Mitigating Biofouling in Cooling Water System: Actibromide® to Combat Perna viridis Infestation and environmental impact

doi:10.21203/rs.3.rs-5121015/v1

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Mitigating Biofouling in Cooling Water System: Actibromide® to Combat Perna viridis Infestation and environmental impact

https://doi.org/10.21203/rs.3.rs-5121015/v1

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Heavy settlement and fouling of green mussels were observed in the process seawater heat exchangers (PSWHX) and conduits at the Madras Atomic Power Station (MAPS), despite the use of a continuous low-dose chlorination (CLDC) regime. This regime involved maintaining total residual oxidant (TRO) levels at 0.2 ± 0.1 mg/L and performing twice-weekly booster dosing at 0.4 ± 0.1 mg/L. To enhance the efficiency of these heat exchangers, supplemental targeted dosing of Actibromide® was considered. The efficacy of this biocide on adult green mussels was evaluated to determine the appropriate in-plant concentrations (0.2, 0.5, and 1.0 mg/L TRO). The results showed 100% mussel mortality after 12 days at 0.2 mg/L, 7 days at 0.5 mg/L, and 4 days at 1.0 mg/L. Actibromide® exerted toxic effects on mussels by generating reactive oxygen species (ROS), which inhibited cellular processes in various tissues, including the gills, mantle, digestive gland, and foot. The highest ROS generation was observed in the digestive gland compared to other tissues. Hydrogen peroxide (H2O2) production increased in a dose-dependent manner under Actibromide® stress, and superoxide dismutase (SOD) and catalase (CAT) activity were highest in the digestive gland. DNA damage, expressed as % tail DNA in a comet assay, indicated that even the lowest dose of Actibromide® (0.2 mg/L) induced significant DNA damage (34%). Additionally, acetylcholinesterase (AChE) activity, a sensitive biomarker for neurotoxic stress, showed reduced activity (80–91%) at all tested biocidal concentrations. This study clearly demonstrates that Actibromide® penetrates green mussels at the cellular level, causing severe damage to the gills and digestive glands, reducing feed consumption, and inducing both neurotoxic and genotoxic effects. Therefore, supplemental targeted dosing of Actibromide® (0.2 to 0.5 mg/L) is recommended for effective green mussel control in PSWHX systems.

Perna viridis

actibromide®

Antioxidative enzymes

Genotoxicity

Neurotoxicity

Biofouling by green mussels in the cooling water system (CWS) of Madras Atomic Power Station (MAPS), located on the South East Coast of India (Lat long: 12.5238° N, 80.1568° E) is more pronounced and possess severe operational problems (Rajagopal et al., 1991a; Venkatesan et al., 2008; Venugopalan et al., 2011; Murthy et al., 2011; Venkatnarayanan, 2018). The power station operates on a continuous low dose chlorination regime (0.2 ± 0.1 mg L^− 1 TRO) and twice-a-week booster dosing (0.4 ± 0.1 mg L^− 1). The green mussel Perna viridis is reported as the most dominant macro-foulant found in the CWS of MAPS (Murthy et al., 2011; Venkatnarayanan, 2018). Heavy fouling by green mussels and barnacles (Amphibalanus reticulatus) induced pressure drop across the sub seabed intake tunnel (Nair et al., 1988), blockage of condenser tubes (Rajagopal et al., 1996) and biofouling loading of 3–14 kg m^− 2 y^− 1 have been observed in the cooling water conduits of this power station (Rajagopal et al., 1991b; Murthy et al., 2011; Venkatnarayanan, 2018). Reports suggests that, the conditions prevailing inside the CWS provides an ideal environment for these organisms to settle and grow due to the availability of nutrients, oxygen, and the removal of wastes by the flowing seawater. (Sasikumar, 1991; Rajagopal et al., 1991a; Satpathy et al., 1994). Blockage of condenser tubes and heavy loading of green mussels has been observed in the inlet conduits and the inlet tube sheet area of the process sea water heat exchangers (PSWHX) at the power station resulting in loss of heat transfer efficiency (Murthy et al., 2011).

