg/L NO3- during the same period. On W4, plants exposed to 20 mg/L NO3- displayed the lowest GR activity whereas plants exposed to 200 mg/L NO3- showed the highest GR activity. Leaf Superoxide Dismutase (SOD) Activity The specific activity of SOD (U/ mg protein) exhibited a sharp increase in all plants on W2, and it was lowest in all plants at W0 (Fig. 45A). In plants exposed to 0 and 20 mg/L NO3-, SOD specific activity decreased from W2 to W4, but increased from W4 to W6, after which they decreased again. In plants exposed to 50 and 100 mg/L NO3-, SOD specific activities increased from W2 to W6, but decreased on W8. SOD activity in plants exposed to 200 mg/L NO3- peaked on W4, but decreased from W6 onwards. On a per g FW basis, SOD activity also increased by about 3-fold in all plants from W0 to W2 (Fig. 45B). SOD activity in plants exposed to 0, 20, 50 and 200 mg/L NO3- then decreased from W2 to W4, but increased on W6, after which they decreased again on W8. In plants exposed to 100 mg/L NO3-, SOD activity remained relatively constant (601.96 – 636.15 U/ g FW) from W2 to W6, but dropped from W6 to W8. Leaf Guaiacol Peroxidase (POD) Activity The specific activity of POD (U/ mg protein) generally decreased from W0 to W8 in all plants; it was highest on W0 (Fig. 46A). POD specific activity was the lowest on W4 in plants exposed to 0, 20, 50 and 100 mg/L NO3- . In plants exposed to 108 (A) (B) Fig. 44. Effects of different NO3- concentrations on the activity of leaf glutathione reductase, expressed in (A) U/ mg protein and (B) U/ g FW, during eight weeks of exposure. Each value is represented as the mean ± SE (n = 4). 109 (A) (B) Fig. 45. Effects of different NO3- concentrations on the activity of leaf superoxide dismutase, expressed in (A) U/ mg protein and (B) U/ g FW, during eight weeks of exposure. Each value is represented as the mean ± SE (n = 4). 110 200 mg/L NO3-, POD specific activity increased at W4, but it dropped on W6 and remained relatively constant at 0.01 U/ mg protein. On a per g FW basis (Fig. 46B), POD activity was also lowest on W4, in plants exposed to 0, 20, 50 and 100 mg/L NO3-. In plants exposed to 100 mg/L NO3-, POD activity peaked on W2, fell to its lowest on W4, but it then increased from W6 to W8 to a level higher than that of W0. POD activity in plants exposed to 200 mg/L NO3- continuously decreased from W0 to W8. 111 (A) (B) Fig. 46. Effects of different NO3- concentrations on the activity of leaf guaiacol peroxidase, expressed in (A) U/ mg protein and (B) U/ g FW, during eight weeks of exposure. Each value is represented as the mean ± SE (n = 4). 112 4.3 Phytoremediation of Bisphenol A by S. aureus 4.3.1 Removal of BPA by S. aureus To ensure that only S. aureus plants were removing BPA from the BPAspiked water, the entire experiment was conducted under aseptic culture conditions (Fig. 47). Under such conditions, phytoremediation of BPA started even after 3 h of exposing S. aureus plants to BPA-containing water (Fig. 48). After 24 h (Day 1; D1) of exposure, S. aureus removed more than 50% of the BPA in the 50 and 100 µM BPA-containing waters. Complete phytoremediation of BPA by S. aureus plants was observed after 48 h (Day 2; D2) of exposure. However, when the plants were exposed to 250 µM BPA, only about 50% of BPA was removed by the plants by the end of 168 h (Day 7; D7) of exposure. The profiles from HPLC analyses showed that BPA was not only taken up by the plant tissues, but it could be converted into another compound in the water (Fig. 49C, 50). Secondary peaks (retention time 15.93 – 16.7 min) were observed very close to the peak corresponding to BPA (retention time 16.97 – 17.05 min) on the HPLC profiles and they increased in height with decreasing BPA concentration throughout the study period. The water samples were also observed to change to a yellowish colour (Fig. 51), which became darker with increasing initial BPA concentration and duration of exposure. The new peak and change in color were not observed in the controls without plant material. The plants were harvested and frozen in liquid nitrogen only after seven days of exposure. The frozen plants were ground in 100% methanol and the extract was centrifuged at 2,500 g for 10 min. The supernatant was then removed and evaporated 113 Fig. 47. S. aureus plants exposed to water containing (A) 0, (B) 50, (C) 100 and (D) 250 µM BPA under aseptic conditions. Bar = 1cm. Photographs are representative of 4 replicates. 114 Fig. 48. Removal of BPA from the growth medium (BPA-spiked water) by S. aureus. Roots of the plants were submerged in water containing 0, 50, 100 and 250 µM of BPA (closed markers, solid lines). Experimental controls with the same BPA concentrations were also set up but without plants growing in them (open markers, dotted lines). Each value is represented as the mean ± SE (n = 4). 115 to dryness and resuspended in pure ethanol. One hundred µl of the resultant extract were analysed by HPLC with the same method used to quantify BPA for the polluted growth medium. No trace of BPA was detected in the plant extracts (Fig. 49 D, E, F), even from plants exposed to 250 µM of BPA. 4.3.2 Growth Parameters Fresh weight and Dry Weight Since all the plants were cultivated from one cm long terminal shoot tips and provided with the same amount of growth medium for the same period of time, they were approximately of the same size on Day 0 (D0) at the beginning of the study. Intact plants (including roots) were used throughout the entire experiment to ensure that changes in the plant biomass of the plants could be more accurately determined. A total of 12 replicates for each treatment were used to determine the changes in fresh weight (FW) and dry weight (DW). Exposure of the plants to BPA for 7 days led to a general decrease in both FW and DW of the plants. Both leaf and root FW (Fig. 52A, B, C, D) and DW (Fig. 52E, F, G, H) were clearly affected by the culture conditions. Plants exposed to 100 µM BPA for 7 days exhibited the largest decrease in leaf FW (27%) and leaf DW (40%), compared to those at the beginning of the study (D0). The largest decrease in root FW (18%) and root DW (63%), from D0 to D7, were observed in plants exposed to 250 µM BPA. High BPA concentrations resulted in larger decreases in both total FW (Fig. 53A) and DW (Fig. 53B) of whole plants as compared to those exposed to 0 µM BPA. Plants grown in 100 µM BPA seemed to be the most affected by BPA exposure, with total FW and DW of the whole plant decreasing by 22% and 47% respectively from 116 Fig. 49. HPLC profiles of water samples (50 µM) on D7 of (A) BPA (1 mM) standards, (B) control set-up (without plants) and (C) set-up with S. aureus plants. HPLC profiles of (D) BPA (1 mM) and plant extracts on D7 obtained from (E) leaves and (F) roots are also shown (different gradient). The HPLC profiles indicated the depletion of BPA (black arrow) and the appearance of a secondary peak (dashed arrow) in the plant set-up but not the control set-up. Each profile is representative of four replicates for each BPA concentration. 117 Fig. 50. Close-up of HPLC profile of water samples (50 µM) on D7 of set-up with S. aureus plants. Peaks 1, 2 and 3 are peaks that appeared only in plant set-ups but not control set-ups. Peak 4 is representative of the presence of BPA. HPLC profile is representative of four replicates for each BPA concentration. 118 Fig. 51. Changes in the colour of liquid growth medium containing BPA concentrations of 0 µM (A), 50 µM (B), 100 µM (C) and 250 µM (D) by the end of D1. Bar = 1 cm. Photographs are representative of 4 replicates. 119 Fig. 52. Effects of various BPA concentrations on fresh weight (FW) and dry weight (DW) of S. aureus. Fig. 49A, B, C, D represent FW, whereas Fig. 49E, F, G, H represent DW of leaves and roots from plants exposed to 0, 50, 100 and 250 µM BPA, respectively, during seven days of exposure. Each value is represented as the mean ± SE (n = 12). 120 Fig. 53. Effects of various BPA concentrations on (A) total FW and (B) total DW of whole plants of S. aureus during seven days of exposure. Each value is represented as the mean ± SE (n = 12). 121 D0 to D7. However, the decreases in total FW and DW of the whole plant in plants exposed to 0 µM BPA were also high at 17.1% and 46.6% respectively, from D0 to D7. FW: DW ratios were calculated to determine whether the plants would show any changes in plant-water relationships (Table 10). In all plants, FW: DW ratios increased from D0 to D7. The largest increase (24%) was observed in plants exposed to 0 µM BPA. Plants exposed to 50, 100 and 250 µM BPA showed smaller increases in FW: DW ratio at 16%, 18% and 17% respectively, compared to those at D0. Root: shoot ratio showed an increasing trend from W0 to W8 in all plants (Table 11). The largest increase was observed in plants exposed to 100 µM BPA, whereas the smallest increase was observed in plants exposed to 0 µM BPA. Root growth On D7, roots of plants exposed to 0 (Fig. 54A, E) and 50 µM BPA (Fig. 54B, F) were still growing with white root tips. Root growth was also observed in plants exposed to 100 µM BPA but the tips of these roots became slightly black in colour (Fig. 54C, G). The root tips of plants exposed to 250 µM BPA were entirely black and roots did not show new growth (Fig. 54D, H). 4.3.3 Photosynthetic Pigment Content Chlorophyll a (Chl a), Chlorophyll b (Chl b) and Total Chlorophyll (Total Chl) Concentrations Changes in concentrations (mg/ g FW) of chl a (Fig. 55A) and chl b (Fig. 56A), and total chl (Fig. 57A) were similar. Compared to D0, a decrease in total chl 122 Table 10. Effects of various BPA concentrations on FW: DW ratio of S. aureus during seven days of exposure. Each value is represented as the mean ± SE (n = 12). BPA concentration FW : DW Ratio (µM) Day 0 Day 1 Day 4 Day 7 0 12.2 ± 1.1 a 13.2 ± 0.2 ab 13.8 ± 0.2 abc 15.2 ± 0.2 c 50 12.2 ± 1.1 a 12.8 ± 0.3 ab 13.7 ± 0.3 abc 14.2 ± 0.2 bc 100 12.2 ± 1.1 a 13.2 ± 0.4 ab 13.7 ± 0.2 abc 14.4 ± 0.3 bc 250 12.2 ± 1.1 a 12.9 ± 0.2 ab 14.0 ± 0.2 bc 14.2 ± 0.3 bc 123 Table 11. Effects of various BPA concentrations on root: shoot ratio of S. aureus during seven days of exposure. Each value is represented as the mean ± SE (n = 12). BPA concentration Root : Shoot Ratio (µM) Day 0 Day 1 Day 4 0 0.37 ± 0.03 abc 0.52 ± 0.05 d 0.36 ± 0.03 abc 0.37 ± 0.03 50 0.37 ± 0.03 abc 0.35 ± 0.05 abc 0.32 ± 0.04 ab 0.40 ± 0.05 bc 100 0.37 ± 0.03 250 0.37 ± 0.03 abc abc 0.37 ± 0.03 abc 0.39 ± 0.04 bc 0.27 ± 0.04 Day 7 a 0.40 ± 0.03 bc 0.47 ± 0.05 abc cd 0.41 ± 0.05 bcd 124 Fig. 54. Root tips of S. aureus submerged in liquid growth medium containing BPA concentrations of 0 µM (A, E), 50 µM (B, F), 100 µM (C, G) and 250 µM (D, H) by the end of D7. Bar = 1 cm. Photographs are representative of 4 replicates. 125 concentration was observed on D1 for all plants, and it decreased further on D4 for plants exposed to 50, 100 and 250 µM BPA. From D0 to D4, the decreases in chl a and chl b concentrations in plants exposed to 250 µM BPA resulted in the greatest fall in total chl concentration. By D7, total chl concentration significantly increased (compared to D4) in plants exposed to 50, 100 and 250 µM BPA. On D7, plants exposed to 100 and 250 µM BPA exhibited almost similar total chl concentrations as those on D0. This recovery, however, was not observed in plants exposed to 50 µM BPA. In plants exposed to 0 µM BPA, both chl a and chl b concentrations increased from D0 to D7 resulting in an increase in total chl concentration. When expressed in terms of per g DW, chl a (Fig. 55B), chl b (Fig. 56B) and total chl (Fig. 57B) concentrations showed similar trends as those expressed in per g FW. Chlorophyll a: b (Chl a: b) Ratio Chl a: b ratio (Table 12) generally increased in all plants upon transfer to liquid growth medium with BPA. It was, however, observed to be the lowest on D7 of exposure to BPA. The chl a: b ratio in all the plants was observed to rise slightly in those exposed to 0, 50 and 100 µM BPA from D0 to D4, but the values decreased on D7. In plants exposed to 250 µM, the chl a: b ratio showed a slight increase from D0 to D1, but it decreased from D4 to D7. Carotenoid Concentrations A decrease in carotenoid concentration (mg/ g FW) was observed in plants exposed to 50, 100 and 250 µM BPA from D0 to D4 (Fig. 58A). Carotenoid concentration continued to decrease in plants exposed to 100 and 250 µM BPA on D7, 126 (A) (B) Fig. 55. Effects of various BPA concentrations on the concentrations of chlorophyll a during seven days of exposure. Concentrations are expressed in (A) mg per g FW and (B) mg per g DW. Each value is represented as the mean ± SE (n = 4). 