Disturbance of photosystem II-oxygen evolution complex induced the oxidative damage in Chlorella vulgaris under the stress of cetyltrimethylammonium chloride

Chemosphere 223 (2019) 659e667

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Disturbance of photosystem II-oxygen evolution complex induced the oxidative damage in Chlorella vulgaris under the stress of cetyltrimethylammonium chloride Han Zhang, Na Liu, Jinfeng Zhao, Fei Ge*, Yin Xu, Yuehui Chen Department of Environment, College of Environment and Resources, Xiangtan University, Xiangtan 411105, PR China

h i g h l i g h t s

g r a p h i c a l a b s t r a c t


Electron transfer and oxygen evolution in PSII-OEC was disturbed by CTAC.
Disturbance of PSII-OEC increased the production of ROS.
Excessive ROS destroyed OEC performance and chloroplast structure.

a r t i c l e i n f o

a b s t r a c t

Article history: Received 28 October 2018 Received in revised form 8 January 2019 Accepted 23 January 2019 Available online 28 January 2019

Oxygen evolution complex (OEC) in photosystem II (PSII) is sensitive to environmental stressors. However, oxidative damage mechanism in PSII-OEC is still unclear. Here, we investigated photosynthetic performance of PSII, oxidative stress and antioxidant reaction induced by reactive oxygen species (ROS) in a unicellular green alga Chlorella vulgaris (C. vulgaris) under the stress of cetyltrimethylammonium chloride (CTAC). From the changes of chlorophyll fluorescence parameters and PSII activity, it was proved that the electron transport, which occurred initially at the electron donor side of OEC, was disturbed by CTAC. Moreover, a significant decrease of the oxygen evolution rate in OEC (40.95%) while an increase of ROS (50.50%) was obtained after the exposure to 0.6 mg/L CTAC compared to the control (without CTAC), confirming that more oxygen transferred to ROS under the stress. Furthermore, the increased ROS in chloroplast and the structural destruction in thylakoid membrane were observed, respectively. These results proved that oxidative damage mechanism in PSII-OEC is mainly through the reduction of oxygen evolution and the production of excessive ROS, thus leading to the destruction of OEC performance and chloroplast structure. © 2019 Published by Elsevier Ltd.

Handling Editor: A. Gies Keywords: Oxygen evolution activity Reactive oxygen species Oxidative damage Photosystem II Surfactant Algae

1. Introduction Photosynthesis is fundamental to the cellular metabolic process which regulates the survival of primary photosynthesis organisms.

* Corresponding author. E-mail address: [email protected] (F. Ge). https://doi.org/10.1016/j.chemosphere.2019.01.135 0045-6535/© 2019 Published by Elsevier Ltd.

In this process, the light energy captured by pigment molecules (e.g. chlorophyll a, chlorophyll b and carotenoid) can be used to aid electron transfer in photosystem II (PSII) and photosystem I (PSI) (photochemical quenching), dissipated as heat (non-photochemical quenching) or re-released at a slightly longer wavelength (chlorophyll fluorescence) (Salleh and Mcminn, 2011; Melis, 2012; Zhang et al., 2017b). Nevertheless, the photosynthesis performance

