Yellow clay modulates carbohydrate and glutathione responses in the harmful dinoflagellate Cochlodinium polykrikoides and leads to sedimentation

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ScienceDirect European Journal of Protistology 71 (2019) 125642

Yellow clay modulates carbohydrate and glutathione responses in the harmful dinoflagellate Cochlodinium polykrikoides and leads to sedimentation Jang-Seu Kia,∗ , Vinitha Ebenezera , Weol-Ae Limb a b

Department of Biotechnology, Sangmyung University, Seoul 03016, South Korea Ocean Climate and Ecology Research Division, National Institute of Fisheries Science (NIFS), Busan 46083, South Korea

Received 15 July 2019; received in revised form 11 September 2019; accepted 27 September 2019 Available online 7 October 2019

Abstract The marine dinoflagellate Cochlodinium polykrikoides is a harmful algal bloom (HAB) species that severely impacts the environment and causes huge economic losses. Yellow clay (YC), considered to be a non-toxic and naturally-occurring material, represents an important step towards the direct control of HABs. In the present study, we evaluated the physiological and biochemical effects of YC on C. polykrikoides after exposures of up to 72 h. We observed little physiological changes in growth rate, chlorophyll a, lipid peroxidation, antioxidant enzymatic activities of superoxide dismutase and catalase, and activity of alkaline phosphatase after exposure to YC. Interestingly, YC significantly increased total carbohydrate and glutathione levels, affecting the physiology of the cells. These results indicate that total carbohydrate content may play an important role in cell–clay aggregation and it could be the main underlying mechanism that mitigates HAB cells via sedimentation. © 2019 Elsevier GmbH. All rights reserved.

Keywords: Antioxidant enzyme activity; Chlorophyll autofluorescence; Cochlodinium polykrikoides; Harmful algal blooms; Yellow clay

Introduction Dinoflagellates are a large group of fresh and marine water microalgae, more than half of which are photosynthetic, and thus play an important role in aquatic ecosystems (Taylor et al., 2008). Dinoflagellate algal blooms cause serious marine disasters that may lead to detrimental environmental effects in oceans, resulting in huge economic losses (Wells et al., 2015). In addition to aquatic ecosystems and the economy, harmful algal blooms (HABs) also adversely impact human health (Willis et al., 2018; Yu et al., 2017). Among HAB

∗ Corresponding

author. E-mail address: [email protected] (J.-S. Ki).

https://doi.org/10.1016/j.ejop.2019.125642 0932-4739/© 2019 Elsevier GmbH. All rights reserved.

species, the dinoflagellate, Cochlodinium polykrikoides Margalef, is ichthyotoxic, causing large-scale finfish mortality via copious amounts of mucous secretion and dissolved oxygen depletion (Kim et al., 1999). In recent years, C. polykrikoides has been found in oceans worldwide, ranging from tropical to temperate zones (Kudela and Gobler, 2012). Cochlodinium polykrikoides HABs, particularly in South Korea, occur almost every year, leading to serious environmental problems and severe economic losses for aquacultures (Park et al., 2013). To help overcome these losses, the Korean government and aquaculture industry spends over US$20 million per year (Jeong et al., 2017). Thus, the development of effective management strategies to control HABs directly, and not just reduce or prevent their effects, is of utmost importance (Anderson, 1997).