127 Fig. 56. Effects of various BPA concentrations on the concentrations of chlorophyll b during seven days of exposure. Concentrations are expressed in (A) mg per g FW and (B) mg per g DW. Each value is represented as the mean ± SE (n = 4). 128 (A) (B) Fig. 57. Effects of various BPA concentrations on the concentrations of total chlorophylls during seven days of exposure. Concentrations are expressed in (A) mg per g FW and (B) mg per g DW. Each value is represented as the mean ± SE (n = 4). 129 Table 12. Effects of various BPA concentrations on chlorophyll a: b ratio of S. aureus during seven days of exposure. Each value is represented as the mean ± SE (n = 4). BPA concentration (µM) Chlorophyll a : b Ratio Day 0 Day 1 Day 4 Day 7 0 2.38 ± 0.12 abc 2.53 ± 0.03 cd 2.82 ± 0.14 d 2.23 ± 0.11 ab 50 2.38 ± 0.12 abc 2.38 ± 0.12 abc 2.60 ± 0.09 cd 2.20 ± 0.07 ab 100 2.38 ± 0.12 abc 2.45 ± 0.12 bc 2.67 ± 0.12 cd 2.10 ± 0.04 a 250 2.38 ± 0.12 abc 2.57 ± 0.03 cd 2.53 ± 0.10 c 2.22 ± 0.08 ab 130 but plants exposed to 50 and 250 µM BPA showed signs of recovery. Plants exposed to 0 µM BPA showed a slight decrease in carotenoid concentration from D0 (0.28 mg/ g FW) until D4 (0.27 mg/ g FW), but were able to restore the carotenoids concentration close to that of D0 by D7. Trends observed for carotenoid concentrations expressed on a per g DW basis (Fig. 58B) was similar to those expressed on a per g FW basis. 4.3.4 Chlorophyll Fluorescence Fv/Fm and Fv/Fo ratios All plants exhibited high Fv/Fm values (0.78 – 0.85) throughout the 7 days of exposure to BPA (Fig. 59A), but these values decreased slightly with exposure to increasing BPA concentrations. To obtain a clearer picture of whether PSII photosynthetic efficiency was affected by exposure to BPA, Fv/Fo values were calculated and plotted (Fig. 59B). When the plants were initially transferred from agar to the liquid growth medium (D1), Fv/Fo increased (compared to D0) in all the plants, including plants exposed to 0 µM BPA. On D7, plants exposed to 100 and 250 µM BPA showed significant decreases in Fv/Fo as compared to those on D0, with the largest decrease in Fv/Fo values observed in plants exposed to 250 µM BPA (16%). Plants exposed to 50 µM BPA showed an almost similar Fv/Fo value as those on D0. Fv’/Fm’ ratio, ФPSII, photochemical quenching (qP) and non-photochemical quenching (NPQ) When initially transferred from solid agar medium to the liquid growth medium (D1), increases in Fv’/Fm’ (compared to D0) were observed in all plants (Fig. 60A). On D7, plants exposed to 50, 100 and 250 µM BPA showed significant 131 (A) (B) Fig. 58. Effects of various BPA concentrations on the concentrations of carotenoids during seven days of exposure. Concentrations are expressed in (A) mg per g FW and (B) mg per g DW. Each value is represented as the mean ± SE (n = 4). 132 (A) (B) Fig. 59. Effects of various BPA concentrations on (A) Fv/Fm and (B) Fv/Fo during seven days of exposure. Each value is represented as the mean ± SE (n = 4). 133 decreases in Fv’/ Fm’ as compared to the respective controls on D0 (8%, 11% and 13% respectively). Changes in ФPSII were similar to those of Fv’/Fm’ (Fig. 60B). When initially transferred from agar to the liquid growth medium (D1), ФPSII first increased, and then decreased from D4 to D7 in all plants. All plants exposed to BPA showed significant decreases in ФPSII as compared to the controls on D0. On D7, ФPSII of plants exposed to 100 and 250 µM BPA decreased by 24% and 28 % respectively, when compared to the controls on D0. On D7, ФPSII of plants exposed to 50 µM BPA was similar to that of D0. Changes in qP were also similar to those of Fv’/Fm’ and ФPSII (Fig. 61A). When initially transferred from agar to the liquid growth medium (D1), qP increased (compared to D0) in all plants. On D7, qp of plants exposed to 50, 100 and 250 µM BPA decreased by 11%, 14% and 18% respectively, compared to those of control plants on D0. NPQ is calculated via the Stern-Volmer equation (Eickmeier et al., 1993). When initially transferred from agar to the liquid growth medium (D1), NPQ (compared to D0) decreased in plants exposed to 0 and 250 µM BPA, but not for plants exposed to 50 and 100 µM BPA (Fig. 61B). On D7, NPQ increased significantly in all plants compared to their controls on D0. This increase was high for plants exposed to 50 µM BPA (30%), as compared to the other BPA concentrations (100 µM = 15%; 250 µM = 19%). Plants of the control set-up (not exposed to BPA) also showed a 31% increase in NPQ on D7 compared to that of D0. 134 Fig. 60. Effects of various BPA concentrations on (A) Fv’/Fm’ and (B) ФPSII during eight seven days of exposure. Each value is represented as the mean ± SE (n = 4). 135 (A) (B) Fig. 61. Effects of various BPA concentrations on (A) qp and (B) NPQ during seven days of exposure. Each value is represented as the mean ± SE (n = 4). 136 4.3.5 Total Soluble Sugar (TSS) Concentration The leaf TSS concentration (expressed as both per g FW and per g DW) in plants exposed to 50, 100 and 250 µM BPA showed significant increases on D1 compared to D0 (Fig. 62). No increase in TSS concentration was observed in plants grown in 0 µM BPA. However, on D4, the TSS concentration in all the plants showed a 2-fold decrease as compared to those on D1. They then continued to decrease on D7 in plants exposed to 0 and 50 µM BPA. The scatterplots obtained from plotting TSS (mg/ g FW) against Fv/Fm, Fv/Fo and NPQ showed that these parameters were not correlated (Fig. 63A, B, F). But scatterplots obtained from plotting TSS against Fv’/Fm’, ФPSII and qp showed a positive linear relationship between the parameters (Fig. 63C, D, E). The correlation coefficients (r) were 0.388, 0.482 and 0.395 respectively, which suggested positive but weak correlations between the various chlorophyll fluorescence parameters and total soluble sugar concentrations. The scatterplot obtained from plotting TSS against NPQ however, suggested a weak negative correlation with a correlation coefficient of -0.177 (Fig. 63F). 4.3.6 Total Soluble Protein (TSP) Concentration The concentrations of TSP (expressed in both per g FW and per g DW) decreased in all plants from D0 to D7 (Fig. 64). TSP concentration on D7 was relatively similar for plants exposed to 0, 50, 100 and 250 µM BPA. 4.3.7 Lipid Peroxidation Level (TBARs) TBARs (expressed as both per g FW and per g DW) were determined for both leaves (Fig. 65) and roots (Fig. 66). In leaves, TBARs increased in all plants from D0 137 (A) (B) Fig. 62. Effects of different BPA concentrations on total soluble sugar concentration in leaves during seven days of exposure. All concentrations are expressed in (A) mg glucose equivalent per g FW and (B) mg glucose equivalent per g DW. Each value is represented as the mean ± SE (n = 4). 138 Fig. 63. Scatterplots of total soluble sugar concentration (mg/ g FW) against chlorophyll fluorescence parameters, (A) Fv/Fm, (B) Fv/Fo, (C) Fv’/Fm’, (D) ФPSII, (E) qp and (F) NPQ of leaves of S. aureus plants exposed to BPA for 7 days. 139 (A) (B) Fig. 64. Effects of different BPA concentrations on total soluble protein concentration in leaves during seven days of exposure. All concentrations are expressed in mg BSA equivalent (A) per g FW and (B) per g DW. Each value is represented as the mean ± SE (n = 4). 140 (A) (B) Fig. 65. Effects of different BPA concentrations on lipid peroxidation levels (TBARs) in leaves of S. aureus, expressed in (A) µM MDA/ g FW and (B) µM MDA/ g DW, during seven days of exposure. Each value is represented as the mean ± SE (n = 4). 141 (A) (B) Fig. 66. Effects of different BPA concentrations on lipid peroxidation levels (TBARs) in roots of S. aureus, expressed in (A) µM MDA/ g FW and (B) µM MDA/ g DW, during seven days of exposure. Each value is represented as the mean ± SE (n = 4). 142 to D4. Surprisingly, the increases in TBARs on D4 (compared to D0) in plants exposed to 100 and 250 µM BPA were not as large as those of plants exposed to 0 and 50 µM BPA. TBARs in leaves decreased for all plants on D7. No significant differences were observed in TBARs (expressed as both per g FW and per g DW) in roots over the 7 days of BPA exposure. TBARs values in the roots were much lower than those of leaves. 4.3.8 Leaf Antioxidant Enzymes Leaf Ascorbate Peroxidase (APX) Activity The specific activity of APX (U/ mg protein) showed a drastic increase in all plants on D1 compared to D0 (Fig. 67A) and this increase (1.5-fold) was the largest in plants exposed to 250 µM BPA. APX specific activity for all the plants (including plants exposed to 0 µM BPA) started to decrease from D4 to D7. APX specific activity of all plants on D7 was slightly lower compared to those on D0. When expressed in per g FW (Fig. 67B), APX activity showed the same changes as those expressed in per mg protein. Leaf Glutathione Reductase (GR) Activity The changes in GR specific activity (U/ mg protein) were very similar to those of APX. GR specific activity drastically increased in all plants on D1 compared to those on D0 and this increase (94%) was the largest in plants exposed to 100 µM BPA (Fig. 68A). GR specific activity decreased in plants (including 0 µM BPA) from D4 to D7. Unlike APX, GR specific activity in all plants on D7 was higher than that of D0. 143 (A) (B) Fig. 67. Effects of different BPA concentrations on the activity of leaf ascorbate peroxidase, expressed in (A) U/ mg protein and (B) U/ g FW, during seven days of exposure. Each value is represented as the mean ± SE (n = 4). 144 (A) (B) Fig. 68. Effects of different BPA concentrations on the activity of leaf glutathione reductase, expressed in (A) U/ mg protein and (B) U/ g FW, during seven days of exposure. Each value is represented as the mean ± SE (n = 4). 145 When expressed in per g FW (Fig. 68B), GR activity showed the same changes as those expressed in per mg protein. Leaf Superoxide Dismutase (SOD) Activity SOD specific activity (U/ mg protein) remained relatively constant until D4 (Fig. 69A). On D4, it started to increase in all plants (including those at 0 µM BPA), and the largest increase (40% as compared to that on D0) was observed in plants exposed to 250 µM BPA. On D7, SOD specific activity for plants exposed to 0 and 100 µM BPA continued to increase but it decreased in plants exposed to 50 and 250 µM BPA. When expressed in per g FW (Fig. 69B), SOD activity showed the same changes as those expressed in per mg protein. Leaf Guaiacol Peroxidase (POD) Activity Plants exposed to 0 and 100 µM BPA exhibited a decrease in POD specific activity from D0 to D1 but it increased on D4 up to D7 (Fig. 70A). POD specific activity for plants exposed to 50 µM BPA decreased from D0 to D7. POD specific activity of plants exposed to 250 µM BPA decreased from D0 to D4, but it started to increase on D7. When expressed in per g FW (Fig. 70B), POD activity showed the same changes as those expressed in per mg protein. 146 (A) (B) Fig. 69. Effects of different BPA concentrations on the activity of leaf superoxide dismutase, expressed in (A) U/ mg protein and (B) U/ g FW, during seven days of exposure. Each value is represented as the mean ± SE (n = 4). 147 (A) (B) Fig. 70. Effects of different BPA concentrations on the activity of leaf guaiacol peroxidase, expressed in (A) U/ mg protein and (B) U/ g FW, during seven days of exposure. Each value is represented as the mean ± SE (n = 4). 