Thermoluminescence as a complementary technique for the toxicological evaluation of chemicals in photosynthetic organisms

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 1 Thermoluminescence as a complementary technique for the toxicological evaluation of chemicals in photosynthetic organisms Guillermo Repettoa, Jorge L. Zuritaa,b, Mercedes Roncelc, José M. Ortegac aDepartamento de Biología Molecular e Ingeniería Bioquímica, Universidad Pablo de Olavide, Carretera de Utrera km. 1, 41013 Seville, Spain bPresent address: Fundación Progreso y Salud, Junta de Andalucía, Américo Vespucio 5, 41092 Seville, Spain cInstituto de Bioquímica Vegetal y Fotosíntesis, Universidad de Sevilla-CSIC, Américo Vespucio 49, 41092 Seville, Spain Corresponding author: G. Repetto, Departamento de Biología Molecular e Ingeniería Bioquímica, Área de Toxicología, Universidad Pablo de Olavide, Carretera de Utrera km. 1, 41013 Seville, Spain. Fax: 34-954349813; Email: gr[email protected] Keywords: Chlorella vulgaris; chemical toxicity; lipid peroxidation, photosystem II; thermoluminescence Abbreviations: B-band: TL band due to S2/3QB- recombination; BrB: bromobenzene; C-band: TL band due to D+QA- recombination; CQ: chloroquine; D: tyrosine D of photosystem II; DEA: diethanolamine; EC50: concentration of test chemical that modified the biomarker by 50% in comparison with appropriate untreated control; HTL: high temperature thermoluminescence; InNO3: indium nitrate; PpG: propyl gallate; PSII: photosystem II; Q- band: TL band due to S2QA- recombination; QA and QB: primary and secondary quinonic electron acceptors of PSII; S2 and S3: oxidized states of the manganese oxygen evolving Manuscript Click here to view linked References 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 2 complex of PSII; SMFA, sodium monofluoroacetate; STL: standard thermoluminescence; TL: thermoluminescence; tmax: temperature of the maximum intensity of a TL band. Abstract Thermoluminescence is a simple non-invasive technique very useful for study electron transfer reactions on photosystem II (standard thermoluminescence) or the level of lipid peroxidation in membranes (high temperature thermoluminescence) in photosynthetic organisms. Both techniques were used to investigate the effects produced on Chlorella vulgaris cells by six compounds: the chemical intermediates bromobenzene and diethanolamine, the antioxidant propyl gallate, the semiconductor indium nitrate, the pesticide sodium monofluoroacetate and the antimalarian drug chloroquine. Electron transfer activity of the photosystem II significantly decreased after the exposure of Chlorella cells to all the six chemicals used. Lipid peroxidation was slightly decreased by the antioxidant propyl gallate, not changed by indium nitrate and very potently stimulated by diethanolamine, chloroquine, sodium monofluoroacetate and bromobenzene. For five of the chemicals studied (not bromobencene) there is a very good correlation between the cytotoxic effects in Chlorella cells measured by the algae growth inhibition test, and the inhibition of photosystem II activity. The results suggest that one very important effect of these chemicals in Chlorella cells is the inhibition of photosynthetic metabolism by the blocking of photosystem II functionality. In the case of sodium monofluoroacetate, diethanolamine and chloroquine this inhibition seems to be related with the induction of high level of lipid peroxidation in cells that may alter the stability of photosystem II. The results obtained by both techniques supply information that can be used as a supplement to the growth inhibition test and allows a more complete assessment of the effects of a chemical in photosynthetic organisms of aquatic ecosystems. 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 3 1. Introduction Model systems and bioassays are used in environmental toxicology to provide information for risk assessment of chemicals. To improve the efficiency in evaluating the hazard of a large number of chemicals, providing mechanistic information, it would be helpful to carry out batteries of rapid and mode of action based screening tests (Katsumata et al., 2009; Repetto, 2013). Standard (STL) and high temperature thermoluminescence (HTL) measurements are useful techniques for the study of the effects of stress conditions in photosynthetic organisms (Ducruet, 2003; Havaux, 2003). Its use in aquatic toxicology has been however very limited. Luminescence is a general term which indicates a light emitting process when the molecules in an electronically excited state decay to the stable ground state. Luminescence from organic or inorganic material can be also thermally induced and enhanced by heating the