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in algal species was extremely sensitive to stressors, mainly reflected in the reduced light capture ability of PSII, weakened reaction center activity, disrupted water splitting process of OEC and damaged thylakoid membrane structure, etc. (Balasaraswathi et al., 2017; Zhang et al., 2018). Therefore, PSII is the most sensitive evaluation indicator in response to stressful conditions (Daum et al., 2010). The continuous application of surfactants has resulted in a high toxicity to primary photosynthetic organisms, such as algae. However, most of the current studies have merely focused on the algal photosynthesis performance (Yamamoto et al., 2006; Brimfield et al., 2012). Also, it is not clear that oxidative stress mechanism is explained by OEC performance. As a basic biological process in ecosystem, water splitting is the foremost source of both electrons and molecular oxygen (Yanykin et al., 2010). This process takes place in OEC which contains a Mn4Ca cluster repeatedly oxidized by reaction center pigment P680 (Aksmann et al., 2011; Vogt et al., 2015). Aksmann et al. (2011) have shown that anthracene inhibited the OEC performance of Chlamydomonas reinhardtii which hindered the photosynthetic performance. In addition, the OEC is more sensitive to environment stress than other photosynthetic apparatus. Thus, the oxygen evolution and electron transfer are generally accepted in assessing the activity of PSII-OEC. Previous studies have shown that environmental stress always stimulates the organism to produce a stress response. For example, reactive oxygen species (ROS) have been increasingly produced by chemical stressors, which can poison and kill cells when they defeat defense system (Halliwell and Gutteridge, 2007; Qian et al., 2009). The mechanism of ROS-production is mainly attributed by following: 1) the reduction of carotenoid contents giving rise to the aggregation of protoporphyrin IX in the cells that suffers oxidation by reacting with the oxygen, ultimately causing
the formation of singlet oxygen (1O2) and superoxide radical (O2); (Kilinc et al., 2009). 2) the electrons in electron transfer chains of chloroplasts, mitochondria and plasma membranes seem to target
and activate O2 production by PSII with formation of O2; 3) the fundamental interaction with atoms or molecules in the cell to
produce highly O2, hydroxyl radical (
OH), and hydrogen peroxide (H2O2) (Das and Roychoudhury, 2014; Almeida et al., 2017) Increasing ROS produced by stressors would respond quickly with the cell, causing disorder of cellular mechanisms (Apel and Hirt, 2004; Jahns and Holzwarth, 2012; Reisz et al., 2014). However, to date, few studies have been conducted on the relationship between oxygen evolution and oxidative stress in the presence of chemical stressors. Surfactants, ubiquitous class of organic contaminants in aquatic environments, have recently produced in increasing amounts, and over 1.96 million tons were used in China in 2013 (Ruan et al., 2014). Quaternary ammonium compounds (QACs) are one type of the typical cationic surfactants, which contain at least one hydrophobic long alkyl chain attached to a positively charged nitrogen atom (Zhang et al., 2017a). Due to its special molecular structure, QACs has a certain toxic effect on the growth and reproduction of aquatic organisms, which may lead to changes in the number of biological populations in water, or even change the structure of the ecological community, and eventually lead to the destruction of the ecosystem balance. Our previous studies have showed that cetyltrimethylammonium chloride (CTAC) which showed a relatively obvious inhibition on the growth and physiological activity of C. vulgaris had more toxic effects than other tested QACs (Zhu et al., 2010; Liu et al., 2018). Thus, we selected the CTAC as the representative QACs. Based on the background, this studied investigated the photosynthetic oxidative damage on a unicellular green alga C. vulgaris under the stress of cetyltrimethylammonium chloride (CTAC), a ubiquitous surfactant in aquatic biota. Firstly, we determined the

chlorophyll fluorescence parameters under the stress of CTAC. Additionally, ROS production was also examined by fluorescence probe technology. Lastly, oxygen evolution levels, ROS levels and the activity of antioxidant enzymes were investigated under light and dark conditions, respectively. Through the above investigation, we will try to explore and clarify the oxidative damage mechanisms in PSII-OEC. 2. Materials and methods 2.1. Chemicals Cetyltrimethylammonium chloride (CTAC, >98.0%) was obtained from Nanjing Robiot Co. Ltd., China. 2, 7-Dichlorofluroescein diacetate (H2DCFDA, 97%) was obtained from Shanghai Xibao Co. Ltd., China. N, N-dimethylformamide (N, N-DMF, 99%) was purchased from Tianjin Hengxing Co. Ltd., China. 2, 6-Dichlorophenol indophenol (DCPIP, 99%) and 1, 5-Diphenylcarbazide (DPC, 99%) was purchased from Sigma-Aldrich (Milan, Italy). 2-(Nmorpholino) ethanesulfonic acid (MES, 99%) and 2, 6-dichloro-1, 4-benzoquinone (DCBQ, 99%) were purchased from Shanghai Aladdin Industrial Co. Ltd., China. All other chemical reagents used in this experiment were AR grade and purchased from Shanghai Macklin Biochemical Co. Ltd., China. 2.2. Algae culture medium and growth conditions The unicellular green alga C. vulgaris (FACHB-1068) applied in this study was purchased from the Freshwater Algae Culture Collection at the Institute of Hydrobiology (Chinese Academy of Science, China). It was inoculated in standard OECD medium according to the Organization for Economic Cooperation and Development Guidelines 201 (OECD, 2006; Yu et al., 2013). All algal solutions were pre-cultured in Erlenmeyer flask (250 mL) and kept in a temperature-controlled illumination incubator (HZ-200LG, Wuhan Ruihua Instrument and Experiment Co. Ltd., China) under continuous white fluorescent lights (2500 lux) with a 12 h light/ dark cycle. The temperature was kept at 25 ± 1  C. To prevent the algal cells sedimentation, all samples were shaken thrice a day. The cells were harvested in the logarithmic growth phase and gathered by centrifugation at 4000 rpm for 10 min. After being washed twice with pure water, the algal cells were incubated in 100 mL of OECD medium. Then, CTAC suspensions were added into the test flask with the final measured concentrations of 0, 0.1, 0.2, 0.4 and 0.6 mg/ L, respectively. The primary cells densities were maintained at 1.0  106 cells/mL (Liang et al., 2013; Xu et al., 2019). The test flasks were placed in an incubator for 96 h under the condition in accordance with pre-cultured condition. 2.3. Quantification of pigments and chlorophyll autofluorescence The chlorophyll a (Chla), chlorophyll b (Chlb) and carotenoid (Car) were determined by spectrophotometer (UV-1000, Shanghai AOE Co. Ltd, China). The detailed steps are as follows. The algal harvested in the logarithmic growth phase were transferred to a centrifuge tube, and then centrifuged at 6000 rpm for 10 min. After the supernatant was discarded, 4 mL N, N-dimethylformamide was put into each sample pipe and mixed. All sample tubes were placed in the darkness at 4  C for 24 h to extract the chlorophyll. Extracting solution (3 mL) was taken from each centrifuge pipe and then the value of absorption was measured at 647 nm and 664.5 nm by a spectrophotometer (722S, Shanghai Precision Scientific Instrument Co. Ltd., China) (Liu et al., 2018). The contents of Chla, Chlb and Car were calculated from the following formulae:

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Total contents ðmg=LÞ ¼ 17:9  OD647nm þ 8:08  OD664:5nm (1) Chla contents ðmg=LÞ ¼ 12:7  OD664:5nm  2:79  OD647nm (2) Chlb contents ðmg=LÞ ¼ 20:7  OD664:5nm þ 4:62  OD647nm (3) Caro contents ðmg=LÞ ¼ Total contents  Chla contents  Chlb contents

(4)

Under five level of CTAC stress, the algal cells were harvested by centrifugation (4000 rpm, 10 min) after 30 min, 3 h, 6 h, 12 h, 24 h, 96 h and 168 h culture, and washed twice with PBS (pH 7.2). Algal re-suspension (3 mL) was measured chlorophyll autofluorescence by fluorescence emission spectrum (Hitachi, Japan) excited at 480 nm. 2.4. PSII performance Chlorophyll fluorescence parameters were investigated by pulse amplitude modulated fluorometer (PAM-2000, Heinz Walz GmbH, Effeltrich, Germany) (Cosgrove and Borowitzka, 2006). Chlorophyll fluorescence parameters were calculated according to the formulae. Under the stress of CTAC, the algal cells were darkadapted for 30 min to allow complete oxidation of PSII reaction centers. The minimum fluorescence yield of dark-adapted cells (F0) was measured under light conditions (photosynthetic active radiation (PAR) less than 2 mmol photons/(m2$s)). The maximum fluorescence yield of dark-adapted cells (Fm) was measured after a 0.6 s saturating pulse light. The minimum (Fʹ0) and maximum (Fʹm) fluorescent yields of PSII in the light-adapted state were recorded. According to the fluorescent yields, non-photochemical quenching (NPQ), non-photochemical quenching (qN), the coefficients of photochemical (qP) and relative photosynthetic electron transport rate (rETR) were calculated. They were calculated from the following formulae:

NPQ ¼ Fm  Fm 0




qN ¼ 1  Fm 0  F0 qP ¼ Fm 0  F


 t

Fm 0

(5)


 0

(6)

rETR ¼ Y*PAR*0:84*0:5

5 mM CaCl2, 2 mM EDTA, 10 mM NaCl and 50 mM Tricine (pH 7.5)). After that, the pellet was broken down using a syringe and centrifuged at 1000 g for 1 min to obtain the chloroplast. The hill reaction activity was determined with DCPIP (Wu et al., 2015). In brief, 20 mM DCPIP (2 mL) was put into a pigment solution (terminal concentration, 10 mg chl/mL). After reaction under the illumination of 4000e5000 lux for 10 min, the absorbance value at the wavelength of 590 nm was detected. It is noteworthy that the reaction of chlorophyll solution with the buffer solution was set as control. Diluted samples were incubated for 3 h with CTAC at concentrations of 0, 0.1, 0.2, 0.4 and 0.6 mg/L on ice and in the darkness in the presence or absence of an electron donor DPC (10 mM) and an electron acceptor K3[Fe(CN)6] (0.6 mM). In vivo, using chloramphenicol (20 mg/mL) that inhibits the PSII repair though inhibiting the synthesis of the D1 protein. After CTAC treatment, samples were gathered by centrifugation (6000 rpm, 10 min), resuspended in Buffer A and adjusted to 10 mg chl/mL. PSII reaction center activity and PSII total activity were measured spectrophotometrically in the presence of the artificial electron acceptor DCPIP (10 mM) at 25  C with and without an artificial electron donor DPC (100 mM), respectively, using a UV-vis spectrophotometer (Cary 300, Varian). Illuminating by LED 2500 lux of light intensity and monitoring the activity by UV-vis spectrophotometer at 600 nm (Zavafer et al., 2015). 2.6. Oxygen evolution activity The photosynthetic oxygen evolution was investigated by a Clark-type oxygen electrode (Chlorolab 1, Hansatech Ltd., UK), companied with a muddler and thermoregulatory by circulating water. In detail, the algal cells (25 mg chl/mL) were treated with CTAC in a buffer comprising 20 mM MES-NaOH (pH 6.3), 1 mM NaCl, 0.5 mM MgCl2, 0.35 mM DCPIP. K3[Fe(CN)6] was considered as a PSII electron acceptor. The mixture solution was illuminated with saturating light (2000 lux) and the rates of oxygen evolved were determined using an oxygen electrode at 25  C. Besides, we also determined the oxygen evolution under the conditions of saturating light (0 Lux) and the temperature was 25  C (in darkness). The samples to be tested were wrapped in tin foil and placed in an incubator at a constant temperature of 25  C for the determination of oxygen release and ROS activity. 2.7. ROS production