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In the recent past, several methods have been used to control and suppress HAB formation. They can mainly be classified into three strategies: chemical, physical, and biological countermeasures (Yang et al., 2015; Yu et al., 2017). However, these methods were not widely used due to their negative ecological impacts, high cost, and poor flexibility (Yu et al., 2017). To overcome these obstacles and control HABs, the development of an alternate approach is important. A promising alternate strategy utilises a non-toxic, natural clay that is inexpensive, easily and readily available, and easy to use. Typical clay is mostly composed of aluminium and iron salts, which produce cationic hydrolysis products that are strongly adsorbed by negative particles, thereby forming hydroxide precipitates (Murray, 2006). Clay also attaches to cationic metals and forms a bridge that absorbs algae, bacteria and other particulates with a net negative surface charge (Hjorth and Jorgensen, 2012). As such, clay sprayed over water surfaces to control HABs leads to the flocculation of clay particles with algal cells and entrainment of algal cells, which eventually leads to the formation of larger aggregates that settle down quickly (Sengco and Anderson, 2004). This technique has proved effective in Japan and South Korea in several HAB trials (Yu et al., 2017). Furthermore, the technique was also implemented in Korea by spraying 60,000 tons of yellow clay (YC) over a coastal aquaculture sector to control and treat red tides caused by Cochlodinium growth (Lee et al., 2013). Reactive oxygen species (ROS) are generally induced in cells by alteration in various external stress factors such as biocides, light, temperature, and pH (Sharma et al., 2012). Increased ROS productivity in cells can cause several deliterious effects such as peroxidation of lipids, oxidation of proteins, and damage to nucleic acids (Ebenezer and Ki, 2014). However, cells employ an eclectic range of defence mechanisms that include enzymatic and nonenzymatic mechanisms involving glutathione, tocopherols, phenolics, carotenoids, and flavonoids (Sathasivam et al., 2018). These two defence mechanisms efficiently reduce cellular oxidative damage by scavenging or removing excess ROS (Sathasivam and Ki, 2018; Sharma et al., 2012). Within cells, the superoxide dismutase (SOD) enzyme constitutes the first component of the antioxidant defence system (Alscher et al., 2002). Other antioxidant enzymes, such as catalase (CAT), peroxidase (POD), and/or glutathione peroxidase (GPx), have also been shown to reduce oxidative stress in microalgae (Ebenezer et al., 2014; Guo et al., 2016) and terrestrial plants (Ahmad et al., 2010). Most studies regarding algicide clay focus on the physical and chemical properties, mitigation efficiency, and/or environmental toxic effects (Beaulieu et al., 2005; Choi et al., 1998; Sengco et al., 2001; Yu et al., 1995). Sedimented cells after clay flocculation have also been thoroughly studied and more information regarding them is available (Sengco et al., 2001; Sun and Choi, 2004). However, biochemical variations in the harmful dinoflagellate, C. polykrikoides, after exposure to YC have not yet been studied sufficiently, until now. Hence,

in the present study, we evaluated the efficiency of YC on the cellular responses of C. polykrikoides, with emphasis on cell number, pigments, total carbohydrate content, and antioxidant enzymatic activities. This study aims to increase our understanding of the role YC plays with respect to harmful algal cells.

Material and methods Cell culture, growth conditions and maintenance A strain (Cp-01) of Cochlodinium polykrikoides was obtained from the National Institute of Fisheries Science (NIFS), Korea, cultured in f/2 medium (Guillard and Ryther, 1962), and maintained at 20 ◦ C using a 12:12-h light/dark cycle with a photon flux density of 65 ␮mol photons m−2 s−1 . YC-treated experiments were performed triplicate in 300 mL flask, using exponential phase cells.

Source and treatment of yellow clay (YC) The YC was obtained from the South Sea Fisheries Research Institute, Korea. The composition of the clay is as follows: Si, 48%; Al, 35%; Fe, 11%; other, 6% (Kim, 1999). Prior to start of experiment, the clay was dried in a hot air oven to remove moisture, then weighed and packed into small aluminum foil pouches. After then, it was dry heat sterilized (110 ◦ C for 24 h) in the hot air oven, cooled to room temperature, and then added to culture flasks containing exponential growth phase Cochlodinium polykrikoides culture. Previous study reported that the exposure of clay in the concentration of 10 g L−1 effectively controlled dinoflagellate growth and the HABs (Sun and Choi, 2004). However, in the present study, we had exposed C. polykrikoides to YC up to 10 times less than the previous study concentration, i.e., 0.01, 0.05, 0.10, 0.50, and 1.00 g L−1 , and the cells were harvested at different time intervals 0, 6, 12, 24, and 72 h. The initial cell concentration was 1 × 106 cells mL−1 .