Much of the literature emphasizes the use of chlorine, which has been the most commonly employed biocide due to its ease of handling and cost effectiveness. In recent years, alternative biocidal methods have been investigated to overcome the drawbacks associated with chlorine-based disinfection (Bartholomew, 1998; Jenner et al., 1997; Venkatnarayanan, 2017). Chlorine reacts with organic maters that produce chlorine by-products (CBPs), including trihalomethanes (THMs), haloacetic acids (HAAs), haloacetonitriles (ANs), and halophenols (HPhs) (Allonier et al., 1999; BEEMS, 2011; Cui et al., 2022). Chlorine dissociates to hypochlorous acid (HOCl), is a pH dependent (at higher pH the biocidal effects reduces). THMs are the first halogenated by-products, comprising of important compounds such as chloroform, monobromodichloromethane, dibromochloromethane, and bromoform (Jenner et al., 1997). Bromine-based biocides have advantages over chlorine, due to their effectiveness at seawater pH 8.2 and strong oxidation potential than chlorine (Flemming et al., 2009; Jenner et al., 1985).

Active bromide-based biocides are widely used in water treatment processes, including cooling towers, industrial water systems and swimming pools (Bartholomew, 1998). They are chosen for their effectiveness in controlling biofilm formation, preventing the spread of waterborne pathogens, and inhibiting the growth of algae and other microorganisms. However, the oxidants convert bromide to oxidized forms of bromine which subsequently form a variety of brominated compounds. In seawater the bromine concentration is relatively low (65–70 mg L^− 1) compared to chlorine (18,980 mg L^− 1) (Bartholomew, 1998), which can get oxidized by the added chlorine resulting in the formation of hypobromous acid (HOBr) and hypobromite (OBr⁻) ion (Helz et al., 1981; Taylor 2006). Hypobromous acid is likely associated with bromine's lower oxidation potential compared to hypochlorous acid from chlorine, leading to a reduced tendency of THMs formation (Faulkner 1988). Bromine is known to be highly effective in treating biofilms and biofoulings; intermittent application of 13–56% less oxidant and with continuous application of 33–81% less oxidant has been known to kill 50% of the organisms (Liden et al., 1980). Because of their relatively low bond strengths, bromine residuals show lower stability and are expected to decay more quickly (Fisher et al., 1999). Additionally, they are more reactive than chlorine residuals and may be more effective as alternative biocides (Bongers et al., 1978).

Despite using CLDC, the inlet water boxes of process seawater heat exchangers (PSWHX) faced heavy fouling and flow blockage by mussels, making it ineffective for controlling adult green mussel settlement (Nair, 1999). The power plant later resorted to supplement another biocide viz. Actibromide® was dosed upstream of the PSWHX at residual levels of 0.2 ± 0.1 mg L^− 1 for one hour during each eight-hour shift. While subsequent studies and biomonitoring of the CWS indicated a significant reduction in fouling, the remaining fouling load still posed challenges for power plant PSWHX operations, necessitating periodic maintenance and annual shutdowns (Venkatnarayanan, 2018). The findings from the previous study clearly indicate the present chlorination regime is sufficient to keep the main condensers clean, whereas the biocidal regime followed was found to be inadequate for preventing fouling in the conduits and the process heat exchangers (Murthy et al., 2011). Therefore, fine tuning the active bromide dosing from intermittent to continuous regime could be a possible way to overcome this fouling issue faced in the CWS. In order to comply with the environmental discharge regulations and to combat the adult green mussels in PSWHX and its conduits, targeted supplementary dosing of actibromide® has been investigated along with the existing continuous low dose chlorination regime in practice.

In comparison to use of biocides like chlorine and chlorine dioxide, data on the efficacy of actibromide® on adult green mussels and their interaction and response at cellular and physiological levels have not been investigated (Badakumar et al., 2024; Badakumar et al., under communication). Valve closure has been an important strategy for green mussels to overcome unfavorable biocidal dosages in CWS (Venkatnarayanan et al., 2021). However, studies by Badakumar et al., (2024) have shown that even minor ingress of chlorine into the mantle cavity of mussels, induces stress at the cellular levels, resulting in altered physiology and weakening the mussel resulting in gradual mortality in the green mussel. Such studies investigating the mechanistic action of biocides at cellular levels inducing mortality are much needed to understand from an environmental standpoint.