148 Chapter 5. Discussion 5.1 Preliminary Screening of Plants Five different plant species were screened, namely Echinodorus palaefolius, Ipomoea aquatica, Cryptocoryne ciliata, Hydrocotyl sibthorpioides and Scindapsus aureus, to determine if they were able to phytoremediate high concentrations of NO3under partially-submerged conditions. E. palaefolius, C. ciliata and H. sibthorpioides are semi-aquatic macrophytes sold in aquarium stores as ornamental plants. These plants are known to be able to withstand partial-submergence and if used for in situ phytoremediation, they could also be used for landscaping. I. aquatica is also semiaquatic, but is a commonly consumed leafy vegetable, hence lacking ornamental value. However, this plant is extremely fast growing, and high growth rate is considered an indicator of efficient phytoremediating plant species (Brix and Schierup, 1989; Forni et al., 2001). S. aureus is the only terrestrial plant among the five species, but it is able tolerate partial submergence in water (Kamel et al., 2007). The reasons of choice for these five plant species are summarized in Table 13. Data obtained from the preliminary screen showed that S. aureus was the best plant to use for in-depth study of its potential for NO3- phytoremediation. E. palaefolius and I. aquatica plants were unable to adapt to the set-up conditions during the acclimatization phase. C. cilliata plants were unable to remove NO3- from water spiked with high NO3- concentrations and C. cilliata as well as H. sibthorpioides plants showed signs of senescence (yellowing) and necrosis when exposed to high NO3- concentrations for four weeks. Only S. aureus plants exhibited removal of NO3coupled with increases in plant fresh weight (FW) over the period of screening. 149 Table 13. Summary of characteristics of five different plant species that make them potential candidates for use in the phytoremediation of nitrate from water. Plant Species Semi-Aquatic E. palaefolius  I. aquatica  C. ciliata  H. sibthorpioides  S. aureus Terrestrial Fast Growing Ornamental Low Cost            150 5.2. Phytoremediation of Nitrates by S. aureus 5.2.1. Removal of Nitrate by S. aureus The data obtained further strengthened the conclusion that S. aureus was able to remove NO3- from water. By the end of W8, more than 80 % of NO3- (even from 200 mg/L-containing water) were removed by S. aureus under partially submerged conditions. Although results showed that algae from the control setups were also able to clear NO3- from water by W8 with similar efficiency, the presence of S. aureus allowed the rate of NO3- removal to be faster, particularly in water containing 20, 50 and 100 mg/L NO3- ( as observed in Table 4). It has been reported that NO3- concentrations can exceed 25 mg/L in surface waters and 100 mg/L in groundwater (Camargo et al., 2005). Dodds et al. (1998) also suggested the upper limits of total nitrogen for eutrophic temperate lakes to be 1.26 mg TN/L (5.58 mg NO3-/L) and for eutrophic temperate streams to be 1.50 mg TN/L (6.65 mg NO3-/L). All these concentrations were within the NO3- limits of this study, meaning that S. aureus plants could be utilized for the removal of NO3- and the prevention of eutrophication in these freshwater bodies. Results obtained on algal growth indicated that the presence of S. aureus helped to reduce algal concentrations, even in water containing 200 mg/L NO3-. The presence of S. aureus plants in the set-up containing 200 mg/L NO3- resulted in the reduction of chlorophyll a (chl a) and pheophytin a concentrations to levels comparable to those of control set-ups (with algal growth) containing 50 and 100 mg/L NO3-. The S. aureus plants not only removed NO3- from the water, hence, limiting nutrient availability for algal proliferation, but the leaves of the plants might 151 have an additional shading effect, reducing light available for algal photosynthesis. The combined effect resulted in the severe reduction in algal growth in set-ups containing S. aureus plants. Eutrophication of water bodies is commonly accompanied by harmful algal blooms (HAB), which consist of potentially toxic species and high biomass producers (Heisler et al., 2008). HAB can not only poison, but also cause hypoxic and anoxic conditions in the water, damaging the water ecosystem (Heisler et al., 2008). Dodds et al. (1998) suggested the upper limits of chl a concentration of planktons in eutrophic temperate lakes to be 25 µg/L and in eutrophic temperate streams to be 30 µg/L. In this study, the water in the set-ups was stagnant, which could account for the much higher chl a concentration (in mg/L range), as compared to lakes and streams with moving water. Yet, the presence of S. aureus plants reduced the high amount of algal growth (based on chl a and pheophytin a concentrations) by 50% or more. The longterm use of S. aureus plants for NO3- phytoremediation might lower algal growth alongside NO3- removal and therefore, can result in preventing eutrophication. The values of the conductivity of water provided data about the concentration of dissolved salts and high conductivity levels are indicators of eutrophic waters (Bellos and Sawidis, 2005). S. aureus was able to reduce or maintain the conductivity of the water containing 20, 50, 100 and 200 mg/L NO3- by the end of W8 to levels below or close to their corresponding levels at the beginning of the study at W0. Such reductions in conductivity readings corresponded with the reduction of NO3- levels in the water. However, the conductivity readings of the equivalent algal controls on W8 generally increased to levels much higher than that of W0. These changes indicated 152 that the algal growth in the water might have released high levels of electrolytes, resulting in the elevated levels of water conductivity. The growth of S. aureus plants in the experimental set-ups containing 0 – 200 mg/L NO3- resulted in increased pH of the water from W0 to W5, but pH fell to acidic levels from W5 to W8. The initial increase in pH observed was probably due to NO3removal as NO3- uptake is a proton-consuming process and usually causes alkalinisation of the surrounding medium (Goodchild and Givan, 1990; Marschner, 1995; Brix et al., 2002). The subsequent decrease in pH might be due to phosphate deficiency experienced by the plants. The growth medium supplied contained only 0.01 mM PO43-, which could be insufficient to sustain the increased growth of S. aureus from W5 onwards. Phosphate-deficient plants could lead to the acidification of the rhizosphere (and surrounding medium) when grown with NO3- and this acidification might arise from the protons released by the dissociation of organic acids synthesized by enhanced phosphoenolpyruvate carboxylase (PEP carboxylase) activity (Pilbeam and Kirkby, 1992). In the experimental algal controls, however, pH levels of water containing 0 and 20 mg/L NO3- were relatively neutral, but it became alkaline in water containing 50, 100 and 200 mg/L NO3- by the end of W8. Sooknah and Wilkie (2004) also noted a similar increase in pH when algae were allowed to grow in wastewater for 31 days. The alkalinisation observed on W8 was probably caused by algal uptake of NO3-. 5.2.2. Growth Parameters Plant FW and DW increased in all plants over the 8 weeks of study, and the increase was larger in plants exposed to 100 and 200 mg/L NO3-. The inorganic ion, 153 NO3-, provides nitrogen (N), which is an essential nutrient for plant metabolism and development, and high levels of NO3- in the growth medium (i.e. NO3- fertilization) could lead to increased levels of amino acids and proteins, and increased growth of plants (Stitt, 1999; Chen et al., 2004). However, increases in FW and DW in plants not supplied with NO3- (0 mg/L) indicated that S. aureus was still able to grow despite NO3- limitations, albeit the growth was not as great compared to plants exposed to NO3-. From W0 to W8, FW: DW ratios (an indication of water relations) decreased in plants exposed to 0, 20 and 50 mg/L NO3- but increased in plants exposed to 100 and 200 mg/L NO3-. Several plant adaptations to drought such as decreased succulence (% water) are the same as or similar to responses of N-deficiency (Radin and Parker, 1979a). Radin and Parker (1979a) reported that in Gossypium hirsutum, low-N leaves were more resistance to dehydration compared to high-N plants, with increased stomatal sensitivity to water stress. This resulted in low N-leaves having higher relative water content compared to high N-leaves. However, the reverse was observed in the S. aureus plants of this study. This indicated that other factors might be responsible for the changes in water relations of the S. aureus plants in this study. Ottosson and Welander (1996) observed that transpiration rates in Begonia X hiemalis cuttings increased after the formation of roots. Bal-Tal et al. (1994) reported that root pruning of Lycopersicon esculentum resulted in the reduction of transpiration rates. These studies suggested that changes in transpiration rates and therefore, plant water status, might be related to rooting. In the S. aureus plants, higher root growth was observed in plants exposed to low concentrations of NO3-, as compared to high concentrations. The increased root growth might have resulted in increased water 154 uptake, but also increased loss of water through transpiration. These water losses might possibly be higher than those lost through bigger leaf areas (observed in plants exposed to high NO3- concentrations), resulting in lower FW: DW ratio. The root: shoot ratio on W8 was highest in plants exposed to 0 mg/L NO3-, but lowest in plants exposed to 200 mg/L NO3-. Nitrogen deficiency causes increases in root: shoot ratios as plants promote root growth at the expense of shoot growth, in order to explore the medium for more nutrients (Améziane et al., 1997; Touraine et al., 2001; Hermans et al., 2006). This would explain the large increase observed in plants exposed to 0 mg/L NO3-. In waters containing 200 mg/L NO3-, plants were supplied with abundant NO3- concentrations, from which NO3- was not fully removed by W8. Fewer resources would then be channeled to root growth in these plants, resulting in the lowest root: shoot ratio. Leaf growth was also affected by exposure to NO3-. From the data obtained, exposure to high concentrations of NO3- promoted an increase in total number of leaves as well as total leaf area. Similar observations were made in Coffea arabica plants exposed to high N concentrations (Nunes et al., 1993). Also, the DW per cm2 total leaf area increased in plants exposed to low NO3- concentrations throughout the 8 weeks of study, but decreased in plants exposed to high NO3- concentrations. This indicated that the leaves of plants exposed to low NO3- concentrations were thicker than those exposed to high NO3- concentrations. These changes were also observed when additional nitrogen (N) was supplied to Solanum tuberosum (Vos and van der Putten, 1998) and Zea mays (Vos et al., 2005). High NO3- concentrations not only increased the number of leaves produced, but also increased the individual leaf size/area. Radin and Parker (1979a, 1979b) reported that in G. hirsutum, low N levels 155 can decrease root hydraulic conductance and reduce water delivery to growing leaves, thereby reducing leaf cell expansion and total leaf area. Abundant N (or NO3-) supply can increase the number of meristems produced by plants and encourage formation of shoots (Lawlor et al., 2001). A large N supply could also increase the number of cells per leaf and their size through cell expansion (Lawlor et al., 2001). Leaf area is also dependant on plant growth conditions. Lawlor et al. (1988; 2001) reported that in Triticum aestivum, increasing N supply increased the area of individual laminae, and the maximum size was dependent on environmental conditions such as water and temperature. Ismail and Noor (1996) also reported that soil flooding resulted in decreased leaf area in Averrhoa carambola. In the present study, partial submergence of S. aureus plants might have resulted in a smaller maximum leaf laminar size, but on W8, the area of each leaf of plants exposed to 100 and 200 mg/L NO3- was still larger compared to those of plants exposed to 0, 20 and 50 mg/L NO3-. 