Cell count and pigment analysis Cell counts in each test flask were determined using a plankton-counting chamber (HMA-S6117, MatsunamiGlass, Japan) and were plotted against the exposure times. Chlorophyll a (Chl a) was measured by using 10 mL of samples cultured at different time intervals. The pigments were extracted with 90% acetone after overnight incubation in the dark. The supernatants extracted were measured using a DU730 Life Science UV/vis spectrophotometer (Beckman Coulter, Inc., Fullerton, CA). The Chl a concentration was estimated according to Parsons et al. (1984). In addition, chlorophyll autofluorescence (CAF) was measured using a fluorescent microscope (Axioscope, Carl Zeiss, Oberkochen, Germany) at 400× magnifications. UV dichoric

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(G365/395–488 nm) source was used for the excitation and the emission was collected by setting the detection bandwidth between 630 and 750 nm. Digital image analysis was performed using the image analysis software ImageJ 1.29 (NIH, Bethesda, MD).

Estimation of lipid peroxidation and antioxidant enzymatic activities Lipid peroxidation was measured according to the method of Heath and Packer (1968). Specifically, the cells were harvested by centrifugation at 4217 × g for 10 min, and then 2 mL of 10% trichloroacetic acid (TCA) was added to the pellet. The cells were kept on ice and homogenized using a Teflon pestle tissue homogenizer. Then the tube was placed in a water bath at 40 ◦ C for 5 min (modified from Soto et al., 2011). The mixture was centrifuged at 4217 × g for 10 min. An equal volume of 0.25% thiobarbituric acid freshly prepared in 10% TCA solution was added to the supernatant. The tube was heated at 95 ◦ C for 30 min in a water bath. The mixture was then cooled to room temperature and centrifuged for 10 min at 4217 × g. The absorbance of the solution was measured at 532 nm. Results were expressed as micromoles of malondialdehyde (MDA) per 104 cells. SOD was assayed according to the method of Beauchamp and Fridovich (1971). Algal cells were harvested by centrifugation at 4217 × g for 10 min, and then 5 mL of 100 mM dihydrogen phosphate buffer was added to the algal cell pellet. The cells were kept on ice and homogenized using a Teflon pestle tissue homogenizer. Then the tube was placed in a water bath at 40 ◦ C for 5 min (modified from Soto et al., 2011). The mixture was centrifuged at 4217 × g for 10 min. To the supernatant, 2.6 mL of the reaction mixture (0.5 M phosphate buffer, 130 mM methionine, 750 ␮M nitroblue tetrazolium, 100 ␮M Na2 EDTA, and 20 ␮M riboflavin) was added. The tubes were incubated in the light (65 ␮mol photons m−2 s−1 ) for 30 min. The absorbance was read at 560 nm. One unit of SOD (U) was defined as the amount of enzyme resulting in 50% inhibition of photochemical reduction of nitroblue tetrazolium (NBT). SOD levels were represented as U per 104 cells (U 104 cells−1 ). CAT is a tetrameric heme-containing enzyme, which catalyzes the breakdown of H2 O2 to water and molecular oxygen (Aebi, 1984). Five milliliters of the algal culture were centrifuged at 4217 × g for 10 min. Two milliliters of extraction buffer (1 M phosphate buffer) were added to the pellet. The cells were homogenized using a Teflon pestle tissue homogenized in ice. The tube was placed in a water bath at 40 ◦ C for 5 min (modified from Soto et al., 2011). The homogenate was centrifuged at 4217 × g for 20 min. To 100 mL of the supernatant, 1.6 mL of 1 M phosphate buffer, 0.2 mL of 0.3% H2 O2 , and 3 mM EDTA were added in a test tube and the mixture was shaken well for 3 min. Enzyme activity was calculated using an extinction coefficient of 0.036 per mM cm−1 and expressed as (unit mg−1 protein). One unit of enzyme is

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the amount necessary to decompose 1 mL of H2 O2 per min at 25 ◦ C. The absorbance of the supernatant was read at 240 nm.