sample in the dark. This process is called thermoluminescence and describes the emission of light at characteristic temperatures from samples containing chemiluminescence active species, radical pair states or electron hole pairs (Ducruet, 2003; Havaux, 2003). STL is normally used to study the photochemical activity of photosystem II (PSII). PSII is a large pigment-protein complex which uses light energy to water splitting producing molecular oxygen as by-product and providing an unlimited supply of reducing power needed to convert CO2 into the organic molecules of life (for a review see Diner and Babcock, 1996). The reaction centre (RC) of PSII is composed of an heterodimer formed by the D1 and D2 membrane proteins that bears all the cofactors needed for charges separation and stabilization: six chlorophylls (Chl) (including four of the Chl centre, P680), two pheophytins, two plastoquinones QA and QB, one non-heme iron atom, one tetranuclear manganese cluster (Mn4CaO5) and two redox active tyrosines. Photons are captured by the light-harvesting pigments and driven as excitation energy to P680 forming an excited state called P680*. The 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 4 P680* state is highly reducing, and an electron is rapidly transferred to the terminal plastoquinone (PQ) electron acceptor bound to the QB site within the D1 protein. On receiving two electrons, the PQ is protonated to plastoquinol (PQH2) and leaving the QB site in the PSII complex. The P680 radical cation (P680+) formed in primary charge separation has a very high redox potential, estimated to be approx. +1.3 V. This high-oxidation state is required to drive the water-splitting reaction that occurs at a catalytic site, the manganese cluster (for a review see Rappaport et al., 2002; Diner and Rappaport, 2002). The photochemical activity of PSII complex can be measured by using the STL technique. Photosynthetic luminescence is a weak light emission originating from recombination of positive and negative charge pairs generated by a prior illumination in PSII, some of these leading to the recreation at a low yield of an exciton in the chlorophyll antenna, with a probability to deactivate as fluorescence. Luminescence decay phases can be better resolved by TL emission technique, which consists in illuminating the sample at a temperature sufficiently low to make negligibly small the recombination rate of the charge pairs under investigation, then to reveal them successively as TL bands by a progressive warming (for a review see Vass and Inoue, 1992; Ducruet, 2003; Sane, 2004). After a sequence of short flashes, a so-called B-band of TL, located between 15 and 40 °C depending on species and conditions, is observed in healthy dark-adapted photosynthetic material, such as leaves and algae. This emission is due to the recombination of S2/S3QB- pairs, QB being the secondary quinonic electron acceptor and S2/S3, the states of the oxygenevolving complex storing positive charges (Rutherford et al., 1982; Demeter and Vass, 1984). The changes on the intensity and temperature maximum of this band reflect alterations in the functionality of PSII complex. Herbicide treatment (diuron, atrazine), which blocks the QA to QB electron transfer, induces the appearance of a Q-band peaking at about 5 °C, due to the S2QA- recombination. A C-band at about 55 °C can also be detected in particular conditions. 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 5 This TL band is due to D+QA- recombination (Demeter et al., 1993, Johnson et al., 1994), D+ being the oxidized form of tyrosine D on the inactive branch of PSII. STL measurements are useful for the study of the effects of stress conditions on PSII electron transport in intact leaves, in algal and cyanobacterial cells, as well as in thylakoids and isolated membrane complexes (Rutherford and Inoue, 1984; Rutherford et al., 1984a; Zurita et al., 2005c; Ducruet and Vass, 2009). The technique is simple, inexpensive and powerful in monitoring several aspects of the PSII electron transport. The fact that alterations in the peaks of the TL curves may be interpreted as changes in the redox potentials of donors and acceptors on PSII makes this technique a useful tool for understanding the process of adaptation, responses to abiotic and biotic stresses, and structural modifications in the PSII polypeptides. HTL is a useful technique for the study of lipid peroxidation processes in photosynthethic organisms (Havaux, 2003). Lipid peroxidation is a complex process known to occur in both plants and animals. This process involves a chain reaction initiated by the hydrogen abstraction or addition of an oxygen radical, a rearrangement of the double bonds in unsaturated