5.2.3. Photosynthetic Capacity Photosynthetic capacity is extremely dependent on plant N status in the long run because nearly 75% of the N in leaves of higher plants are stored in the photosynthetic machinery mainly as photosystems and carboxylases (Ferrario et al., 2001). Also, 20 – 30% of total leaf N (Sage et al., 1987), and 50 – 70% of total leaf proteins (Lawlor et al., 1987) are made up of ribulose-1,5-bisphosphate carboxylaseoxygenase (Rubisco). Rubisco is the enzyme required in carbon fixation, and is usually rate-limiting for photosynthesis (Seemann et al., 1987). Decreasing N levels could lead to a reduction in chlorophyll concentrations and Rubisco concentrations. The combined effects could cause a decrease in photosynthesis, resulting in a reduction in growth (Seemann et al., 1987). In this study, changes in the 156 concentrations of photosynthetic pigments and chlorophyll fluorescence parameters of S. aureus were affected by NO3- concentrations. In plants not exposed to NO3- (0 mg/L), concentrations of chlorophylls (chl a, chl b and total chl) and carotenoids decreased from W0 to W8. Plants exposed to 20 and 50 mg/L NO3- showed increased in chlorophylls and carotenoid concentrations from W0 to W4. Beyond W4, when NO3- was completely removed from the water, levels of chlorophylls and carotenoids started to decrease until W8. In plants exposed to 100 and 200 mg/L NO3-, however, levels of chlorophylls and carotenoids increased from W0 to W8. The relationship between chlorophyll concentration and N levels has been extremely well-documented over the years, so much so that recent studies report the use of chlorophyll content as a gauge for plant N content (Hunt et al., 1985; Khamis et al., 1990; Chikaraishi et al., 2005; Demotes-Mainard et al., 2008). Chlorophyll molecules contain N (Sechley et al., 1992) and chlorophyll concentrations are closely related to N content within leaves because photosynthetic proteins account for more than half of total N within the leaf (Demotes-Mainard et al., 2008). N limitations can result in decreases in chlorophyll concentrations by downregulating synthesis of antenna chlorophyll protein complexes (Skillman and Osmond, 1998). Under NO3- deficiency, as observed in S. aureus exposed to 0 mg/L NO3- from W0 to W8, as well as 20 and 50 mg/L NO3- from W4 onwards, these plants were unable to produce sufficient chlorophylls to replace those depleted or destroyed during chlorophyll turnover. Sønsteby et al. (2009) observed that chlorophyll concentrations in Fragaria x ananassa plants increased with NO3- fertilization, but decreased when fertilization was terminated. Khamis and Lamaze (1990) also 157 observed a decrease in chlorophyll concentration with decreasing NO3- levels in Zea mays. N-deficient Olea europaea plants were observed to have lower chlorophyll a concentrations in the leaves (Boussadia et al., 2010). Heuchera American plants grown in low N-conditions had relatively lower chlorophyll concentrations as compared to those grown in higher N-conditions (Skillman and Osmond, 1998). Also, as a response to limiting N due to NO3- deficiency, plants tend to promote root growth at the expense of shoot growth, and existing N resources in the leaves are channelled to the roots, resulting in reduction in concentrations of chlorophylls and other photosynthetic proteins, including Rubisco (Améziane et al., 1997; Touraine et al., 2001; Hermans et al., 2006). However, as observed in S. aureus plants exposed to 100 and 200 mg/L NO3-, when N levels were abundant, the concentrations of chlorophylls were higher and plants appeared ‘greener’ in colour (Hunt et al., 1985). In this study, TSP concentrations in the leaves of S. aureus plants were generally higher in plants exposed to high NO3- concentrations than low NO3concentrations. Since Rubisco makes up 50 – 70% of total leaf proteins (Lawlor et al., 1987), higher TSP levels also indicated higher Rubisco concentrations. In Phaseolus vulgaris and Alocasia macrorrhiza, the leaf concentration of Rubisco was linearly dependent on leaf N content (Seemann et al., 1987). Pérez et al. (2005) also reported increased levels of Rubisco in Triticum aestivum when supplied with increased N. The increased chlorophyll and Rubisco concentrations in S. aureus plants exposed to high NO3- concentrations could result in elevated photosynthetic rates. This increase in photosynthesis could then account for the greater increase in biomass 158 observed in plants exposed to high NO3- concentrations as compared to those exposed to low NO3- concentrations. Carotenoid concentrations are also dependent on the N supply in plants, as observed in this study. Ramalho et al. (2000) noted that the increase in carotenoid concentrations when Coffea arabica plants were treated with high NO3- content. Smoleń and Sady (2009) also reported that carotenoid concentrations in Daucus carota increased with NO3- fertilization and foliar nutrition. In Neoregelia cruenta, carotenoid levels were lower in plants experiencing limiting N, compared to plants exposed to high N conditions (Fernandes et al., 2002). Various chlorophyll fluorescence parameters obtained suggested that the efficiency of PSII was dependent upon the NO3- concentrations provided to S.aureus plants. The Fv/Fm ratio is defined as the maximum quantum efficiency of PSII (photosystem II) in using light absorbed for the reduction of the primary quinone acceptor (QA) (Baker, 2008). Fv/Fm decreased slightly from W0 to W8 in plants exposed to all NO3- concentrations. On the other hand, the decreases from W0 and W8 for Fv/Fo were comparatively larger. Both Fv/Fm and Fv/Fo values are indicators of photosynthetic efficiency of PSII, but Fv/Fo appeared to be a more sensitive indicator. In Zea mays (Khamis et al., 1990; Lu and Zhang, 2000), Spinacia oleracea (Verhoeven et al., 1997), Coffea arabica (Nunes et al., 1993) and Heuchera americana (Skillman and Osmond, 1998), Fv/Fm decreased with low N-levels and increased when provided with high-N levels. In this study, however, Fv/Fm values decreased for plants exposed to both low (0, 20, 50 mg/L) and high (100 and 200 mg/L) NO3- levels. Yet upon closer observation, Fv/Fo ratios in plants exposed to 100 and 200 mg/L NO3- actually increased from W0 to W6 and W4 respectively, before 159 decreasing to W8. This indicated that exposure of S.aureus plants to high NO3- levels increased the maximum quantum efficiency of PSII and Fv/Fo only decreased towards the end of the experiment when NO3- was reduced significantly by S. aureus. The maximum efficiency of PSII, Fv’/ Fm’, is an indication of the efficiency of light capture by ‘open’ PSII reaction centers (oxidized QA) (Baker, 2008). Similar to Fv/Fm and Fv/Fo, as NO3- levels depleted towards W8, Fv’/Fm’ also decreased on W8, compared to W0, in all plants. However, in plants exposed to 200 mg/L NO3-, Fv’/Fm’ increased from W0 to W2 before decreasing from W2 to W8. Also, the decrease from W0 to W8 in Fv’/Fm’ was smallest in plants exposed to 100 mg/L NO3-. Both observations suggested that high concentrations of NO3- used did not constitute a stress to decrease the efficiency of light capture by oxidized QA when the plants were exposed to NO3- earlier in the experiment. Only towards the end of the study did the decreases in Fv’/Fm’ become larger. Similar changes in Fv’/Fm’ with N-levels were observed in Z. mays (Lu and Zhang, 2000), S. oleracea (Verhoeven et al., 1997) and C. arabica (Nunes et al., 1993), as Fv’/Fm’ values were high in plants supplied with high N-levels but much lower when the plants were supplied with low N-levels. The PSII operating efficiency, ФPSII, gives an estimate of the efficiency by which light captured by PSII is used for the reduction of QA and is also an estimate of the quantum yield of the linear electron flux through PSII (Baker, 2008). The value of qp (photochemical quenching) gives an indication of the photochemical capacity of PSII at the steady state and is also reflective of the proportion of oxidized QA (Baker, 2008). Both ФPSII and qP decreased from W0 to W8 in all plants. This showed that a decrease in NO3- concentration could lead to a drop in efficiency of the PSII operating system. Similar observations were made in Z. mays (Lu and Zhang, 2000), S. oleracea 160 (Verhoeven et al., 1997) and C. arabica (Nunes et al., 1993), with plants exposed to low N-levels exhibiting low ФPSII and qP values. In plants exposed to 100 mg/L NO3-, both ФPSII and qP values remained relatively constant from W0 to W2, but decreased from W4 onwards. In plants exposed to 200 mg/L NO3-, however, both ФPSII and qP values decreased sharply from W0 to W2, then increased sharply again from W2 to W4, before decreasing again. This suggested that initial exposure to extremely high concentrations of NO3- negatively affected ФPSII and qP, but the plants slowly acclimatized to the high concentrations by W4, leading to a sharp increase in PSII operating efficiency. This acclimatization period was not necessary for plants exposed to 100 mg/L NO3-, indicating that this concentration was optimum for maintenance of PSII operating efficiency. NPQ (non photochemical quenching), the non-radiative energy dissipation via the xanthophyll cycle, increased sharply for plants exposed to 0 and 20 mg/L NO3from W0 to W2, and also the largest increases in NPQ were from W0 to W8 compared to the other NO3- concentrations. In Z. mays, NPQ values were much higher in N-deficient plants compared to plants grown with adequate N (Khamis et al., 1990; Lu and Zhang, 2000). In S. oleracea (Verhoeven et al., 1997) and C. arabica (Nunes et al., 1993), exposure of the plants to low N-supply also resulted in higher non photochemical quenching values (NPQ and qNP) as compared to plants exposed to high N-supply. Limiting N caused the interconversion of violaxanthin to zeaxanthin (Demmig-Adams and Adams, 1992; Verhoeven et al., 1997), which led to increased NPQ (Skillman and Osmond, 1998). This explanation could also account for the increase in NPQ levels for plants exposed to 50, 100 and 200 mg/L NO3- towards the end of the study. These plants, however, experienced a decrease in NPQ levels from 161 W0 to W2, with the percentage decrease becoming larger with increasing NO3concentrations. Although PSII efficiency decreased in plants exposed to high NO3concentrations from W0 to W8 (as indicated by decreasing ФPSII values), this was compensated by the increase in chlorophyll concentrations over the same period of time. Therefore, overall photosynthetic capacity in these plants might not have been drastically affected and this accounted for the large increase in growth (increase in FW, DW and leaf area) throughout the 8 weeks of study. 5.2.4. Carbon (C) and Nitrogen (N) metabolism Results show that in general, TN in S. aureus plants increased with increasing NO3- concentration. When NO3- was completely phytoremediated by S. aureus towards the end of the study for water containing 20, 50 and 100 mg/L NO3-, TN decreased accordingly. These results indicated that S. aureus could take up and assimilate NO3- within its tissues and that TN was a good reflection of the amount of NO3- absorbed into the plant tissues. Similar changes in TN were observed in Zea mays exposed to different concentrations of NO3- (Khamis and Lamaze, 1990). Nunes et al. (1993) also reported that Coffea arabica plants exposed to high N-levels had higher TN as compared to plants exposed to low N-levels. In Daucus carota plants, increased NO3- fertilization resulted in increased TN in the roots (Smolén and Sady, 2009). N-deficient Olea europaea plants were also observed to have lower TN in the leaves (Boussadia et al., 2010). The total root nitrogen: total shoot nitrogen ratio (TNR: TNS) was highest in plants exposed to low NO3- concentrations compared to high NO3- concentrations. 