Estimation of alkaline phosphatase activity and total carbohydrates Alkaline phosphatase (APase) was measured using a modified procedure (Gage and Gorham, 1985; Hernández, 1996). 10 mL of culture was harvested by centrifugation at 4217 × g for 10 min, and then 5 mL of 100 mM dihydrogen phosphate buffer was added to the algal cell pellet. The cells were kept on ice and homogenized using a Teflon pestle tissue homogenizer. Then the tube was placed in a water bath at 40 ◦ C for 5 min (modified from Soto et al., 2011). The mixture was centrifuged at 4217 × g for 10 min. To the supernatant, 2 mL of Tris-HCl buffer (pH 8.5) and 2 mL 0.3 mM p-nitronphenylphosphate (p-NPP) was added and incubated at 37 ◦ C for 4 h and 0.1 mL 0.1 M NaOH was added into the mixture after 4 h. The release p-nitrophenol from p-nitronphenylphosphate was determined by absorbance at 410 nm using a DU730 Life Science UV/vis spectrophotometer (Beckman Coulter, Inc., Fullerton, CA) and APA was calculated in nM mL−1 min−1 . Total carbohydrate was estimated in the media, following Dubois et al. (1956). One milliliter sample was reacted with 3 mL of concentrated sulfuric acid (72% w/v) and 1 mL of phenol (5% w/v) in a water bath. The mixtures were incubated for 5 min at 90 ◦ C. The absorbance at 490 nm was then measured using an DU730 Life Science UV/vis spectrophotometer (Beckman Coulter Fullerton, CA). The absorbance measurements were then compared to a standard curve based on glucose.

Estimation of reduced glutathione level Glutathione (GSH) level was measured according to Rathod and Balkrishna (2011). For this analysis, 5 mL aliquots of algal culture were mixed with 2 mL of 5% TCA and centrifuged at 4217 × g for 10 min. The cells were homogenized using a Teflan pestle on ice. The tube was placed in a water bath maintained at 40 ◦ C for 5 min (modified from Soto et al., 2011). To the supernatant, 0.6 mM of 5 ,5 -dithio-bis-2-nitrobenzoic acid (DTNB) in (0.1 mM) phosphate buffer was added. Thiol anions were measured at 412 nm using a DU730 Life Science UV/vis spectrophotometer (Beckman Coulter, Fullerton, CA).

Statistical analysis Statistical analyses were done using the software Instat® (Graphpad Software, Inc., San Diego, CA). The data were analyzed using one-way analysis of variance (ANOVA), followed by the Student–Newman–Keuls multiple comparisons test. P < 0.05 was considered statistically significant, and data are presented as mean ± standard deviations (SD). The corre-

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tory settings. Since the movement of ocean water is more prominent, it is possible that this leads to delays in the settling time of clay particles, thereby facilitating longer exposure times of algal cells to YC and causing stress to these cells (https://www.clearias.com/movements-ocean-waves-tidescurrents/). Stressed algal cells considerably increase their resistance due to mucous secretion (Gala and Giesy, 1991). Perhaps, this allows algal cells to aggregate with clay even more strongly and results in decreased cell concentrations (Kremp, 2001). Finally, cell–clay aggregates sink into deep and dark waters. These could be some of the reasons as to why YC effectively mitigates HABs under environmental conditions. Field surveys in South Korea showed that Cochlodinium polykrikoides blooms did not reoccur for the rest of the season after YC dispersion in the environment (Lee et al., 2013; Sun and Choi, 2004). This suggests that YC can inhibit the growth of C. polykrikoides only by flocculation. However, the substance responsible for the cell–clay aggregation remains undetected.

Effects of YC on morphology and chlorophyll autofluorescence

Fig. 1. Variation in cell number (A) and chlorophyll a levels (B) of Cochlodinium polykrikoides after exposure to different concentrations of yellow clay at different time intervals. Error bars represent ± SD; n = 3. Significant difference as determined by Student–Newman–Keuls test are represented as *P < 0.05 level when compared to control (CK).

lation between cell count and Chl a was tested using Pearson’s correlation coefficient and a Microsoft Excel spreadsheet.