Fm  F0

Fm 0  F0

661

0


(7) (8)

2.5. PSII activity The algal cells were collected to extract the chloroplast. In detail, the algal cells were harvested in the logarithmic growth phase and collected by centrifugation at 5000 g for 5 min, then treated by removing supernatant and adding 10 mL enzyme solution (0.5 M mannitol, 0.2% bovine serum albumin (BSA), 50 mM 2-(N-morpholino) ethanesulfonic acid (MES) (pH 5.8), 2% cellulose and 2% macerozyme) (Deng et al., 2013). The density of algal cells was kept in 1.0  106 cells/mL. The solution was shaken in the dark with 60e70 rpm for 24 h at 37  C. Then, it was filtered with a 400-mesh cell strainer and then centrifuged at 120 g for 5 min. The cells were collected and kept suspended in a buffer solution (0.33 M mannitol,

The ROS production was investigated by microplate reader (iMark Bio-Rad, American). Algal cells (in the logarithmic phase) were cleaned three times with 400 mL of 10 mM Na2EDTA solution, and then delivered to a 96-well black plate containing 250 mL of 5% H2DCFDA in 10% Hoagland solution. All samples were saved in the darkness (light intensity: 2500 lux, 25  C) for 30 min. The production of ROS was determined by microplate reader at 530 nm with an excitation wavelength of 485 nm. The CAT and SOD enzyme activity determined by kits (Kemin Jiangsu, China). 20 mL algal cells were gathered by centrifugation at 4000 rpm for 10 min, and then added in pre-cooled PBS (pH 7.6). Then they were broken by ultrasonic cell disruptor in ice-bath, and then centrifuged at 4000 g for 10 min. The activity of SOD was determined according to Alexander and Chridtine (2000). In addition, CAT activity was determined according to Jiang et al. (2017). 2.8. Fluorescence microscope images After culture for 96 h, algal cells (in logarithmic growth phase) were washed with sterile water three times and then delivered into a 1.5 mL pipe. 10 mL 0.5 mM H2DCFDA dye and 1 mL 10% Hoagland's

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solution were subsequently added in the tube. All sample tubes incubated for 30 min in the darkness. After incubation, the cells were transferred into 5 mL 10% sucrose for 20 min and then dehydrated in 20% sucrose for 20 min. Fluorescence measurements were performed with a fluorescence microscope with a CCD camera (Nikon ECLIPSE Ti2, Tokyo Japan).

2.9. Cell ultrastructure Algal cells exposed in CTAC in logarithmic growth phase were collected. The cells were centrifuged at 4000 rpm for 10 min, then cleaned twice. Sample slices were dehydrated successively through a series of alcohol solutions of 30%, 50%, 70%, 90%, and 100% for 10 min, then embedded in Epon 812 resins and polymerized in the oven of 70  C for 24 h. Samples were prepared by a micro-tome (Leica EM UC6, Leica, Wetzlar, Germany), and stained with uranyl acetate and lead citrate. Algal cells were cut into slices with the thickness of 70 nm by ultramicrotome (Leica EM UC6, Leica, Germany) and observed by transmission electron microscopy (TEM, JEM-2100EX, JEOL, Japan).

2.10. Statistical analysis All the experiments were set as three replications and the data were mean ± standard deviation. Figures were analyzed by oneway analysis of variance and Duncan's new complex range test. Points denoted by different lower case letters and capital letters on each curve differ significantly at P < 0.05 and at P < 0.01, respectively.

3. Results 3.1. Disturbance of photosynthetic performance in PSII induced by CTAC As shown in Fig S1, the pigments of photosynthetic containing Chl a, Chl b and Car significantly declined after the 96 h exposure of 0.6 mg/L CTAC (P < 0.05). Additionally, Chla/b ratio in algal cells declined by 35.02% of the control and Chla/car ratio was only 27.01% of the control. However, there were insignificant changes between the sample treated with 0.1 mg/L CTAC and the control (0 mg/L CTAC). Also, it concurrently induced changes in cells Chla/b ratio and Chla/Car ratio. The variations of photosynthetic parameters are illustrated in Fig. 1. As shown in Fig. 1A, there was insignificant difference between the control group (0 mg/L CTAC) and different CTAC-treated groups after 24 h culture. However, with the extending cultivation time, rETR significantly declined with the increasing CTAC concentration. When exposed to 0.6 mg/L of CTAC, the rETR decreased by 39.25% at the 96 h and 46.01% on the 168 h compared with the control. Nevertheless, there was no significant difference at the low concentration of CTAC (0.1 mg/L) (P > 0.05). In addition, the values of qP (in Fig. 1B) also markedly decreased at concentrations of CTAC higher than 0.2 mg/L compared with the control. The maximum inhibition efficiency of qP was 25.65%, 36.90% and 49.26% on the 24, 96 and 168 h, respectively. Whereas, the production of qP displayed an almost insignificant difference when the CTAC concentration was less than 0.2 mg/L (P > 0.05). Meanwhile, NPQ of PSII was significantly increased under the stress of CTAC (Fig. 1C, P < 0.05), accompanied by the increase of qN (Fig. 1D). Similar to rETR, there was an insignificant difference between the control group and the different CTAC-treated groups on the 24 h. With the exposure