lipids and the formation and propagation of lipid radicals. As a result of these reactions takes place an eventual destruction of membrane lipids with the production of a variety of breakdown products, including alcohols, ketones, alkanes, aldehydes and ethers (Dianzani and Barrera, 2008). In stress situations the reactive oxygen and nitrogen species are generated at higher than normal rates, and as a consequence, lipid peroxidation occurs. In photosynthetic organisms a large amount of oxygen is formed in thylakoids in the vicinity of powerful oxidation-reduction systems that readily reduce oxygen to dangerous superoxide and produce harmful singlet oxygen which could initiate lipid peroxidation processes (Dalton, 1995; Repetto et al., 2012). 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 6 The lipid peroxidation is commonly detected and quantified by measuring by-products of the reaction, such as ethane, conjugated diens, malondialdehyde, and other carbonyl compounds (Heath and Packer, 1968; Pryor and Castle, 1984; Chatterjee and Agarwal, 1988; Havaux and Niyogi, 1999). The most commonly used test is called TBARS Assay (thiobarbituric acid reactive substances assay) where the concentration of malondialdehyde is determined by reaction with thiobarbituric acid to yield a fluorescent product. However, there are other sources of malondialdehyde, so this test is not completely specific for lipid peroxidation. In recent years, the development of immunochemical detection of 4- hydroxynonenal histidine adducts has opened more advanced methodological possibilities for qualitative and quantitative detection of lipid peroxidation in various human and animal tissues (Weber et al., 2013). These chemical methods cannot be however routinely used for the assessment of lipid oxidation in phytoplankton because the analysis requires high amounts of biological material, and they are often time-consuming and, therefore, not adequate for high throughput screening. The second problem is the difficulty of separating the peroxidative reactions that take place in phytoplankton from the reactions occurring in no phytoplankton materials. Recently, a new TL-based technique has been proposed as a useful method to detect lipid peroxidation in photosynthetic organisms that may solve some of the aforementioned problems (Vavilin and Ducruet, 1998; Havaux, 2003; Ducruet and Vass, 2009). When chlorophyll-containing material is gradually heated to temperatures above 70 °C, a rise of chlorophyll luminescence emission is often observed. This increase in the luminescence intensity at high temperature was named HTL (Matorin et al., 1989; Venediktov et al., 1989). A broad HTL band peaking around 135 °C (HTL2 band) is generated in the reaction of thermal decomposition of peroxidized lipids that, in turn, leads to the formation of carbonyls in triplet state followed by migration of excitation energy toward chlorophylls (Venediktov et 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 7 al., 1989; Vavilin et al., 1991; Vavilin and Ducruet, 1998). Its amplitude has been correlated with the accumulation of malodialdehyde, an indicator of lipid peroxidation in standard chemical tests (Vavilin and Ducruet, 1998). Consequently, HTL2 band measurements have been used to study in vivo lipid peroxidation in plants subjected to various oxidative stress conditions (Merzlyak et al., 1992; Havaux and Niyogi, 1999; Ducruet, 2003; Baroli et al., 2004; Havaux, 2003). In a future perspective HTL might be applied as a fast screening tool before more sophisticated methods like mass spectrometry are applied to identify the most sensitive molecular target in a given tissue. STL and HTL techniques have been applied in several in vitro toxicological studies, mainly using thylakoids and cell cultures (Ducruet et al., 2007). Using thylakoids isolated from leaves of greenhouse grown peas (Pisum sativum L.), Mohanty et al. (1989) suggested that Cu (II) does not block electron flow between the primary and secondary quinonic electron acceptor but modifies the secondary quinonic acceptor site in such a way that it becomes unsuitable for further PSII photochemistry. Horváth et al. (1998) reported that the amplitude of the B-band was gradually decreased, which corresponds to the Cu (II) induced inactivation of Tyrosine Z. The simultaneous stepwise shift in the peak position of the B-band indicated, however, that S3QB− charge recombination is more resistant to Cu (II). Nitrite, but not formate, fluoride or nitrate anions, inhibited PSII in spinach thylakoids, probably at the donor side (Sahay et al., 2006). Toxicological applications of HTL have been reported by several authors. A rise of thermoluminescence at 120-130 °C