Mortality of marine organisms is influenced by alterations in the aquatic environment, such as temperature (Syafaat et al., 2021), salinity (Zhang et al., 2020), pH; feed (Mohan et al., 2019), oxidizing biocides (Chavan et al., 2016) and by xenobiotic chemicals. Biomarkers serve as sentinel indicators in response to external stress factors. Assessment criteria for the interpretation of activity levels of biomarkers in mussels have been established (ICES, 2011). A broad spectrum of cellular biomarkers has been studied like the oxidative stress induced by environmental pollutants which results in the generation of reactive oxygen species (ROS) (Irani, 2000). Other reactive species include superoxide, hydroxyl and peroxyl radicals and non-radicals like hydrogen peroxide and singlet oxygen (Ames et al., 1993). These have shown to act on intracellular components like DNA, proteins and lipids (Grune, 2000; Ghezzi and Bonetto, 2003). The organismal response to oxidative stress includes generation of antioxidant enzymes like super oxide dismutase (SOD), catalase (CAT) and glutathione peroxidase (GPx) which quench / neutralize the ROS. Neurotoxicity of xenobiotics have been measured using the acetylcholinesterase (AChE) activity (Mora et al., 1999; Ellman et al., 1961). Recently oxidizing biocides like chlorine have also been shown to act on neuronal synapses altering the titer of acetylcholinesterase (AChE) in green mussel affecting their physiological process (Badakumar et al., 2024). In addition to different cellular toxicity mechanisms, induction of genotoxicity by xenobiotic substances has been documented in studies by Almeida et al., (2011) and Chavan et al., (2016), and is widely accepted as a biomonitoring tool in environmental toxicity studies (Bolognesi et al., 2014).

In the present study, the effect of continuous actibromide® dosing (0.2, 0.5 and 1.0 mg L^− 1, TRO) on adult green mussels was studied by conducting a laboratory-based seawater flow through system to determine the time taken for mortality and its effects using biomarkers at molecular and cellular levels which are indicative of general, oxidative stress, genotoxic effects and neuronal stress. Experiments were designed to analyze the changes in the levels of different oxidative stress biomarkers and their quenching enzymes in response to different concentration of actibromide® in adult green mussels. In this study, several tissues of the green mussel were selected for analysis based on their specific functions: the respiratory organ (gills); the digestive gland; the mantle tissue (which plays a role in immunological defence against predators) and the foot tissue (involved in production of byssus threads). These are the major sites of action of xenobiotic and oxyradical generating biotransformation enzymes, as documented by Livingstone et al., (1992). The parameter measured include: (1) ROS and H₂O₂ as oxidative stress markers; (2) activity of SOD and CAT as antioxidant, (3) GSH as a conjugation substrate and free radical scavenger, (4) DNA integrity by doing comet assay and (5) AChE activity as a neuronal effect to the nervous system.

2.1 Collection of green mussels and maintenance of brood stock

Adult green mussels (Perna viridis), size: (30–70) mm were collected from an unpolluted coastal mussel farm at Cuddalore (11.7390^◦ N, 77.7866^◦ E), in Tamil Nadu, India. Post collection; the mussels were shifted to the laboratory and cleaned thoroughly using a brush to remove the epibionts and debris. Cleaned green mussels were transferred to the fibre rein-forced plastic (FRP) tanks with flowing raw seawater and maintained as stocks in a temperature-controlled environment (27.0 ± 1.5°C, pH 8.22 ± 0.5) for acclimation. The mussel stocks were fed daily twice with the algae Isocrysis galbana (obtained from central marine fisheries research centre, Chennai, Tamil Nadu, India) grown with Walne’s medium (Walne, 1976) at a final concentration of 10⁶ cells mL^− 1 along with the algae available in continuous flowing raw seawater. Mussels which were producing good byssal thread and reattachment during the acclimation period (10 days) were considered as healthy and used for the toxicity experiments.

2.2 Experimental setup

The laboratory experimental set up was designed to control the flow of seawater and allow continuous actibromide® dosing throughout the experimental period. The experimental setup comprised of two set-up tanks (each, 25 L capacity); (a) control/ untreated tanks (in triplicate), and (b) experimental actibromide® treatment tanks (6 replicates). In the experimental tanks (6 replicate treatment tanks), one set (i.e., 3 tanks) was used for performing the physiological assays and the other set (i.e. 3 tanks) were used for biomarker assays. The main seawater flow connecting to each of the tanks during the entire experiment was maintained at the rate of 190 ml min^− 1. In treatment tanks, three different concentrations of actibromide® (0.2 mg L^− 1, 0.5 mg L^− 1 and 1.0 mg L^− 1) were maintained for the study. The commercial biocide actibromide® was obtained from M/s Ion exchange India Ltd. The biocide was activated by mixing 1 part of the actibromide® with 3 parts of 12% sodium hypochlorite solution.