Fig. 1. Effect of CTAC on chlorophyll fluorescence parameters of C. vulgaris. (A) rETR: relative photosynthetic electron transport rate. (B) qP: coefficient of photochemical quenching. (C) NPQ: non-photochemical quenching. (D) qN: coefficient of non-photochemical quenching. Exposure time was 168 h. The data was presented as mean ± standard deviation. Error bars indicate the standard deviations (n ¼ 3). Points denoted by different lower case letters and capital letters on each curve differ significantly at P < 0.05 and at P < 0.01, respectively.

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concentration of CTAC from 0.1 to 0.6 mg/L, the NPQ was significantly increased from 0.061 to 0.252 on the 96 h compared to the control group, meanwhile, it was increased from 0.194 to 0.392 on the 168 h. To estimate the algal cell reactions exposed to CTAC, chlorophyll autofluorescence of C. vulgaris were scanned at 684 nm and 735 nm during the 96 h cultivation respectively (Table 1). It was found that, the chlorophyll autofluorescence emission intensity both enhanced at the two different wavelength by extending the cultivation time. When CTAC concentrations ranged from 0 to 0.6 mg/L, the chlorophyll autofluorescence emission intensity gradually increased from 9.73 to 11.91 a.u. at 684 nm, accompanied by an increase from 1.06 to 1.12 a.u. at 735 nm on the 30 min (Fig. S2). Then, the electron donor DPC and electron acceptor K3[Fe(CN)6] were added to detect the total activity of PSII and the activity of the reaction center upon the exposure of CTAC. As shown in Fig. 2, in the absence of CTAC, the addition of electron donor and acceptor had no obvious effect on the total activity of PSII on the 96 h. However, with the exposure of CTAC at 0.2 mg/L and 0.6 mg/L, the total activity of PSII decreased to 39.55% and 25.26% of the initial activity in the absence of DPC, respectively, while slightly increased to 41.10% and 28.15% after the addition of the electron donor DPC. Furthermore, the co-addition of the electron donor and acceptor did not significantly enhance the total activity of PSII, with the value increased to 45.54% and 30.41% at 0.2 mg/L and 0.6 mg/L CTAC, respectively. Diversely, the co-addition of electron donor and acceptor significantly affected the activity of the PSII reaction center. With the exposure concentration of CTAC at 0.2 mg/L and 0.6 mg/L, the activity of PSII reaction center decreased to 56.02% and 49.46% of the initial activity in the presence of DPC, respectively, while significantly increased to 75.71% and 68.45% after the addition of the electron acceptor K3[Fe(CN)6].

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which increased from 55.24% to 96.19% with CTAC ranged from 0 to 0.6 mg/L. However, there was no significant difference under dark conditions (P > 0.05). Meanwhile, we found that the catalase activity of the experimental group treated with 0.6 mg/L CTAC was 57.94% of the control group under light conditions, and the superoxide dismutase activity was 58.37% of the control group (Fig. S4). Similarly, there was almost no change of enzyme activity under dark conditions. The H2-DCFDA stained cells upon visualization under a fluorescent microscope showed the emission of green fluorescence because of dichlorofluorescein (DCF) formation through oxidation by CTAC-induced ROS. As shown in Fig. 4, there was a slight ROS production in the control group, whereas, stronger fluorescence signals were displayed in the CTAC-treated samples (Fig. 4B1 and C1). As shown in Fig. 4B2 and C2, most of the red autofluorescence of chlorophyll were overlapped by the green fluorescence. 3.4. Chloroplast structure damaged by CTAC Alterations in cellular morphology and chloroplast ultrastructure were further studied by TEM. The size of algal cells reduced with the elevated exposure concentration of CTAC (Fig. 5). Moreover, a phenomenon of plasmolysis and damage membrane was observed in the presence of CTAC stressor. Furthermore, the cell organelle damage level was induced by CTAC. The cells in the control group demonstrated a normal healthy chloroplast thylakoids membrane with an extended shape (Fig. 5D), which was the main site for photosynthetic reactions. Chloroplasts were partly jammed with complete thylakoids in the presence of 0.2 mg/L CTAC, while chloroplasts of 0.6 mg/L CTAC-treated group demonstrated loss of membrane completeness with misrepresented thylakoids. As a result, the empty gaps were filled inside the membrane.