was detected when cultures of the unicellular green alga Chlorella (C.) pyrenoidosa were exposed for 5 h to 10 µM of the oxidative stress promoter methyl viologen (Vavilin and Ducruet, 1998). Hörcsik et al. (2007) found photodestruction of D1 protein in C. pyrenoidosa caused by Cr (VI) due to enhanced oxidative stress and lipid peroxidation, as indicated by the appearance of a HTL2 band. Schumann et al. (2009) have 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 8 developed a rapid and easy to handle TL based technique for evaluation of carbon tetrachloride-induced oxidative stress on rat hepatocytes. The aim of this work has been the application of two simple, non-invasive and inexpensive TL techniques (STL and HTL) to study the alterations produced on the freshwater green alga C. vulgaris by six chemicals. This green alga was selected as a representative of primary producers. It is a genus of single-cell green algae, widely distributed in freshwater and seawater. It has a short cycle of growth, which makes it ideal for aquatic ecotoxicity studies (Wong et al., 1997). The chemical intermediates bromobenzene and diethanolamine, the antioxidant propyl gallate, the semiconductor indium nitrate, the pesticide sodium monofluoroacetate and the antimalarian drug chloroquine were selected as representative of several chemical groups according to their use. The results have been compared with those obtained by our group by using the algae growth inhibition test (Zurita et al., 2005a,b; Zurita et al., 2007a,b,c,d). 2 Material and Methods 2.1 Cultures The unicellular green algae Chlorella vulgaris var. viridis, kindly provided by Dr. Muñoz- Reoyo (CISA, Spain), were grown as previously described (Ramos et al., 1996) in Bold’s Basal Medium (BB medium) (Culture Collection of Algae and Protozoa, Cambridge, UK; http://www.ccap.ac.uk/media/recipes.htm) under a water-saturated sterile atmosphere containing 5% CO2 at 22 °C, constant agitation and a light dark cycle of 16:8 h (white light of 40 E m-2 s-1). 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 9 2.2 Chemicals Stock solutions of bromobenzene (CAS number 108-86-1), indium nitrate (CAS number 13770-61-1), sodium monofluoroacetate (CAS number 62-74-8), diethanolamine (CAS number 111-42-2) and chloroquine (CAS number 54-05-7) were prepared daily in deionised water. Propyl gallate (CAS number 121-79-9) was solubilised in dimethyl sulfoxide. All the chemicals were from Sigma-Aldrich (USA), unless sodium monofluoroacetate (Honeywell´s Riedel-de Haën). They were sonicated for 30 min and sterilized by filtration through a 0.22 µm Millipore filter. The concentrations of the chemicals used in this work were chosen taking into account the EC50 values obtained for growth inhibition (biomarker) of C. vulgaris cells after 24 h exposure to the different chemicals, as previously published (Zurita et al., 2005a,b; Zurita et al., 2007a,b,c,d). The EC50 value is the concentration of the chemical that modified the biomarker (growth inhibition, for C. vulgaris) by 50% in comparison with appropriate untreated control. Two different concentrations of chemicals were used, one below EC50 values and the second one above or close to the EC50 values. The two concentration used and the EC50 values for the six chemicals are shown in Table 1. STL and HTL experiments were carried out using cultures of C. vulgaris in exponential phase of growth (with about 106 cell ml-1). The cultures were exposed in BB medium to the two different concentrations of chemicals selected in 80 ml flasks for 24 h at 20 °C using constant agitation at 40 r.p.m., under a continuous cold light source of 110 E m-2 s-1. The suspensions were then centrifuged at 2000 r.p.m for 10 min, quantified at 665 nm and adjusted to 0.1 mg chlorophyll ml-1 by using BB medium. Control experiments in the absent of the chemicals were also treated as above described. 2.3 Thermoluminescence measurements 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 16 stimulation on the antioxidant enzyme glucose-6-phosfate dehydrogenase in the PLHC-1 fish cell line after a treatment with 1.8 mM bromobenzene, confirming the situation of oxidative stress. This significant increase in lipid peroxidation induced by bromobenzene in Chlorella cells can be also related to the accelerated membrane phospholipid degradation described by Lamb et al. (1984). The decrease in the B-band observed after the treatment with the highest concentration of bromobenzene (Figure 2, BrB) can also be explained taking into account the effects on the PSII activity of the phospholipid degradation processes. Thus, it seems that the main cytotoxic effects of BrB seem not to be related with the inhibition of PSII activity. Other