162 Under limiting N conditions, when the plants were exposed to low NO3concentrations over a prolonged period of time, plants tend to allocate N resources to roots at the expense of shoots to allow for increased root growth to search for exogenous N sources (Améziane et al., 1997; Touraine et al., 2001; Hermans et al., 2006). This would therefore result in the higher TNR: TNS in plants exposed to low NO3- concentrations. In plants exposed to 200 mg/L NO3-, both TN and TNR: TNS were extremely low on W2 compared to plants exposed to other NO3- concentrations. Plant NO3uptake could be inhibited if the plant was supplied with high concentrations of NO3-, particularly if the uptake matched the growth rate and the storage pools were saturated (Tischner and Kaiser, 2007.) Two hundred mg/L NO3- might be too high a concentration for S. aureus and resulted in slight inhibition of NO3- uptake and a subsequent decrease in TN. However, when the levels of NO3- decreased on W4 due to phytoremediation, the substrate inhibition was removed and the TN and TNR: TNS in these plants increased drastically. The concentrations of TSS decreased from W0 to W8 in all plants, but this decrease was smallest in plants exposed to 100 and 200 mg/L NO3-. This coincided with the decreases in concentrations of chlorophylls and PSII capacity (as determined from chlorophyll fluorescence parameters) from W0 to W8, particularly in plants exposed to 0, 20 and 50 mg/L NO3-. This indicated that the decrease in TSS levels was due to a decrease in photosynthetic capacity. Towards the end of the study (W6 to W8), plants exposed to 100 and 200 mg/L NO3- exhibited high TSS levels (expressed in per g DW). Compared to S. aureus plants exposed to low NO3- conditions, photosynthetic capacity in plants exposed to high NO3- concentration was 163 comparatively higher, resulting in the higher TSS concentrations in those plants. Also, increased NO3- in plant tissues could shift carbohydrate metabolism from starch synthesis to increased sugar synthesis (Tischner and Kaiser, 2007), which was possibly one reason for the higher TSS concentrations in these particular plants. Similar differences in TSS levels were observed in Typha angustifolia plants grown under different trophic conditions. Steinbachová-Vojtíšková et al. (2006) reported that T. angustifolia plants grown under hypertropic conditions (high N- levels) had the highest levels of TSS (sucrose, glucose and fructose) compared to plants grown under eutrophic condititions (moderate N-levels), but plants grown under oligotrophic conditions (low N-levels) had the lowest concentrations of TSS. The concentrations of TSP in old existing leaves in plants (OL) slightly decreased from W0 to W8 in plants exposed to 0, 20 and 50 mg/L NO3-, as the NO3levels in waters were phytoremediated towards the end of the study. In plants exposed to 100 and 200 mg/L NO3-, however, TSP concentrations increased about 2-fold on W8 compared to W0. A similar trend was observed in the newly formed leaves (NL) after NO3- addition. NO3- assimilated into the plant tissues is converted into NH4+ and subsequently used for protein synthesis. In Zea mays, TSP levels were observed to increase with increasing NO3-, but as NO3- concentrations decreased towards the end of the experiment, TSP levels also decreased (Khamis and Lamaze, 1990). Lawlor et al. (1987) showed that in wheat supplied with sufficient nitrogen fertilizer, Rubisco made up 50 – 70% of total leaf TSP levels. Rubisco acts as a store of N when excess N is available than required for growth (Warren et al., 2000), which could account for the drastic increase in TSP levels in for plants exposed to high NO3- concentrations. Increases in total N led to increases in Rubisco N in Triticum aestivum (Evans, 1983), 164 Chenopodium album and Amaranthis retroflexus (Sage et al., 1987), Phaseolus vulgaris and Alocasia macrorrhiza (Seemann et al., 1987) and Spinacia oleracea (Terashima and Evans, 1988) and these increases were consistant with the putative role of Rubisco as a storage protein (Warren et al., 2000). High levels of Rubisco can also help compensate for the low rate of enzyme catalysis and low affinity of CO2, leading to increased photosynthesis (Warren et al., 2000). Also, high levels of N have also been reported to decrease the degradation of proteins in the reaction center of PSII (Kolber and Falkowski, 1988; Cai et al., 2008). These reasons could account for the high TSS levels in S. aureus plants exposed to 100 and 200 mg/L NO3-, as a result of increased photosynthesis. The NR activity in all OL and NL showed a general increase from W0 to W8, but this increase was larger in plants exposed to lower NO3- concentrations than higher concentrations. Similarly, root NR activity also increased from W0 to W8, and NR activity was higher in plants exposed to low NO3- concentrations, compared to high NO3- concentrations. NR mRNA expression is induced by NO3- uptake (Faure et al., 2001; Meyer and Stitt, 2001; Tischner and Kaiser, 2007), but levels of NR activity do not necessarily correlate with NO3- levels in the plant tissues (Sechley et al., 1992) or with exogenous NO3- concentration (Chen at al., 2004). Ivanshikina and Sokolov (1997) reported that NR activity increased with increasing NO3- within Zea mays tissues whereas Chen et al. (2004) observed that in Brassica chinensis, Brassica campestris and Spinach oleracea, NR activity was only induced at low levels of NO3- but beyond a certain concentration, NR activity did not increase further. In the S. aureus plants, NR activities might be influenced by other factors aside from NO3- uptake. NR is 165 sensitive to several environmental factors and its activity can differ under the effects of light intensity, CO2 levels, temperature, water availability and NO3- supply (Shaner and Boyer, 1976). For instance, in Betula pendula plants, light intensity can influence transcription of the NR gene, and NR mRNA levels increased when the plants were transferred from dark to light conditions (Strater and Hachtel, 2000). NR activity in Gracilaria chilensis fluctuated when the plants were placed under a light:dark cycle, with NR activity increasing by more than 2-fold in the light phase (Chow et al., 2004). Anoxia can also result in increased NR activity, as observed in the roots of Lycopersicon esculentum grown under anoxic conditions (Morard et al., 2004). Agüera et al. (2006) reported that in Cucumis sativus plants, NR expression decreased in very low CO2, but at normal and elevated CO2 conditions, both NR expression and activity increased, even though NO3- levels supplied to the plants were low. The effects of temperature on NR activity have also been studied. Calatayud et al., (2008) observed that exposure to low temperatures could induce higher NR activity in Rosa hybrids, leading to higher NO3- uptake. The influence of NO3- supply on NR activity is particularly prominent only when other environmental factors remain constant (Beevers and Hageman, 1969; Shaner and Boyer, 1976). As this present study was conducted in a greenhouse environment, light intensity and the surrounding temperature might be high but fluctuating. Also, the S. aureus plants were partially submerged in water, resulting in a waterlogged environment, of which pH changed with time (as indicated by results above) and conditions of hypoxia or anoxia might occur in the rhizosphere. All these environmental factors might have influenced the induction of NR activity, resulting in the higher leaf NR activities in plants exposed to lower NO3- concentrations. 166 The activity of NR in the roots in general was lower than that detected in the leaves, particularly in OL. Samuelson et al. (1995) observed that NR activity in the roots of Hordeum vulgare plants were unaffected by exposure to increased NO3concentrations, but the increased NO3- levels led to increased NR activity in shoots. In most herbaceous plants, assimilation of NO3- mainly takes place in the leaves (Faure et al., 2001). The roots take up NO3- from the water, but most of the NO3- was channelled to the leaves for N-assimilation, resulting in higher NR activity in the leaves than roots of the S. aureus plants. Also, shoot NO3- reduction (leaves included) might regulate NO3- uptake and, hence, NR activity, in the roots via phloemtranslocated amino acids (Muller and Touraine, 1992; Ivanshikina and Sokolov, 1997). Although root NR activity (µM NO2-/ g FW/ hr) only showed significant differences from W0 to W8 in plants exposed to 50 and 100 mg/L NO3-, a closer observation of NR activity expressed on a per mg protein basis showed that root NR activity increased significantly from W0 to W8 in all plants. This indicated that the increase in root NR activity detected was not due to increase in protein concentrations. Morard et al. (2004) reported that when the roots of L. esculentum underwent anoxia, NR activity increased. A similar situation might have occurred in this study as prolonged growth under partially submerged conditions might cause anoxic conditions, resulting in increased NR activity in the roots of S. aureus plants. 5.2.5. Lipid Peroxidation and Antioxidant Enzyme Activities On W2, lipid peroxidation levels in plants exposed to 0 (expressed in per g DW), 50 and 200 mg/L NO3- showed huge increases as compared to W0. The increases in TBARs in these plants coincided with large decreases in ФPSII and qP values. Plants exposed to 0 mg/L NO3- might have exhibited this increase due to the 167 plants experiencing oxidative stress due to NO3- and hence N deficiency. Tewari et al. (2004) reported similar increases in leaf lipid peroxidation levels of Zea mays plants during N deficiency. Lipid peroxidation levels in S. aureus plants exposed to 0 mg/L NO3- first decreased, then remained relatively constant as the study progressed, which might be an indication of acclimatization to the stressor. In plants exposed to 200 mg/L NO3-, the initial NO3- concentrations might be too high for the plants to tolerate, resulting in oxidative stress for the plants on W2, similar to plants experiencing salinity stress (Parida and Das, 2005). High salinity levels are known to cause osmotic stress in plants, leading to increases in ROS production (Azevedo Neto et al., 2006) which can cause lipid peroxidation. In Solanum melongena seedlings exposed to an excess of calcium nitrate, lipid peroxidation levels in the leaves increased due to salinity stress (Wei et al., 2009). As the high NO3- levels decreased towards the end of this study due to phytoremediation, lipid peroxidation levels decreased accordingly. Since plants exposed to 0, 50 and 200 mg/L NO3- showed decreases in PSII capacity (as determined from ФPSII and qP values) as well as increased TBARs on W2, the higher lipid peroxidation levels could be due to excess light energy absorbed by the PSII system but was not fully transmitted for photosynthesis. This excess light energy was then diverted to oxygen, and it might have resulted in the formation of reactive oxygen species (ROS) and increased lipid peroxidation levels. Lipid peroxidation levels in plants exposed to 50 mg/L NO3- decreased on W4 compared to W0, but remained high on W6 (especially when TBARs were expressed in per g DW basis). The high lipid peroxidation levels were most probably a result of N deficiency in the plants as on W4, NO3- levels remaining were less than 35% of the 168 original concentration, whereas by W6, NO3- levels in the water set-up decreased to less than 2% of the original concentration. Lipid peroxidation levels in plants exposed to 100 mg/L NO3- decreased from W0 to W2 and levels remained low until W4. However, TBARs showed a large increase on W6. A concentration of 100 mg/L appeared to be the optimal NO3concentration for S. aureus as membrane integrity appeared to be the least disrupted as compared to the other NO3- concentrations from W0 to W4. However, by W6, NO3- levels in the water set-up decreased to less than 10% of the original concentration, resulting in the plants experiencing N deficiency and an increased oxidative stress in the leaves. Plants exposed to 20 mg/L NO3- also showed a sharp increase in lipid peroxidation levels on W4, which was most probably a result of oxidative stress caused by N deficiency as NO3- levels in the water set-up on W4 had decreased to less than 5% of the original concentration. In general, APX levels increased from W0 to W8 in all plants, with the largest increased observed in plants exposed to 0, 20 and 50 mg/L NO3-. APX levels have been reported to increase with N deficiency in wheat plants during grain filling (Cai et al., 2008). Similarly, the prolonged exposure to NO3- deficiency might have caused the large increase in APX activity in S. aureus plants. On W2, APX levels were high in plants exposed to 0, 50 and 200 mg/L NO3- and this coincided with the high TBARs levels also observed on W2. APX acts specifically to scavenge for H2O2 (Asada, 1992) and H2O2 is a ROS, which can result in lipid peroxidation. The high lipid peroxidation levels might have led to the induction of APX activity in these plants on W2. Exposure to high NO3- levels might also account for