3.2. Oxygen evolution activity inhibited by CTAC 4. Discussion The oxygen evolution rates in C. vulgaris exposed at the tested CTAC concentrations under light or dark conditions were investigated. As plotted in Fig. 3, with the elevated exposure concentration of CTAC, the oxygen evolution rate declined from 505.67 to 250.33 mmol O2/(mg$chl$h) under light conditions. However, there was little difference between CTAC-treated sample and the control sample and under dark conditions, which just varied from 310.01 to 300.04 O2/(mg$chl$h). 3.3. ROS production induced by CTAC ROS production in the presence of CTAC stressor was significantly higher than that of the control under light conditions (Fig. 3),

Algae, as important primary producers, play vital roles in environmental protection and nutrient cycling in both aquatic and terrestrial ecosystems. Thus, they are often regarded as a model organism to assess the ecological risk. Surfactants have been reported to negatively impact on algae. Nevertheless, although most of the studies have focused on cultivating inhibition, the photosynthesis performance and the accompanying oxidative damage in algae are still poorly examined, and the oxidative stress mechanism by OEC performance remains unknown. Therefore, in this study, we investigated the oxidative damage mechanisms of in C. vulgaris under the stress of CTAC by evaluating disturbance with photosynthesis and ROS production.

Table 1 Fluorescence spectrum analysis of Chlorella vulgaris PSII system under 480 nm (EM Wavelength). EX Wavelength (nm)

CTAC contents (mg/L)

Fluorescence intensity (a.u) 30 min

3h

6h

12 h

24 h

96 h

684

0 0.1 0.2 0.4 0.6

11.91 12.35 11.23* 10.89* 9.77*

12.43 12.09 11.88* 10.89* 10.08*

12.46 11.54 10.04* 7.60* 7.17*

17.60 17.14 14.60* 9.72* 9.39*

20.14 19.05* 16.37* 15.62* 12.52*

31.35 29.37* 22.18* 19.47* 17.01*

735

0 0.1 0.2 0.4 0.6

1.29 1.32 1.21 1.16 1.06

1.35 1.32 1.27 1.18* 1.12*

1.25 1.17 1.05* 0.82* 0.79*

1.70 1.65 1.46* 1.04* 1.03*

2.06 1.94 1.89* 1.73* 1.52*

3.91 3.03* 2.52* 2.04* 1.82*

* means P < 0.05 compared to control..

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Fig. 2. Variations of PSII total activity (A, B, C) and PSII reaction center activity (OEC activity) (A1, B1, C1) in C. vulgaris upon the exposure of CTAC. PSII samples were exposed to different concentrations of CTAC in the presence of electron acceptor K3[Fe(CN)6] with or without electron donor DPC. Values are mean ± SD of three separate experiments. Exposure time was 96 h. The data was presented as mean ± standard deviation. Error bars indicate the standard deviations (n ¼ 3). Points denoted by different lower case letters and capital letters on each curve differ significantly at P < 0.05 and at P < 0.01, respectively.

Fig. 3. The relationship of oxygen evolution and ROS production under the stress of CTAC. (A) Variations of oxygen evolution (Blue bar) and ROS production (Orange line) influenced by CTAC with light. Exposure time was 96 h. The data was presented as mean ± standard deviation. Error bars indicate the standard deviations (n ¼ 3). Points denoted by different lower case letters and capital letters on each curve differ significantly at P < 0.05 and at P < 0.01, respectively. (B) Variations of oxygen evolution (Blue bar) and ROS production (Orange line) influenced by CTAC under dark. (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.)

Currently, chlorophyll fluorescence technology as a noninvasive and non-destructive assay has been widely applied in physiological and toxicological studies to detect photosynthetic responses of algae. The chlorophyll fluorescence parameters such as F0, Fm, and Fm’ are frequently used to evaluate the PSII activity due to their sensitive towards various stresses (Salleh and Mcminn, 2011; Herlory et al., 2013). In our investigations, these parameters in CTAC-treated groups had obviously decreased, indicating that CTAC resulted in the impairment of PSII function. This destruction may be connected either to a decrease of the electrons transfer chain or the electron consumption requirement of the CalvinBenson cycle. As expected, the reduction of rETR confirmed that the electron transport process was inhibited by CTAC. Also, this impact on electron transport was associated with the PSII-OEC

electron donor site. In our study, the addition of electron donor DPC and electron accepter K3[Fe(CN)6] did not alleviate the decrease of PSII total activity under the stress of CTAC, whereas alleviated the decrease of PSII reaction center activity. Thus, this confirmed that the inhibition of the electron transfer by CTAC was mainly located in PSII-OEC. It is well known that OEC is a complex protein composed of a Mn4Ca cluster, which functions in initial electron transfer and oxygen evolution. However, the OEC activity was often disturbed by stressors, thus inhibiting the electron transport to the reaction center pigment P680þ. To alleviate the disturbance of electron transport caused by PSIIOEC impairment, algae have developed a protective mechanism of NPQ under various stresses. Since NPQ demonstrates that the excess energy in algal chloroplast can be dissipated in the form of

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Fig. 4. Fluorescence microscope images of C. vulgaris with CTAC. The red fluorescence corresponds to the chloroplast (AeC). The green fluorescence indicates the location of reactive oxygen species (ROS) (A1-C1). The merged fluorescence indicates the relationship between chloroplast and ROS (A2-C2). (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.)