effects on the Chlorella metabolism induced by BrB (as lipid peroxidation) must be responsible for the growth inhibition measured in previous work (Zurita et al., 2007a). Propyl gallate is an antioxidant widely used to preserve and stabilize the freshness, nutritional value, flavour and colour of foods, fats and oils, cosmetics and medicinal preparations (Van der Heijden et al., 1986). Propyl gallate is not very toxic to Chlorella, showing much more toxicity in other systems (Zurita et al., 2007c). A consistent decrease in PSII functionality (B-band) was detected in parallel with cytotoxicity (Table 1 and Figure 2, PpG). Propyl gallate uncouple oxidative phosphorylation by increasing the permeabillty of mitochondrial inner membranes to protons (Fusi et al., 1991). A similar effect on chloroplast may explain the inhibition of TL B-band of PSII. Although there are many studies that demonstrate the antioxidant capacity of propyl gallate (Karthikeyan et al., 2005), other authors have reported oxidant properties (Kobayashi et al., 2004; Kawanishi et al., 2005). The mitochondria alternative oxidase, that catalyzes a cyanide-resistant reduction of oxygen to water without translocation of protons, is specifically inhibited by propyl gallate, stimulating H2O2 production in plant mitochondria (Josse et al., 2003). High concentrations of propyl gallate have been reported to produce loss of mitochondrial membrane potential and to be oxidant in non-vegetal systems due to depletion 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 17 in glutathione, stimulating glucose-6-phosphate dehydrogenase activity (Zurita et al., 2007c; Han and Park, 2010). Therefore, in order to clarify this discrepancy, the protective and toxic effects of propyl gallate, we have investigated the possible lipid peroxidation induced by this chemical by using HTL technique (Figure 4, PpG). Our results clearly support the antioxidant properties of propyl gallate. The observed slight reduction on lipid peroxidation (Figure 4, PpG), in comparison to the control, is in agreement with its proposed antioxidant capacity (Kawanishi et al., 2005). In fact it has been recently demonstrated in PSII membranes isolated from spinach leaves, that the scavenging capacity of radicals by propyl gallate significantly suppressed 1O2 formed by oxidation of organic molecules (lipids and proteins) (Yadav and Pospíšil, 2012). Indium nitrate is used in the industry of batteries, to prevent build-up of hydrogen gas within sealed battery casings, plating and as semiconductor in other chemical applications. It is also used for the treatment of Hodgkin and non-Hodgkin's lymphoma and for the management of hypercalcemia (Lipcamon and Sahl, 2002). However, there are no reports on its phytotoxicity. The proliferation of Chlorella cells is very sensitive to indium nitrate exposure, showing a parallel inhibition of PSII functionality as shown by the significant decrease of B-band (Figure 2, InNO3). It has been reported that treatment of rats with indium trichloride produced changes in heme biosynthesis (Woods et al., 1979). A similar effect on Chlorella could explain the decrease on PSII activity. The two concentrations of indium nitrate tested in this work did not affect the lipid peroxidation level in Chlorella cells in comparison with control conditions (Figure 4). These results are not in agreement with the reported induction by indium of lipid peroxidation in mice liver (Verstraeten et al., 1997; Repetto and del Peso, 2012). Indium promotes membrane rigidification and lateral phase separation; both parameters are well correlated with the ability 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 18 of the ions to stimulate Fe2+-initiated lipid peroxidation. According to Zurita el al. (2007d), toxicity to fish cells at higher concentrations (5-7 mM) than those investigated with HTL in alga seems to be mediated by oxidative stress, and particularly by NADPH-dependent lipid peroxidation. Thus, taking into account our results, in Chlorella cells the toxic effect of indium nitrate seems not to be related with the lipid peroxidation process. Sodium monofluoroacetate is one of the most potent pesticides. It is mainly used as a rodenticide and as a vertebrate pest control agent. At least since the 1940s, it is known as one of the most toxic substances around the world, retaining the restricted use classification imposed by the USEPA in 1978. Leaf chlorosis, necrosis and suppression of seeding growth were reported in the plant Helianthus annus exposed to 6.5 mM sodium monofluoroacetate (Cooke, 1976). From the soil organisms evaluated by O’Halloran et al. (2005), including the earthworm Eisenia fetida and microorganisms, lettuce (Lactuca sativa) was more sensitive than oats (Avena sativa). Lettuce seedling emergence and seedling shoot growth were adversely affected at a soil concentration of 7 mg per Kg of sodium monofluoroacetate. Zurita et al. (2007b) reported the lowest non observed adverse effect level in C. vulgaris for sodium monofluoroacetate (0.1 mM) in comparison to D. magna, V. fischeri and fish cell lines. Sodium monofluoroacetate triggers its main toxic action by blocking the Krebs cycle. The compound is incorporated into fluoroacetyl coenzyme A, which is condensed with oxalacetate to form fluorocitrate, a process dubbed as ‘‘lethal synthesis’’ (Buffa and Peters, 1950). The intensity of HTL2 band was increased in Chlorella more than four-fold by sodium monofluoroacetate, reflecting a status of oxidative stress probably generated by the metabolic blockage. In fact, Dojcinovic et al. (2005) demonstrated that inhibition of the mitochondrial electron transport chain of the tricarboxylic acid cycle by sodium monofluoroacetate causes increased expression of nucleus-encoded alternative oxidase genes in plants. In addition, Zurita et al. (2007b) reported that glucose-6-phosphate dehydrogenase activity was the only 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 19 biomarker stimulated in RTG-2 fish cells exposed to sodium monofluoroacetate. Taking into account the protection of fluoroacetate toxicity observed in PLHC-1 cells pre-treated with sulphydryl reagents, another mechanism of action of sodium monofluoroacetate seems to be the binding to sulphydryl groups, including those of glutathione. The 30% reduction on the intensity of B-band of TL with the highest concentration of the chemical may be also related to the oxidative stress processes induced by sodium monofluoroacetate. However, this chemical induced another major change in the characteristics of the TL emission signal. Whereas a single TL band is generally observed in untreated Chlorella cells, sodium monofluoroacetate produced a more broadened TL curve with a significant upshift of the tmax. These results suggest that sodium monofluoroacetate could also induce major changes in the ratio among the two components responsible for the emission of B-band: S2/S3 states of the manganese cluster and/or the reduced state of QB (Vass and Inoue, 1992). Diethanolamine (DEA) is a viscous liquid widely used as chemical intermediate, anticorrosion agent in metalworking fluids and surface-active agent in cosmetic formulations, pharmaceuticals and agricultural products. It presents low cytotoxicity to Chlorella cell proliferation (Zurita et al., 2005b), while the algae Skeletonema costatum and Phaeodactylum tricornutum have been reported to be more sensitive in a recent ecotoxicological evaluation (Libralato et al., 2010). However, the TL emission bands in Chlorella were very sensitive and dose dependent. At concentrations around the EC50 for cell proliferation, the intensity of B- band showed a reduction of 15%, and disappeared when the concentration was doubled (30 mM). In adition, at this last concentration, another major change was a potent stimulation in lipid peroxidation, with an approximately 100-fold increase. Diethanolamine probably inhibits the mitochondrial fatty acid beta-oxidation. Zurita et al. (2005b) have reported a marked steatosis in PLHC-1 cells exposed to diethanolamine. Toxicity studies on vertebrates have shown that diethanolamine inhibits cellular choline 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 20 uptake, causing a wide range of indirect effects, including increased susceptibility to oxidative damage, negative effects on phospholipid metabolism, altered mitochondrial function and structure, DNA methylation, altered transcription of genes involved in cell growth regulation and tumour development (Hansen et al., 2010). In the marine copepod Calanus finmarchicus, diethanolamine caused the up-regulation of the transcription of genes involved in protein catabolism, metal ion homeostasis, defence against oxidative stress, accompanied by altered expression of fatty acid derivates, amino acids and cholines (Hansen et al., 2010). All these metabolic alterations, including the observed important lost in the PSII activity, can be related to the interference of phospholipids accumulation in the metabolism. It has been reported that phospholipid accumulation produces a loss of mitochondrial integrity (Barbee and Hartung, 1979), a process that can also occur in chloroplasts. The HTL experiments of this work showed enormous levels of lipid peroxidation in Chlorella cells treated with diethanolamine (Figure 4, DEA). This suggests that this chemical besides inducing an accumulation of lipids in the chloroplasts, also