the large increase 169 in APX activity in plants exposed to 200 mg/L NO3- for 2 weeks. Wei et al. (2009) reported increased APX levels in Solanum melongena seedlings exposed to high concentrations of Ca(NO3)2. High NO3- levels can cause salinity stress in plants, leading to increases in ROS production and subsequent increases in antioxidant enzyme activities (Azevedo Neto et al., 2006). However, as NO3- was phytoremediated from W4 onwards in this study, APX activities in the S. aureus plants decreased accordingly. APX and GR are enzymes involved in the Halliwell-Asada pathway and act specifically to scavenge for H2O2 (Asada, 1992) and maintain a high GSH/GSSG ratio (Foyer and Noctor, 2005). An increase in APX activity is expected to be accompanied by an increase in GR activity. However, in this study, while APX activity increased from W0 to W8 in all the plants, GR activity decreased in almost all the plants. In Hordeum vulgare plants subjected to waterlogged conditions, leaf GR activity decreased while leaf APX activity increased (Yordanova et al., 2004). Biemelt et al. (1998) also made a similar observation in the roots of Triticum aestivum plants when they were exposed to hypoxic and anoxic conditions. Therefore, the decrease in GR observed in the present study could be due to hypoxia or anoxia experienced by the S. aureus plants when grown in waterlogged conditions for a prolonged period of time. Also, GR activities showed the largest decrease from W0 to W8 in plants exposed to 100 and 200 mg/L NO3-, which could possibly be a result of high NO3- concentrations promoting increased algal growth and greater hypoxic and anoxic water conditions (Heisler et al., 2008). Similar to APX, SOD activities increased in all plants from W0 to W8. Zea mays plants subjected to N-deficiency showed a 2-fold increase in SOD activities 170 compared to plants not experiencing N-deficiency (Tewari et al., 2004). The S. aureus plants in this study also showed a 2/3-fold increase in SOD activities from W0 to W8, most probably a result of prolonged NO3- deficiency. For plants exposed to 200 mg/L NO3-, however, NO3- levels in the water did not drop to exceedingly low levels by W8. Hence, the increase in SOD activity from W0 to W8 for this plant was not likely due to NO3- deficiency. This increase was also not likely due to prolonged exposure to waterlogged conditions because as reported in studies done on Vigna radiata (Ahmed et al., 2002) and H. vulgare (Yordanova et al., 2004), exposure to waterlogged conditions led to a decrease, rather than an increase, in SOD activity. POD activity in all plants decreased from W0 to W8 when expressed as U/ mg protein, but it only decreased in plants exposed to 0 and 200 mg/L NO3- when expressed as U/ g FW. This indicated that the activity of the enzyme decreased in all plants, but in plants exposed to 20, 50 and 100 mg/L NO3-, the decrease in POD activity was much smaller compared to plants exposed to 0 and 200 mg/L NO3-. Also, there was a possibility that the amount of enzymes produced per g FW in these plants were high enough to compensate for the decrease in activity. Clearly in plants exposed to 0, 20, 50 and 100 mg/L NO3-, the removal of NO3- from the water led to NO3- deficiency in these plants after 8 weeks and this deficiency resulted in a decrease in the activities of POD. NO3- deficiency also resulted in a decrease in TSP levels from W0 to W8, which might account for the decrease in enzyme levels in plants exposed to 0 mg/L NO3-. Very few studies have reported on the effect of NO3concentrations on POD activities, but POD activities have been observed to increase in plants exposed to high N-levels. Zhang et al. (2010) reported that in Potamogeton 171 crispus plants grown submerged in eutrophic waters, POD activity increased due to the high nutrient (nitrogen) levels. In plants exposed to 200 mg/L NO3-, however, there were no NO3- limitations on W8 and TSP levels increased from W0 to W8 in these plants, yet POD activities decreased drastically. The leaves were also not undergoing senescence as chlorophyll levels increased from W0 to W8. By W8, plants exposed to 200 mg/L NO3- were undergoing minimal stress as NO3- concentrations had fallen to levels that met S. aureus N-requirements. The increases in APX and SOD activities were sufficient to deal with existing ROS, which might have resulted in the down-regulation of POD activities. 5.2.6. The Effectiveness of S. aureus as a Phytoremediator of Nitrate Based on the results obtained from this study, S. aureus definitely had the potential to phytoremediate NO3- from water, even at high concentrations. However, the rates of NO3- removal might not be significantly higher than that of algal controls. In eutrophic waters, blooms of algae have been reported to cause hypoxic and anoxic conditions, as well as release toxic compounds, all of which could cause harm to the ecosystem (Heisler et al., 2008). This study showed that the use of S. aureus in the phytoremediation of NO3- could lead to drastic decreases in the algal population as compared to controls. Furthermore, physiological and biochemical data obtained showed that S. aureus was able to propagate efficiency even when exposed to 200 mg/L NO3-, and only when NO3- concentrations were depleted to levels lower than the plant’s Nrequirements (approximately 0 – 10 mg/L NO3-) , were the plants showing signs of 172 oxidative stress and decreases in photosynthetic capabilities. The loss of chlorophyll pigments (which eventually led to leaf yellowing), decreases in biomass and decreases in leaf area could all act as visible plant indicators of depleted NO3- levels from the water body that S. aureus was used for phytoremediation. In general, S. aureus plants would make excellent phytoremediators of NO3- from polluted water bodies. 5.3. Phytoremediation of Bisphenol A (BPA) by S. aureus 5.3.1. Removal of BPA by S. aureus Studies have shown that microorganisms such as bacteria and fungi are capable of degrading BPA (Kang et al., 2006b). In the present study, S. aureus plants, under aseptic conditions, could tolerate submergence and, at the same time, remove BPA from water. Experimental data obtained indicated that the depletion of BPA from the water samples was solely due to phytoremediation by S. aureus plants. The data obtained indicated that the phytoremediation rate of BPA by S. aureus plants was much faster than degradation via photolysis. In less than 7 days, S. aureus was able to remove almost all of the BPA present in 50 and 100 µM BPAcontaining water and more than 50% of BPA from 250 µM BPA-containing water. The HPLC profiles obtained from water analyses, as well as the colour of the water samples, indicated that during the phytoremediation of BPA, a secondary compound was present and its concentration increased with decreasing BPA levels. This secondary compound might also share some similar chemical characteristics with BPA, due to the close proximity of its peak to that of BPA. Imai et al. (2007) also reported the formation of an unknown product in the BPA-spiked waters containing 173 Portulaca oleracea plants. The concentration of this unknown product also increased with decreasing BPA concentration. Imai et al. (2007) speculated that this unknown product might be the first metabolite of BPA and suggested that it could be 4[1-(4hydroxypheyl)-1-methyl-ethyl]-benzen-1,2-diol, a hydroxylation product of BPA. Another study on freshwater algal metabolism of BPA also reported the presence of new minor peaks in the HPLC profiles of the micro-algal culture media alongside the decreasing peaks of BPA (Nakajima et al., 2007). Further analyses of this peak via FAB-MS and 1H-NMR identified it to be either BPA-mono-O-β-D-glucopyranoside or BPA-mono-O-β-D-galactopyranoside, depending on the species of algae incubated with BPA (Nakajima et al., 2007). Therefore, the secondary compound detected in the BPA-spiked water samples of the present study could possibly be a metabolite of BPA – the result of phytoremediation of BPA by S. aureus. The secondary compound detected in this study could also be a root exudate. Plants can respond to environmental pollutants by exuding compounds at the root tips in order to induce biofilm production for enhancing the breakdown of pollutant(s) in the rhizosphere (Harsh et al., 2006). Plants exposed to high levels of heavy metals have been reported to exude phenolic compounds at the root tips (Barceló and Poschenrieder, 2002; Quartacci et al., 2009). Since the retention time corresponding to this secondary compound was very close to that of BPA, it could be surmised that this compound from S. aureus could also be a phenolic compound. HPLC analysis of S. aureus plant extracts collected on D7 did not show any trace of BPA, indicating that BPA absorbed into the plant tissues would have been metabolized into a different compound. Several studies involving the use of whole plants for the phytoremediation of BPA showed similar results. Imai et al. (2007) 174 showed that after 24h incubation of Portulaca oleracea in water containing 50 µM BPA, no traces of BPA was found in the plant. Noureddin et al. (2004) showed that BPA was detected only in the roots of Ipomoea aquatica at a very low level when the plants were initially exposed to BPA (21.9 µM), but it quickly disappeared after 4 days of exposure. The data obtained with S. aureus indicated that it was a nonaccumulator of BPA and the BPA absorbed into the plant tissues appeared to be quickly metabolized. A number of studies on plant biodegradation of BPA have shown that BPA absorbed into the tissues can be metabolized in a number of ways. Glycosylation of BPA has been determined as the main mechanism of BPA metabolism in plants (Kang et al., 2006b). Nakajima et al. (2004) found out that metabolites of BPA glycosylation included BPA mono-β-D-glucopyranoside, BPA di-β-Dglucopyranoside, BPA mono-β-D-gentiobioside and BPA β-D-glucopyranosyl(1→4)-[β-D-glucopyranosyl-(1→6)]β-D-glucopyranoside. However, other mechanisms have also been reported. Oxidation and oxidative cleavage of BPA can also occur as a result of the activity of fungal laccase (Fukuda et al., 2001; Modaressi et al., 2005), and peroxidases (Caza et al., 1999; Sakuyama et al., 2003; Li and Nicell, 2008) and polyphenol oxidases (Yoshida et al., 2002). Also, because of the phenol group as part of its structure, hydroxylation of BPA via cytochrome P450, an enzyme well-known to be involved in the degradation of various phenolic xenobiotics could also occur (Harvey et al., 2002; Morant et al., 2003), although no evidence for this reaction has yet to be reported. In order to determine the exact mechanisms of BPA metabolism and to identify the BPA metabolites in S. aureus, further studies are required. This will help in determining whether S. aureus is a suitable plant for the 175 long term phytoremediation of BPA, annulling all endocrine-disrupting properties of BPA and its metabolites in water. 5.3.2. Growth Parameters Exposure of S. aureus to BPA led to similar decreases in FW and DW as those of the experimental controls (0 µM BPA). This was unexpected because within the study period of 7 days, new leaves unfolded and expanded, and new root growth was also detected in most of the plants. The decrease in FW was smaller than that of DW, suggesting that water relations of the plants were minimally affected (also indicated by high values of FW: DW ratios). Such decreases in plant biomass could be due to the depletion of plant reserves during the time when they were grown in BPA-spiked water, without an exogenous sugar supply under plant culture conditions. During this period, the photosynthetic capacity of the plants also decreased (as indicated by decreasing values of ФPSII), hence reducing the amount of carbon fixed by the plants. The increase in root: shoot ratios in all plants indicated that partial submergence in water favoured root growth over shoot growth. Higher BPA concentrations also promoted greater root proliferation as compared to lower BPA concentrations Root biomass appeared to be more affected by the culture conditions than leaf biomass. Also, roots of S. aureus exposed to high concentrations of BPA showed browning at the root tips. This was also observed in hydroponically-grown seedlings of Lactuca sativa, Lycopersicon esculentum, Triticum durum and Vicia faba, which also exhibited root browning symptoms when they were exposed to BPA for 21 days (Ferrara et al., 2006). Ferrara et al. (2006) also reported the formation of a jelly-like film in the rhizospheric region and attributed both symptoms to enhanced bacterial activity, as the seedlings of all four plant species were not grown under aseptic 176 conditions in their study. Browning of root tips was also observed in plants exposed to heavy metals like cadmium (Punz and Sieghardt, 1993) and copper (Kopittke and Menzies, 2006), and this could be the result of the deposition of phenolics or enhanced suberization at the root tips (Punz and Sieghardt, 1993; Ghanati et al., 2005) – a plant mechanism to cope with stress. Hence, the data obtained indicated that total plant biomass was not affected by BPA. However, the physiological/biochemical responses of S. aureus could be influenced by the exposure of the plants to BPA, in terms of BPA concentration and exposure duration. 