Prior to commencement of experiments, biocidal demand of the seawater was ascertained using demand free seawater. A stock concentration of biocide (1:3 ratios of actibromide® and sodium hypochlorine) was prepared and depending on the concentration to be dosed the biocide flow was throtled and adjusted to maintain the required TRO in the respective tanks. The treated tanks were regulated from this biocidal stock tanks with the flow rate fixed at 4.0 mL min^− 1 to obtain the desired residual concentration of actibromide® (0.2, 0.5 and 1.0 mg L^− 1, TRO respectively). The outlet flow in the tanks was maintained at the rate of 120 ml min^− 1. The experimental tanks were measured frequently (once every 4 hours) for the biocide concentration using DPD (diethyl-p-phenylenediamine) no-4 tablets (Lovibond, USA) with a hand-held colorimeter. The results were measured as total residual oxidants (TRO) and confirmed by checking the samples obtained from individual outlets.

For obtaining the required residuals at the outlet of the experimental tanks (0.2–0.5 mg L^− 1, TRO [low dose] and 1.0 mg L^− 1 [shock dose]), the biocide stock was renewed twice a day (once in 12 hours). The control tanks were free of biocide. Post setting up the control and treatment tanks, for the toxicity experiments, 20 no’s adult green mussels (size: 30–60 mm) were placed in each of the tanks for acclimation. Feeding of experimental and control tanks were carried out on a daily basis. During the feeding time the biocide flow was stopped and the tanks containing the green mussels were cleaned thoroughly and fed with the algal diet to minimize the interaction of the treated chemicals with the microalgae (Juhel et al., 2017). The biocide dosing was checked and adjusted thrice daily to maintain the required TRO levels at the outlet. The mortality (if any) and the condition index were checked every day in the respective tanks. Randomly 3 nos of mussels were taken out from both control and treatment tanks at every three-day interval, (i.e., 0th, 3rd, 6th, 9th and 12th) for assessment of physiological response and for biomarker assays.

2.3 Effect of residual actibromide® on physiological metabolism of P. viridis

2.3.1 Physiological response

Mortality was checked periodically in both control and treatment tanks and checked throughout the experimental period. The mussels which were opened their valve fully was recorded as dead. The percentage mortality was calculated as the number of dead mussels recorded in treatment tanks multiplied by the total available organisms in the respective tanks (Li et al., 2012). The condition index (CI) was also checked by selecting 3 individuals of each tank of both control and treatment. For checking the CI, the soft tissues were removed from the shell and dried in an oven at 80°C for 6 h and then weighed. The percent CI value was calculated according to Almeida et al., (2013).

2.3.2 Ammonia

The control and experimental mussels were fed on a micro-algal diet during the experimental period. In the treatment tanks, there were no pseudofaeces production. Meanwhile in the control tanks, the pseudofaeces production ranged from 2.9 to 5.0 mg L^− 1 per day. In the experimental tanks instead of pseudofaeces production there was an increase in ammonia production which was quantified both in experimental and control conditions. Ammonia was estimated in treatment and control tanks by taking 50 mL of water samples from each. The samples were estimated using 95% phenol alcohol solution, 0.5% sodium nitroprusside (1 g sodium nitroprusside in 200 mL distilled H₂O), alkaline solution (100 g trisodium citrate and 5 g sodium hydroxide in 500 mL H₂O), sodium hypochlorite solution and oxidizing solution (100 mL alkaline solution and 25 mL sodium hypochlorite solution. Samples were kept in dark for 1h incubation, and the absorbance was read at 640nm (Grasshoff et al., 2009).

2.3.3 Sample preparation

The enzyme activity was analysed using six randomly selected mussels from both the control and treatment tanks. Using a sterile scalpel, the mussel shells were carefully pried open to expose their internal organs. The gill, mantle, digestive gland and foot (figure.1) tissues were excised (Pal et al., 2012). The collected tissues were thoroughly washed with sterile potassium phosphate buffer (50 mM; pH 7.4) and stored at -80°C until further use. The tissue samples were taken and homogenised with liquid nitrogen in a pre-chilled mortar and pestle (Chavan et al., 2016). The homogenisation was carried out using potassium phosphate buffer (50 mM, pH 7.4). The homogenized extract was centrifuged at 20,000×g for 15 min at 4°C. The supernatant was aliquoted for assessment of biomarker assays and the total protein was also estimated using Bradford method (Bradford, 1976).