promotes its degradation by peroxidation processes. Interestingly, the lower concentration (15 mM) of diethanolamine induced the appearance of a more broadened TL glow curve with a shoulder peaking at about 46 ºC (Figure 2, DEA and Figure 3). We tentatively assigned this component appearing at higher temperatures to the AG emission band usually induced by far red illumination (Miranda and Ducruet, 1995). A similar AG-band has been observed in Chlamydomonas cells excited by white light (Ducruet et al., 2011). The presence of an AG-band in TL curves recorded in cells excited by white light flashes reflects the amount of PSII centres in the S2/3QB non-radiative state immediately after preillumination, in which the arrival of an electron transferred from stroma along cyclic/chlororespiratory pathway(s) produces the radiative state S2/3QB- that emits luminescence (Ducruet el al., 2005). This back electron transfer can be induced by warming that activates the cyclic pathway(s) leading to an AG emission as S2/3QB centers are 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 21 converted to S2/3QB- by electrons arriving from an unknown stroma reductant (Ducruet el al., 2005). In Chlorella cells grown in the presence of diethanolamine, phospholipids can provide additional NADPH in the stroma through the respiratory pathway (Alric, 2010), thus stimulating the cyclic versus linear electron transfer pathway to maintain the ATP/NADPH ratio required by the Benson-Calvin cycle. This stimulation of the cyclic/chlororespiratory pathway can explain the appearance of the AG-band in TL emissions induced by white light. In Chlorella cells incubated in the presence of 30 mM diethanolamine there is not photosynthetic electron transfer on PSII (see Figure 2, DEA). However, the growth inhibition test has shown that at this concentration the percentage of Chlorella living cells was still about 40% (Zurira et al., 2005b). So, diethanolamine should activate alternative photosynthetic metabolism, without the participation of PSII (chlororespiration, cyclic electron transport around PSI, etc.) or not photosynthetic metabolism processes that maintain the cells viability. According to Foster (1994), the lysosomotropic drug chloroquine is probably the second or third most widely consumed drug in the world. It is still the main pharmaceutical used for the prophylaxis and treatment of malaria, and is also used for treatment of autoimmune diseases, collagen vascular and dermatological disorders and extraintestinal amebiasis. In addition, CQ has been proposed as a promising inexpensive drug in the management of HIV disease (Romanelli et al., 2004). It is the most potent compound evaluated according to the inhibition of the proliferation of Chlorella cells, with an eight-fold increase in toxicity when the exposure time period was increased from 24 to 72 h (Zurita et al., 2005a). Jjemba (2002) demonstrated that the growth of soybean plants was decreased by increasing concentrations of chloroquine, while the number of bacteria and protozoa in soil was either unchanged or increased. Although the inhibition of bioluminescence in V. fischeri 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 22 is less sensitive, the crustacean D. magna is even more sensitive than Chlorella (Zurita et al., 2005a). Thermoluminescence measurements in Chlorella cells were particularly sensitive to chloroquine, showing important reductions on B-band intensity (Figure 2, CQ), while HTL2 intensity was increased 5.5-fold by 0.25 mM chloroquine, in comparison to control (Figure 4, CQ). These results agree with the previously reported oxidative stress in rats induced by chloroquine, with the consequence of lipid peroxidation as mechanism of the nephrotoxicity (Murugavel and Pari, 2004). According to Sugioka et al. (1987), the ferriprotoporphyrin IX- chloroquine complex remarkably promotes peroxidative cleavage of unsaturated phospholipids. The significant decrease of the activity of PSII can be related to lipid peroxidation on chloroplast and also with the possible binding of chloroquine to porphyrin groups of the PSII cofactors. In conclusion, for five of the chemicals studied (not BrB) there is a very good correlation between the cytotoxic effects in Chlorella cells measured by the algae growth inhibition test (Zurita et al., 2005a,b; Zurita et al., 2007a,b,c,d), and the inhibition of PSII activity measured by STL. Thus, we may propose that one very important effect of these chemicals in Chlorella cells is the inhibition of photosynthetic metabolism by the blocking of PSII functionality. In the case of sodium monofluoroacetate, diethanolamine and chloroquine this inhibition seems to be related with the induction