Results and discussion Effects of YC on cell number and chlorophyll a content In the present study, most algal cells stuck to clay particles and then quickly sank to the bottom after YC treatment. However, viable cells easily escaped from the settled clay flocs because they probably did not bind firmly to YC. For this reason, the cell number and Chl a content in the culture exposed to YC was not significantly different from the control after 6, 24 and 72 h exposures (Fig. 1). However, the settling pattern and easy escape of cells was observed to be different in natural environments when compared to labora-

Previously, most studies reported that algal decease after clay treatment was not due to the release of toxic substances (Beaulieu et al., 2005; Choi et al., 1998; Sengco et al., 2001; Yu et al., 1995). Although, our study showed similar cell growth under YC treatment and control conditions, microscopic observation of the damaged cells showed that algal cell morphology was affected after YC exposure (Fig. 2). This included cell size discrepancies, fragmentation of cell wall, loss of some pigment contents, and/or flagellar damage. A similar result was found in other dinoflagellates (Alexandrium tamarense complex and Scrippsiella trochoidea) after being exposed to a clay matrix (Sun and Choi, 2004). In addition, Sengco et al. (2001) reported that algal cell death occurred after direct physical contact with clay at higher concentrations at increased exposure times. Interestingly, flagellar damage and modification in swimming behaviour were demonstrated to be responsible for growth inhibition (Kremp, 2001). Physiological stress or damage was suspected due to the significant decrease in CAF intensity in YC-treated cells (Fig. 2). Perhaps, cells exposed to YC did not function normally in photosynthesis and cell metabolisms caused by cell signaling disorders, cell cycle dysregulation, and/or oxidative stress (Wang et al., 2018b).

Effects of YC on antioxidant enzymatic and alkaline phosphatase activities The effects of YC on the antioxidant enzymatic and alkaline phosphatase activities of C. polykrikoides were evaluated (Fig. 3). As mentioned previously, SOD acts as the first line of defence against ROS (Ebenezer and Ki, 2014; Sathasivam

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Fig. 2. Morphological changes and chlorophyll autofluorescence (CAF) of Cochlodinium polykrikoides; (A and B) represent the control cells; (C and D) represent the cells exposed to yellow clay (0.5 mg mL−1 ), respectively; (E) represents relative CAF levels per Cochlodinium cells of control and yellow clay (YC) treated culture. Arrow represents damage or loss of pigment. Relative CAF level was measured with ImageJ. Error bars represent ± SD. Significant difference as determined by Student–Newman–Keuls test is represented as *P < 0.05 level when compared to control. Scale bar represent 10 ␮m.

et al., 2016; Wang et al., 2019), whereas CAT, POD, and GPx prevent oxidative stress in both higher plants (Ahmad et al., 2010) and microalgae (Sathasivam et al., 2018; Wang et al., 2018a). The present study showed that after 6, 12, 24, and 72 h exposures to different YC concentrations (0.01, 0.05, 0.1, 0.5, and 1.0 g L−1 ), the CAT, MDA, and SOD levels showed little to no difference when compared to the control (Fig. 3A–C). This indicates that short exposures to YC does not majorly affect algal cell antioxidant enzymatic levels. It is likely that YC uses another important mechanism to arrest or inhibit the growth of C. polykrikoides cells.

A previous study reported that YC absorbed nutrients from water columns, which impacted dissolved oxygen concentrations (Yu et al., 1995), thereby causing severe stress to algal cells. APase plays a vital role in internal phosphorus cycle in water and lake sediments (Zhu et al., 2013; Zhu et al., 2016). Moreover, algal and bacterial cells contain the enzyme phosphatase, which is localized on the surface of cells (Zhu et al., 2016). APase also has an external function: it is excreted into the surrounding culture medium depending on the circumjacent phosphorus concentration, especially under phosphorous deficient conditions (Zhu et al., 2016).