5.3.3. Photosynthetic Capacity The change from a solid to a liquid growth medium also seemed to result in a sharp decrease in chlorophyll concentrations, especially from D0 to D1. Plants exposed to 0 µM BPA exhibited the largest drop in chlorophyll concentrations from D0 to D1, when compared to plants exposed to 50, 100 and 250 µM BPA. BPA present in the growth media could have buffered the plants from the stress of growth medium change, resulting in lesser changes in chlorophyll concentration. The subsequent increase in chlorophyll concentrations observed in plants exposed to 0 µM BPA (from D1 to D7) indicated that the plants were able to acclimatize to new growth conditions and the increase in chlorophyll concentrations were due to increasing plant age. However, exposure to BPA, particularly at high concentrations, was observed to cause reductions in levels of chl a, chl b and carotenoids from D0 to D4. On D7, recovery in chl a and b levels was observed in plants exposed to 50, 100 and 250 µM. Recovery rate of chl b was also observed to be lower than that of chl a. Two possible explanations could account for such changes in chl levels: (1) high BPA 177 concentrations led to increased degradation of chlorophylls or (2), BPA resulted in a decrease in chlorophyll synthesis. Carotenoid concentration was also affected by BPA exposure. Concentrations of carotenoids in plants exposed to BPA decreased from D0 to D4, but carotenoid levels showed signs of recovery on D7. Carotenoids are known to be important quenchers of the triplet state of chlorophyll (3Chl) and they also actively scavenge for singlet oxygen radicals (Demmig-Adams, 1990; Frankart et al., 2003). With decreasing carotenoid content, a buildup of reactive oxygen species (ROS) could occur in the leaves, leading to oxidative damage. The Fv/Fm ratio of S. aureus grown under the present culture conditions was at an average of 0.80 (D0). In most healthy plant species grown in natural conditions, the Fv/Fm ratio is maintained relatively constant within the range of 0.80 – 0.86 (Bjӧrkman and Demmig, 1987). The F v/Fm value for plants exposed to 250 µM BPA was about 0.78, indicating minimal effects of such high levels of BPA on the PSII efficiency of S. aureus. In contrast, exposure of Halophila ovalis to 1 mg/L of DCMU (a PSII inhibiting herbicide) resulted in the lowering of Fv/Fm to a reading less than 0.1 (Ralph, 2000). Li et al. (2009a) reported that Fv/Fo was a more sensitive indicator of potential photosynthetic activity of plants. Compared to Fv/Fm, Fv/Fo also appeared to be a more sensitive indicator of BPA treatment on the photosynthetic efficiency of PSII in this present study. On D1, Fv/Fo of all plants increased, indicating that the potential photosynthetic activity increased upon transfer of S. aureus plants from agar to liquid growth medium. This was probably the result of increased water availability to the plants. The magnitude of this increase in Fv/Fo was reduced when plants were exposed 178 to higher BPA concentrations, indicating that higher concentrations of BPA might suppress the photosynthetic efficiency of the S. aureus plants. Plants treated with 100 µM BPA also showed a significant decrease in photosynthetic efficiency on D7 of the treatment. The changes in Fv’/Fm’ were similar to those of Fv/Fo, indicating that the efficiency of light capture by oxidized QA was also suppressed by high concentrations of BPA. The value of qp gives an indication of the photochemical capacity of PSII at steady state and is also reflective of the proportion of oxidized QA (Baker, 2008). Generally, Fv’/Fm’, ФPSII and qp decreased in all plants from D0 to D7. Such decreases in PSII photochemistry could be linked to a decrease in NPQ, the nonradiative energy dissipation via the xanthophyll cycle or the non-photochemical quenching (Demmig-Adams, 1990; Eickmeier et al., 1993; Young and Frank, 1996). The data also indicated that plants exposed to 100 and 250 µM BPA were not as efficient as those exposed to 0 and 50 µM BPA in the dissipation of excess energy absorbed. BPA is a phenolic compound. Like phenolic herbicides, which are well-known PSII-inhibitors, BPA at high concentrations is expected to inhibit photosynthesis by binding to the secondary plastoquinone (QB) site in the PSII reaction centre and blocking electron transfer from QA to QB (Rutherford and Krieger-Liszkay, 2001). This would then change the thermodynamic properties of QA, lowering its redox potential and leading to excessive oxidative damage (Rutherford and Krieger-Liszkay, 2001). Photosynthetic capacities of Halophila ovalis (Ralph, 2000) and Lemna minor (Frankart et al., 2003) were greatly reduced by PSII-inhibiting herbicides. As expected, at the end of the study, BPA at 100 and 250 µM reduced ФPSII by 24 – 179 28%, even though total chlorophyll concentrations of plants grown in these BPA levels were higher than those at D0. The carotenoid concentration in S. aureus plants decrease upon exposure of the plants to BPA. Carotenoids play important roles in the photoprotection of photosynthesizing plant cells. They are not only able to react with singlet oxygen and quench 3Chl, but also play a role in energy dissipation in the chlorophyll pigment bed (Demmig-Adams, 1990). The formation of zeaxanthin, in particular, is correlated with the formation of non-photochemical quenching (NPQ) as a result of de-epoxidation of violaxanthin via the xanthophyll cycle ((Demmig-Adams, 1990; Eickmeier et al., 1993; Young and Frank, 1996). Therefore, changes in the carotenoid levels would lead to changes in non-photochemical quenching. In this study, this relation was also observed in plants exposed to 0 µM BPA as NPQ levels increased with increasing carotenoid concentration. In plants exposed to 250 µM, carotenoid concentration decreased with decreasing NPQ levels from D0 to D4, and by D7, NPQ levels started to rise together with carotenoid concentration. Plants exposed to 50 and 100 µM BPA, on the other hand, showed increasing NPQ levels with decreasing carotenoid concentration. A major part of non-photochemical quenching is induced by the acidification of the thylakoid lumen associated with the formation of the proton motive force (Young and Frank, 1996). The increase in NPQ levels observed in these S. aureus plants might be due to increased acidification levels whereas zeaxanthin and the xanthophyll cycle might have a smaller effect on non-photochemical quenching in S. aureus. 180 5.3.4. Total Soluble Sugar (TSS) Concentration TSS can act as indicators of plant stress as they can be representative of plant osmotic adjustments, photosynthesis, signal transduction and induction of defence mechanisms (Morgan, 1984; Ehness et al., 1997; Sheen et al., 1999; Saladin et al., 2003). The concentration of TSS in plants exposed to 0 µM BPA decreased from D0 to D7. This could have resulted from the removal of S. aureus plants from the sucrose-supplemented agar. In contrast, TSS levels increased in all plants exposed to BPA on D1. Studies with Eucalyptus perriniana (Hamada et al., 2002), Nicotiana tabacum (Nakajima et al., 2002; Nakajima et al., 2004,) Ipomoea aquatica (Noureddin et al., 2004) and algal species (Nakajima et al., 2007) have shown that BPA is glycosylated upon uptake into the tissues of these plants. The phenolsulphuric assay used is normally employed for the determination of soluble sugars, but it can also react with glycosides (Saha and Brewer, 1994). It was possible that the sudden increase in TSS observed on D1 was partially due to the conversion of BPA into BPA glycosides as the highest levels of TSS coincided with the highest initial concentration of BPA. The subsequent decrease in TSS levels observed on D4 could be due to the depletion of soluble sugars in the plants, as the plants were also exhibiting decreasing photosynthetic capacity. Slight increases in TSS levels in plants exposed to 100 and 250 µM BPA on D7 indicated that prolonged exposure to high BPA concentration might affect sugar transport and/or metabolism within S. aureus. 5.3.5. Total Soluble Protein (TSP) Concentration Despite the increases in enzyme activities during the exposure of S. aureus plants to BPA, TSP concentration decreased in all plants as the duration of exposure to BPA increased. TSP levels in S. aureus might be affected under stress conditions as 181 proteolysis and de novo synthesis of proteins could be inhibited (Gaspar et al., 2002; Saladin et al., 2003). Therefore, the data obtained indicated that the plants were exposed to abiotic stresses. 5.3.6. Lipid Peroxidation and Antioxidant Enzyme Activities Lipid peroxidation was detected in both the leaves and roots of S. aureus. In studies with rats, BPA resulted in the production of reactive oxygen species (ROS) such as hydroxyl radicals (Obata and Kubota, 2000) and hydrogen peroxide (Bindhumol et al., 2003) in rat tissues. As a phenolic compound, BPA might also undergo oxidation by peroxidases to form phenoxyl radicals (Takahashi and Oniki, 1992; Sakihama and Yamasaki, 2002), and together with ROS, could then react with polyunsaturated fatty acids present in plant cells, resulting in an increase in lipid peroxidation (Kabuta et al., 2003). The values of TBARs were generally higher in the leaves than the roots that were immersed in BPA-containing water. In the present study, the level of lipid peroxidation (indicated by TBARs) in control plants (not exposed to BPA) was always higher than, or equivalent to, those detected in plants exposed to BPA. In other words, exposure of S. aureus to BPA did not increase the levels of lipid peroxidation in the leaves. This might be due to the increased activities of the antioxidant-enzyme defence mechanisms. Exposure to BPA elicited increases in APX, GR and SOD activities, which also changed with the concentration of BPA as well as the duration of exposure of the plants to BPA. The specific activities of APX and GR were highest on D1, and they generally increased with BPA concentrations. The specific activities of SOD were highest on D4 and it was also highest in plants exposed to 250 µM BPA. 182 APX and GR are enzymes involved in the Halliwell-Asada pathway to scavenge for H2O2 (Asada, 1992), which might account for the similar trends observed in S. aureus for both enzymes. The data obtained indicated that oxidative stress levels were high in the plants when they were transferred from the solid agar medium to a liquid growth medium, and it was even higher in plants exposed to BPA. By the end of the study, the plants were able to acclimatize to the new growth conditions, in liquid medium with BPA. This, together with the much lower BPA concentrations after phytoremediation, led to the lowering of APX and GR activities. The specific activity of SOD was highest in plants exposed to BPA for 4 days; it then decreased on D7. The increase in SOD specific activity on D4 coincided with the lowest concentrations of chlorophylls. Also, SOD specific activity was highest in plants exposed to 250 µM BPA. Hence, higher BPA levels taken up by S. aureus plants might have led to the degradation of chlorophylls, and reductions in photosynthesis capacity (ФPSII