of high level of lipid peroxidation in cells that may alter the stability of PSII. Other toxic mechanisms different from lipid peroxidation must be however responsible for the inhibition of PSII induced by PpG and InNO3. The inhibition of growth in Chlorella cells induce by BrB seems not to be related by the blocking of photosynthetic electron transfer activity on PSII. Consequently, although no single biomarker is viable in gaining a comprehensive understanding of xenobiotic stress (Brain and Cedergreen, 2009), the quantification of TL 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 23 emissions is a useful technique to study specific effects of environmental chemicals on photosynthetic organisms. STL and HTL techniques provide specific information on photosynthetic electron transport and on lipid peroxidation induced by oxidative stress, respectively. This information is complementary to that obtained by the growth inhibition test and allows a more complete assessment of the effects of a chemical in organisms of aquatic ecosystems, and about the mechanisms of the toxicity. Conflict of interest None of the authors had any conflicts to declare. Acknowledgements This work was supported by grants BFU2007-68107-C02-01/BMC from the Spanish Ministry of Science and Innovation and BIO2012-35271 and CTM2012-31344 from the Spanish Ministry of Economy and Competitiveness, and the PADI CVI-261, BIO-022 and CTS-996 from the Andalusia Government. All of these grants were partially found by the EU FEDER program. J.L. Zurita was the recipient of a grant for the training of research personal from the Ministry of Science and Innovation. The authors thank S. Jiménez and A. 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Pyrogallol-induced calf pulmonary arterial endothelial cell death via caspase-dependent apoptosis and GSH depletion. Food Chem. Toxicol. 48, 558-563. Havaux, M., 2003. Spontaneous and thermoinduced photon emission: new methods to detect and quantify oxidative stress in plants. Trends Plant Sci 8, 409-413. Havaux, M., Niyogi, K.K., 1999. The violaxanthin cycle protects plants from photooxidative damage by more than one mechanism. Proc. Nat. Acad. Sci. USA 96, 8762-8767. 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 32 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 33 Figure 3. Thermoluminescence glow curves from Chlorella vulgaris cells exposed for 24 h to diethanolamine (DEA). Chlorella cells suspensions were exposed to two different concentrations of diethanolamine (15 and 30 mM) as described in Material and Methods section. After the treatment, 50 l of cells suspension were dark-adapted for 2 min at 20 ºC and subsequently cooled to 1 ºC during 1 min. One saturating single turn-over flash was given at the end of this period. Luminescence emission was then recorded while warming samples from 0 oC to 80 oC at a heating rate of 0.5 oC per s. For other technical details see Material and Methods section. Symbols: Control, non-treated samples; DEA 1, Chlorella cells exposed to 15 mM diethanolamine; DEA 2, Chlorella cells exposed to 30 mM diethanolamine. The B and the AG-bands resulting from the decomposition of the TL curve induced in Chlorella cells exposed to the lower concentration of the chemical are shown in dots. 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 34 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 35 Figure 4. High temperature thermoluminescence glow curves from Chlorella vulgaris cells exposed for 24 h to six different chemicals. Bromobenzene (BrB), propyl gallate (PpG), indium nitrate (InNO3), sodium monofluoroacetate (SMFA), diethanolamine (DEA) or chloroquine (CQ). Chlorella cells were exposed to two different concentrations of the chemicals as described in Material and Methods section. After the treatment, 50 l of cells suspension were adsorbed by filtration on a piece of filter paper, dark adapted for 10 min at 20 ºC and cooled to 1 ºC for 1 min. Luminescence emission was then recorded without flash excitation while warming samples from 10 ºC to 160 ºC at a heating rate of 0.1 oC per s. In order to desiccate the samples and prevents any oxidation induced by high temperature, N2 was flushed on the samples during HTL experiments. For other technical details see Material and Methods section. Symbols: (●) Control non-treated samples; (○) Chlorella cells exposed to the lower concentration of the chemical; (Δ) Chlorella cells exposed to the higher concentration of the chemical. Lower and higher concentrations of the chemicals, respectively: BrB, 1 and 2 mM; PpG, 0.5 and 1 mM; InNO3, 0.3 and 0.7 mM; SMFA, 5 and 20 mM; DEA, 15 and 30 mM; CQ, 0.1 and 0.25 mM. 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 36