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Fig. 3. Variation in enzymatic activity of Cochlodinium polykrikoides after exposure to different concentrations of yellow clay at different time intervals; variations in lipid peroxidation (MDA) activity (A), superoxide dismutase (SOD) activity (B), catalase (CAT) activity (C) and alkaline phosphatase assay (APA) activity (D). Error bars represent ± SD; n = 3. MDA, Malondialdehyde; CK, control.

Hence, APase is more likely to be absorbed by clay (Tietjen and Wetzel, 2003), thereby decreasing its level in the medium and initiating severe competition among the cultured species. However, in our study, after 6, 12, 24, and 72 h time periods, cultures exposed to 0.01, 0.05, 0.1, 0.5, and 1.0 g L−1 concentrations of YC showed little to no difference in APase activity when compared to the control (Fig. 3D), which supported the cell number, Chl a level, and antioxidant enzymatic activity results. Therefore, this indicates that short exposure times (up to 72 h) to YC do not cause nutrient starvation (especially with respect to phosphorus) in medium designed to inhibit the growth of C. polykrikoides. Thus, it can be inferred that YC removes HAB cells only via flocculation.

Effects of YC on total carbohydrate content Polysaccharides are carbohydrates that consist of several sugar molecules bound together (Guo et al., 2017). Algae is known to release a variety of extracellular polymeric substances (EPS), especially polysaccharides, that are vital to their various respective functions (Hokputsa et al., 2003). For example, EPS produced by algal cells has the ability to protect them from toxic substances (Costa et al., 2018). In addition, it has the ability to aggregate soil particles due to its slimy texture and ionic charges, and to act like glue that

sticks to clay and ions, inevitably leading to the aggregation of solid particles (Chenu, 1995). Total carbohydrate contents were measured using dglucose and spectrophotometry; here we found significantly correlation with its different concentrations (R2 = 99.0, P < 0.01). Considering the standard curve, different biomass concentrations of non-treated C. polykrikoides were used to analyse the total carbohydrate content in the cells. The results showed that the total carbohydrate content gradually increased with increasing biomass concentration (Fig. 4A). In addition, the exposure of YC to C. polykrikoides induced a wide range of responses depending on the clay concentration. Exposure to YC for 6 and 12 h time periods at all experimental concentrations (0.01, 0.05, 0.1, 0.5, and 1.0 g L−1 ) showed a gradual increase in total carbohydrate levels (Fig. 4B). Interestingly, total carbohydrate levels considerably increased in the 24 and 72 h cultures; a trend that was associated with increasing YC concentrations (from 0.01 to 1.0 g L−1 ). After 72 h of YC exposure, the total carbohydrate content in C. polykrikoides increased to ∼2 times of that in the unexposed culture. The highest total carbohydrate content was observed in the culture exposed to 1.0 g L−1 of YC, whereas the lowest level was observed at the 0.01 g L−1 concentration. What is interesting from this result is that YC induces stress in C. polykrikoides, which causes the cells to trigger EPS production, leading to cell–clay aggregation and formation of large

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Fig. 5. Variation in reduced glutathione levels of Cochlodinium polykrikoides after exposure to different concentrations of yellow clay at different time intervals. Error bars represent ± SD; n = 3. Significant difference as determined by Student–Newman–Keuls are represented as *P < 0.05; ** P < 0.01 level when compared to control.

Fig. 4. (A) total carbohydrate levels of non-treated Cochlodinium polykrikoides cells at different biomass concentrations; and (B) total carbohydrate levels of C. polykrikoides cells after exposure to different concentrations of yellow clay at different time intervals. The dotted line represents mean values of controls at 6, 12, 24 and 72 h. Error bars represent ± SD; n = 3. Significant difference as determined by Student–Newman–Keuls test are represented as *P < 0.05; ** P < 0.01 level when compared to control (CK).

particles that quickly settle down and further entrain cells during their descent. From this study and previous reports, the fundamental HAB-mitigating mechanism via YC is observed to occur via clay flocculation, leading to cell–clay sedimentation (Lee et al., 2013; Sengco and Anderson, 2004).