reduced) and dissipation of excess energy absorbed. This resulted in the formation of O-. and the induction of SOD activity. The specific activity of POD decreased upon transfer of plants from a solid to a liquid growth medium. POD activity increased in control plants (0 µM BPA) as they became acclimatized to the new growth conditions. POD activities of plants exposed to BPA was almost always lower than those of plants not exposed to BPA, indicating that BPA constituted some form of stress. When BPA levels became lower in the growth medium (those with 100 and 250 µM BPA initially) towards the end of the study, POD specific activity increased in the leaves. The initial decrease of POD activity suggested that ROS production, H2O2 in particular, could have increased to such a huge extend that POD became inactivated due to oxidative stress. The 183 subsequent increase in POD activity could be due to the lower BPA levels in the water, after its removal by the plants, leading to a decrease in ROS levels, increasing POD activity. Sun et al. (2008) also observed that POD activity in Ceratophyllum demersum decreased upon exposure to high concentrations of tetrabromobisphenol A (TBBPA), a derivative of BPA, but increased when TBBPA concentrations were low. Interestingly, the maximum specific activities of the various antioxidant enzymes coincided with the lowest levels of leaf lipid peroxidation (lowest TBARs values), particularly in plants exposed to 100 and 250 µM BPA. The data obtained indicated that S. aureus plants depended on both APX and GR for protection against oxidative damage when they were initially exposed to BPA. 5.3.7. The Effectiveness of S. aureus as a Phytoremediator of BPA Based on the results obtained in this study, S. aureus plants were clearly able to remove BPA from water up to a concentration of 250 µM. The rate of BPA removal was also considerably fast as S. aureus plants were able to remove almost completely, the BPA in the 50 and 100 µM BPA-containing water by D2, and half of that present in the 250 µM BPA-containing water by D7 of exposure. BPA exposure resulted in S. aureus plants exhibiting decreases in chlorophyll concentration and photosynthetic capacity, and they also showed an oxidative burst. However, by the end of the study, the plants were able to develop tolerance to BPA, as shown by the upregulation of activities of several antioxidant enzymes (ascorbate peroxidase, glutathione reductase, superoxide dismutase and guaiacol peroxidase) and recovery of chlorophyll levels. 184 Chapter 6. Conclusion This study demonstrated the potential of S. aureus plants to efficiently phytoremediate NO3- and BPA from water, and also demonstrated the physiological and biochemical changes within the plants during exposure to these pollutants. NO3- concentrations in eutrophic waters can range from 6 – 100 mg/L (Dodds et al., 1998; Camargo et al., 2005) and these concentrations were within the NO3limits used for this study. S. aureus plants were able to completely remove NO3- from 20, 50, 100 mg/L NO3--containing water, and more than 80 % of NO3- from 200 mg/L NO3--containing water, by the end of eight weeks, indicating that S. aureus plants would be excellent phytoremediators of eutrophic waters. Furthermore, S. aureus plants were also able to greatly reduce algal growth in water set-ups. Harmful algal blooms are problems commonly associated with eutrophic water bodies and their elimination is beneficial to the phytoremediation of water bodies. Exposure of S. aureus plants to high NO3- concentrations resulted in high growth rates, increases in plant biomass, leaf area, photosynthetic pigment concentrations, TN and TSP levels in the plants. However, PSII efficiency (as determined from chlorophyll fluorescence parameters) and TSS levels in these plants decreased. Signs of oxidative stress, such as elevated lipid peroxidation levels and increased activities of APX were also observed in plants exposed to high NO3concentrations. As the study progressed towards W8, symptoms of N-deficiency were observed in all the plants. Photosynthetic pigment concentrations, chlorophyll fluorescence parameters, TSS and TSP levels decreased, and these were accompanied by increased lipid peroxidation levels and activities of antioxidant enzymes. The changes in NR activity of S. aureus plants were also investigated. Despite decreasing 185 NO3- levels in the water from W0 to W8, NR activity increased in the roots and leaves. Although it is well-known that NR activity increases with increasing NO3concentrations, this was not observed in the S. aureus plants in this study. Several other environmental factors such as light, temperature and anoxia might have larger influences on NR activity in this study. When exposed to water spiked with BPA concentrations of 0, 50, 100 and 250 µM over a period of seven days, S. aureus plants were able to completely remove BPA from 50 and 100 µM BPA-containing waters and 50% of BPA from 250 µM BPA-containing water. Yamamoto et al. (2001) reported that BPA concentrations in leachates of hazardous waste landfills ranged from 1.3 – 17, 200 µg/L (5.69 X 10-3 – 7.53 X 101 µM), whereas Fromme et al. (2002) reported that BPA levels in surface water ranged from 0.0005 – 0.41 µg/L (2.19 X 10-6 – 1.80 X 10-3 µM) and BPA levels in sewage effluents range from 0.018 – 0.702 µg/L (7.89 X 10-5 – 3.08 X 10-3 µM) – all lower than those used in the present study. Hence, S. aureus plants would be good phytoremediators of BPA from these environments. However, BPA, particularly at a high concentration of 250 µM, could cause oxidative stress in S. aureus upon uptake into the plant tissues. The data obtained from assays of antioxidant enzymes and lipid peroxidation indicated an oxidative burst with elevated levels of O-. and H2O2. These ROS caused a decrease in both chlorophyll levels and photosynthesis, subsequently leading to a decrease in TSS levels and plant biomass. In conclusion, S. aureus seemed to be a promising plant species to use for the efficient phytoremediation of NO3- and BPA as it was not only able to acclimatize to partially submerged conditions, but also remove high concentrations of NO3- and BPA 186 from water. Further work should be carried out in order to develop a method or technology for the large-scale, in-field application of S. aureus plants in the phytoremediation of polluted water bodies. 187 Chapter 7. References Agüera E., Ruano D., Cabello P., de la Haba P. 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Dear Dr. Bee-Lian Ong, Your submission entitled "PHYTOREMEDIATION OF BISPHENOL A BY SCINDAPSUS AUREUS (Lindl. & André) Engl." has been received by Environmental and Experimental Botany You may check on the progress of your paper by logging on to the Elsevier Editorial System as an author. The URL is http://ees.elsevier.com/eeb/. Your username is: dbsongbl If you need to retrieve password details, please go to: http://ees.elsevier.com/EEB/automail_query.asp Your manuscript will be given a reference number once an Editor has been assigned. Thank you for submitting your work to this journal. For further assistance, please visit our customer support site at http://epsupport.elsevier.com. Here you can search for solutions on a range of topics, find answers to frequently asked questions and learn more about EES via interactive tutorials. You will also find our 24/7 support contact details should you need any further assistance from one of our customer support representatives. Kind regards, Elsevier Editorial System Environmental and Experimental Botany 208 [...]... and O2 by catalase or enters the Halliwell-Asada pathway and is converted to H2O (Diagram redrawn from Mach and Greenberg, 2004) 26 A variety of antioxidant enzymes are involved in the enzymatic detoxification of ROS (Fig 4) Peroxidases and catalases act specifically to scavenge for H2O2 (Gaspar et al., 2002), but differ in that peroxidases require an additional substrate for catalysis whereas catalases... and wastewater treatment plants (Spivak et al., 1994), and BPA degradation rates were relatively high (Kang et al., 2006b) However, not all bacterial strains have high BPA biodegradability rates Kang and Kondo (200 2a) isolated 10 bacterial strains with BPA biodegradability from river waters, but only a Pseudomonas sp and a Pseudomonas putida strain were able to biodegrade >90% of BPA within 10 days... treatment plants and manufacturing factories, as well as leachates from water landfills (Staples et al., 1998; Yamamoto et al., 2001; Fromme et al., 2002; Kang et al., 200 6a) From these sources, BPA can then make its way into freshwater and marine bodies, and even into groundwater In Japan, BPA pollution is widespread, as reported by the Environmental Agency of Japan (Yamamoto et al 2001) Levels of. .. the standard The concentration of TSP was expressed as mg proteins per g FW and per g DW 3.1.11 Determination of Nitrate Reductase (NR) Activity (EC 1.6.6.1) The activity of NR was determined according to Hageman and Hucklesby (1971), Ahmad and Abdin (1999) and Taghavi and Babalar (2007) with some modifications Leaf and root materials (0.1 g FW each) were ground in 1 ml of extraction buffer containing... Furthermore, BPA degradation by microbes are affected by temperature, bacterial counts (Kang and Kondo, 2002b), as well as aerobic and anaerobic conditions (Kang and Kondo, 200 2a) 20 In comparison to the above methods, phytoremediation of BPA appears to be more desirable As a result, in recent years, research on phytoremediation of BPA has been gaining popularity, with biodegradation studies involving plant enzymes,... Momordica charantia (bitter gourd) (Karim and Husain, 2009), and Cochlearia armoracia (horseradish) (Sakuyama et al., 2003) have shown to be able to oxidize BPA Yoshida et al (2002) also reported that polyphenol oxidases extracted from vegetables (e.g potato, mushroom, eggplants, edible burdock and yacon) were able to oxygenate BPA to quinones Fungal laccases extracted from Trametes villosa (Fukuda et al.,... et al., 2003), as well as increased aggression and anxiety (Kawai et al., 2003; Ryan and Vandenbergh, 2006) In female rats, daily exposure to BPA can disrupt the development of nervous systems, sexual dimorphic behaviours and increased anxiety (Ryan and Vandenbergh, 2006) as well as advanced puberty (Howdeshell et al., 1999) ) It has also been demonstrated in pregnant rats that BPA absorbed into the... chloroplast electron transport chains (Gaspar et al., 2002; Mach and Greenberg, 2004) The reactions that produce ROS are summarized in Fig 3 High levels of ROS can cause protein oxidation, DNA damage and lipid peroxidation (Mach and Greenberg, 2004) Protein oxidation can disrupt the protein structure through side chain alterations and backbone cleavages, leading to denaturation, aggregation and degradation... electrodialysis and chemical adsorption (Menkouchi Sahli et al., 2008) and even microbial treatment (Ayyasamy et al., 2007) These methods may be effective in the removal of NO3-, but they are expensive to set up on a big scale (Kapoor and Viraraghavan, 1997; Ayyasamy et al., 2009) Also, microbial treatments require external supplies of organic carbon and nutrients, which may not be fully consumed and can... physiological, hormonal, reproductive and even behavioural changes (Richter et al., 2007) In male rats, daily exposure to low levels of BPA can cause oxidative stress in the liver (Bindhumol et al., 2003), kidneys (Kabuto et al., 2003), striatum (Obata and Kubota, 2000) and epididymal sperm (Chitra et al., 2003), permanent alterations in the male prostate and hypothalamic-pituitary-gonadal axis (Ramos et al., ... Determination of Nitrate Reductase (NR) Activity (EC 1.6.6.1) The activity of NR was determined according to Hageman and Hucklesby (1971), Ahmad and Abdin (1999) and Taghavi and Babalar (2007)... peroxidase Peroxidases extracted from Glycine max (soybean) (Caza et al., 1999), Momordica charantia (bitter gourd) (Karim and Husain, 2009), and Cochlearia armoracia (horseradish) (Sakuyama et al.,... of NO3- and BPA 30 Chapter Materials and Methods 3.1 Phytoremediation of Nitrates by S aureus 3.1.1 Plant Materials and Growth Conditions Shoot cuttings of Scindapsus aureus were grown partially