Fig. 5. Effect of CTAC on ultrastructure of C. vulgaris and chloroplasts. (A) Ultrastructure of C. vulgaris cells treated with 0 mg/L CTAC. (B) Ultrastructure of C. vulgaris cells treated with 0.2 mg/L CTAC. (C) Ultrastructure of C. vulgaris cells treated with 0.6 mg/L CTAC. (D) Ultrastructure of thylakoid membranes treated with 0 mg/L CTAC. (E) Ultrastructure of thylakoid membranes treated with 0.2 mg/L CTAC. (F) Ultrastructure of thylakoid membranes treated with 0.6 mg/L CTAC.

heat and fluorescence, a high value of NPQ indicated a high photo protective capacity. In this study, increased value of NPQ was also detected with the exposure of CTAC, indicating that CTAC induced protective mechanism of NPQ in C. vulgaris. Moreover, this increase in NPQ was accompanied by an increase in qN and a decrease in qP, thus indicating that the fluorescence quenching of PSII in C. vulgaris was enhanced under the stress of CTAC. The results were consistent with the changes of chlorophyll autofluorescence. The decline of qP was caused by the reduction of Chla and carotenoids contents, in which Chla is responsible for fluorescence emission and carotenoids are involved in energy capture. According to a previous study, the decline of pigment contents will reduce the regulatory capacity of energy dissipation by NPQ and increase the ROS production (Kilinc et al., 2009; Jaspers and Kangasj€ arvi, 2010; Jiang et al., 2014). ROS, such as 1O2, O2
,
OH, and H2O2 are crucial signal transduction cascades which could function in several biological

processes such as photosynthesis, photorespiration, mitochondrial, protein oxidation. Nevertheless, excess ROS induced by contaminants will cause cellular oxidative damage, thus it has been frequently used to assess the toxicity of contaminants in algal species (Nestler et al., 2012; Cheloni and Slaveykova, 2013). As expected, ROS production, upon treatment from 0.1 mg/L up to 0.6 mg/L with CTAC, was significantly higher than that of the control under light conditions. Previous studies demonstrated that the ‘electron leakage’ in electron transport chains (ETCs) of chloroplasts, mitochondria and plasma membrane was the natural source of ROS and contributor to oxidative stress in algae. In normal physiological conditions, 1e5% of electrons can be ‘lost’ in ETCs, while it seems to be leaked out more under environmental stressors (Jiang et al., 2017). Furthermore, we observed that the green fluorescence of ROS overlapped with the red autofluorescence of chloroplast under the stress of CTAC, indicating that

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ROS accumulated mainly in chloroplast. Chloroplast was reported to contribute more ROS in comparison with mitochondria, peroxi€rvi, somes and apoplast. Further, studies (Jaspers and Kangasja 2010) indicated that the major production sites of ROS are located in PSII of thylakoids, because there exist compounds in PSII which
stimulated the generation of active ROS, such as 1O2 and O2-(Asada et al., 2006; Knauert et al., 2008). According to results from chlorophyll parameters and PSII activity, OEC performance in PSII confirms to be related to the ROS production. Previous studies demonstrated that the disruption of OEC performance will prevent PSII photochemistry reaction and then consequently inhibit photosynthetic electron transport. In addition, the alteration in water splitting process performed by OEC can lead to additional ROS production under environment stress. Oxygen, as one of the most important products in the water splitting process, has been frequently used to reflect the OEC activity. We found that oxygen evolution in OEC was significantly inhibited by CTAC in light conditions, indicating the reduction of OEC activity. Nevertheless, there was no significant difference between the control and CTACtreated samples in darkness. The tread of oxygen evolution was consistent with the results of ROS production, confirming that the oxygen evolution of OEC is closely related to the accumulation of free radicals. Previous studies reported that oxygen could be acti
vated to H2O2 and O2 by electron leaked out from ETCs of PSII.
Interaction between H2O2 and O2 can create
OH and 1O2 which are far more destructive to site of origin. It can be explained that the reduction of oxygen evolution was mainly attributed to their activation into ROS under the CTAC stress (Fig. 6). It has been reported that organisms have produced a variety of antioxidant defense mechanisms to counteract the production of ROS, including cleaning oxidative stress such as catalase (CAT) and superoxide dismutase (SOD) (Apel and Hirt, 2004; Halliwell and Gutteridge, 2007). SOD is a valuable scavenger of the antioxidant
system that converts O2 into H2O2 and H2O, it being considered the first line of defense against ROS. CAT plays an important role in oxidative protection by eliminating H2O2, producing H2O and O2 (Abogadallah, 2010). According to our findings, CAT activity and SOD activity were enhanced under CTAC stress in the light, indicating that the antioxidant defense mechanisms were activated in the algae. Once the capacity of antioxidant defense system cannot resist the production of ROS, increasing of ROS can give rise to oxidative stress and consequently induce oxidative damage to necessary proteins and lipid portions in the thylakoid membrane of chloroplasts (Jahns and Holzwarth, 2012). Based on the ultrastructure of C. vulgaris, the chloroplast structure was damaged and