Effects of YC on reduced glutathione levels Although a wide range of responses were induced from C. polykrikoides depending on YC concentration, 6 and 12 h exposures to YC at the lowest experimental concentration (i.e. 0.01 g L−1 ) did not show any change in reduced glutathione levels (Fig. 5). However, after 24 and 72 h YC exposures at this concentration, a slight increase was observed. Notably, the reduced glutathione levels were significantly higher in cells exposed to relatively high concentrations of YC (0.05, 0.1, 0.5, and 1.0 g L−1 ) at all exposure

times (6, 12, 24, and 72 h). Reduced glutathione levels increased to ∼3 times in the culture exposed to 1.0 g L−1 of YC after 6 h when compared to the control. The main role of glutathione is to maintain proper cellular function and prevent oxidative stress in cells (Mezzari et al., 2005). It acts as a scavenger of hydroxyl radicals, singlet oxygen, and various electrophiles. Glutathione reductase is an important enzyme that catalyses the reduction of glutathione disulphide to the sulfhydryl form of GSH, which is a life-threatening molecule that counteracts oxidative stress and the balance of the reducing environment of the cell (Deponte, 2013). This enzyme diminishes the oxidized form of the enzyme glutathione peroxidase, which sequentially decreases the level of hydrogen peroxide, a dangerous ROS produced within cells. Therefore, this result shows that oxidative stress may be induced in YCexposed C. polykrikoides cells, which leads to increased GSH levels in the YC-exposed cells.

Implication of YC in environmental treatments In Korea, the most promising and practical approach in mitigating HABs is the addition of YC, which can be backed by field documentation showing the removal of algae from water columns (Anderson, 1997; Sengco and Anderson, 2004). In the present study, YC effectively settled down C. polykrikoides cells at exponential staged culture. Taken together, our favoured explanation regarding their mechanism-of-action involves YC adhering to the algal cell surface and increasing its stickiness in mucilage (by increasing the total carbohydrate content) on most of the cell surface (Sengco and Anderson, 2004), which triggers cell–cell and cell–clay adhesion, leading to sedimentation of flocs, thereby inhibiting growth and inducing algal cell death (Fig. 6). A

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Fig. 6. A systematic diagram of yellow clay mitigates the Harmful Algal Blooms (HABs).

similar mechanism was reported by Pan et al. (2011). YC also mitigates HABs via another mechanism that involves the induction of oxidative stress in organisms, thereby inhibiting the growth of the remaining cells. This might be one reason why cell lysis occurs after YC exposure. The use of YC in natural environments will involve a balance between removing HABs from the water column and reducing the environmental impacts on benthic fauna (Beaulieu et al., 2005). Hence, the use of clay increases the transparency of water, diminishes toxicity, and limits the extent of hypoxic conditions near the sea floor, thereby mitigating the serious outbreaks caused by HABs and measurably increasing the survival of fish (Shirota, 1989). In conclusion, YC dispersal is a promising strategy in controlling HABs to date. Our results clearly show that YC settles down C. polykrikoides by increasing total carbohydrate production and inducing cellular stress and leads to cell to clay aggregation. Both effects demonstrate effective HAB mitigation using YC. In addition, it may be the main underlying mechanism that mitigates HAB cells via sedimentation. Future research needs to be designed to study other benthic influences in the field and analyse the amount of toxin removal by the clay and algae removal during the tide.

Author contributions VE performed the experiments and analyzed the data; JSK conceived and designed project, analyzed the data, and wrote

the manuscript; WAL designed project and analyzed the data. All the authors read and approved the final manuscript.

Acknowledgements We thank Dr. R. Sathasivam for critical comments on the early version of manuscript. This work was supported by the National Research Foundation of Korea Grant funded by the Korean Government (2015M1A5A1041805) and by a grant from the National Institute of Fisheries Science (R2019037), South Korea, funded to J.-S. Ki.

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