Fig. 6. Mechanism scheme of PSII-OEC performance and oxidative damage induced by CTAC.

thylakoid membranes were rearranged under CTAC stress. Moreover, large spaces between the pairs of stacked thylakoid membranes were observed with the increased CTAC concentration. €rvi (2010) have shown that the oxygen conJaspers and Kangasja centration in chloroplasts by photosynthesis was higher than that of other subcellular organelles, most likely to produce reactive oxygen species. The thylakoid membrane in the chloroplast is rich in unsaturated fatty acid components, and the double bonds contained in the unsaturated fatty acids in the membrane lipid were easily oxidized and decomposed by the attack of free radicals, and finally the membrane system is destroyed. As these results show, chloroplasts are particularly sensitive to membrane peroxidation induced by ROS, which is related to oxygen evolution in PSII-OEC. 5. Conclusions In this study, through investigating photosynthetic performance of OEC, oxidative stress and antioxidant reaction induced by ROS, we proved that oxidative damage mechanism in PSII-OEC under the stress of CTAC is mainly through the reduction of oxygen evolution and the production of excessive ROS, thus leading to the destruction of OEC performance and chloroplast structure. These research results will provide instructive suggestion in deep understanding the oxidative damage mechanism of organism under environmental stress. Acknowledgements This work was financially supported by the National Natural Science Foundation of China (No. 21577117). Appendix A. Supplementary data Supplementary data to this article can be found online at https://doi.org/10.1016/j.chemosphere.2019.01.135. References Abogadallah, G.M., 2010. Antioxidative defense under salt stress. Plant Signal. Behav. 5, 369e374. Aksmann, A., Shutova, T., Samuelsson, G., Tukaj, Z., 2011. The mechanism of anthracene interaction with photosynthetic apparatus: A study using intact cells, thylakoid membranes and PS II complexes isolated from Chlamydomonas reinhardtii. Aquat. Toxicol. 104, 205e210. Almeida, A.C., Gomes, T., Langford, K., Thomas, K.V., Tollefsen, K.E., 2017. Oxidative stress in the algae Chlamydomonas reinhardtii exposed to biocides. Aquat. Toxicol. 189, 50e59. Apel, K., Hirt, H., 2004. Reactive oxygen species: metabolism, oxidative stress, and signal transduction. Annu. Rev. Plant Biol. 55, 373e399. Asada, Y., Tokumoto, M., Aihara, Y., Oku, M., Ishimi, K., Wakayama, T., Miyake, J., Tomiyama, M., Kohno, H., 2006. Hydrogen production by co-cultures of Lactobacillus and a photosynthetic bacterium, Rhodobacter sphaeroides RV. Int. J. Hydrogen Energy 31, 1509e1513. Balasaraswathi, K., Jayaveni, S., Sridevi, J., Sujatha, D., Phebe Aaron, K., Rose, C., 2017. Cr-induced cellular injury and necrosis in Glycine max L.: Biochemical mechanism of oxidative damage in chloroplast. Plant Physiol. Biochem. 118, 653e666. Brimfield, A.A., Soni, S.D., Trimmer, K.A., Zottola, M.A., Sweeney, R.E., Graham, J.S., 2012. Metabolic activation of sulfur mustard leads to oxygen free radical formation. Free Radic. Biol. Med. 52, 811e817. Cheloni, G., Slaveykova, V.I., 2013. Optimization of the C11-BODIPY(581/591) dye for the determination of lipid oxidation in Chlamydomonas reinhardtii by flow cytometry. Cytometry A 83, 952e961. Cosgrove, J., Borowitzka, M., 2006. Applying Pulse Amplitude Modulation (PAM) fluorometry to microalgae suspensions: Stirring potentially impacts fluorescence. Photosynth. Res. 88, 343e350. Das, K., Roychoudhury, A., 2014. Reactive oxygen species (ROS) and response of antioxidants as ROS-scavengers during environmental stress in plants. Front. Environ. Sci. 2, 1e13. Daum, B., Nicastro, D., Austin 2nd, J., McIntosh, J.R., Kuhlbrandt, W., 2010. Arrangement of photosystem II and ATP synthase in chloroplast membranes of spinach and pea. Plant Cell 22, 1299e1312. Deng, C., Zhang, D., Pan, X., Chang, F., Wang, S., 2013. Toxic effects of mercury on PSI

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