2.3.4 Reactive oxygen species

Free radical generation in tissue of exposed green mussels was determined by following the method described in (Gomes et al., 2005) with minor modifications. The protein extracts from the tissue homogenate were used as samples for ROS estimation using the dye DCFDA (2′,7′- Dichlorodihydrofluorescein diacetate). For analysis equal volumes of sample and DCFDA were mixed and incubated in dark for 15 mins. Post incubation the samples were analysed using a 96-well fluorescence microplate reader (Tecan, Germany) at excitation 498 nm and emission with 525 nm. The obtained results were expressed as the fluorescence units (expressed as Arbitrary Units or AU) with respect to control.

2.3.5 Antioxidant enzymes

The activity of total SOD was determined by the method described in (Giannopolitis et al., 1977) using riboflavin- nitro blue tetrazolium (NBT) in potassium phosphate buffer at pH 7.8 at 25°C. The absorbance was measured at 560 nm using a UV-Visible spectrophotometer (Shimadzu UV-1800, Japan). The obtained OD results were calculated and expressed as Units mg^− 1 of protein. CAT activity was assessed by monitoring the decrease in absorbance of hydrogen peroxide (H₂O₂) at 240 nm, following the method as outlined by Aebi (1974). Enzyme activity was defined as one unit when it decomposed 1 µM of H₂O₂ per minute at 25°C. The reduction of H₂O₂ was observed over 1 minute at 10-second intervals using a multimode reader (Tecan, Germany) at wavelength 240 nm. The obtained OD was calculated and results were quantified as Units mg^− 1 of protein. The total glutathione activity was measured in the resulting supernatant by the enzymatic method of Akerboom et al., (1981), and the total protein content determined according to (Bradford, 1976). This involved catalysing the amounts (nmols) of GSH, leading to a continuous reduction of DTNB to NBT. The resulting GSSG was then recycled by glutathione reductase and NADPH. It should be noted that the GSSG present may also yield a positive value in this reaction. The rate of the reaction is directly proportional to the concentration of glutathione. The yellow product, TNB, is measured using spectrophotometer at a wavelength of 412 nm.

2.3.6 Small antioxidant molecule

The mitochondrial fraction was determined by assessing the production of hydrogen peroxide using horseradish peroxidase (HRP) and phenol red by following the method as described by Pick and Keisari, (1981). The total protein content of tissues for enzyme assay was estimated as described by (Bradford, 1976). The absorbance was measured at 610 nm using a UV-Vis spectrophotometer (Shimadzu UV-1800, Japan). The obtained results were expressed as nmols/mg/protein.

2.3.7 Genotoxicity - Comet assay

The comet assay was conducted following the procedure outlined by (Singh et al., 1988). Post experiment after retrieval of mussels, the haemolymph/haemocyte were collected immediately after opening the valve from the posterior adductor mussels using a sterile hypodermic syringe. The haemolymph was gathered and transferred to a microcentrifuge tube, kept on ice and immediate analysed for: (i) Haemocyte count (stained with 4% Giemsa), and (ii) comet assay. Furthermore, cells were separated via centrifugation at 1000×g for 3 minutes (Almeida et al., 2013) for the comet assay. Haemocytes obtained from the harvested samples were combined with 0.65% low melting agarose (LMP), which was coated on pre-coated microscope slides. For alkaline unwinding, the slides were placed in an electrophoresis unit with electrophoresis buffer (consisting of 300 mM NaOH, 1mM Na2 EDTA, pH > 13) for 30 minutes, followed by electrophoresis at 25 V/300 mA for 30 minutes. Following electrophoresis, the slides were neutralized for 15 minutes with freshly prepared chilled neutralizing buffer containing 0.4 M Tris HCl at pH 7.4. Subsequently, dehydration was carried out using 70% ethanol, after which the slides were stored in a moisture-free chamber. For staining, the slides were treated with 80 µL of 20 µg/mL ethidium bromide for 10 minutes in dark and examined at 400x magnification using an epifluorescence microscope. The obtained images were processed using the IMAGEJ plugin software (NIH, USA). DNA damage was quantified as percent tail DNA (% tail DNA), which represents the percentage of DNA migrating from the head to the tail.

2.3.8 Neurotoxicity - Acetylcholinesterase

For assessment of neurotoxicity, a portion of the haemolymph was subjected to centrifugation at 4000×g for 10 minutes at 4ºC (Juhel et al., 2017). The resulting cell-free haemolymph (CFH) were collected, and the obtained aliquoted was stored at -20 ºC for subsequent analysis of acetylcholinesterase activity (AChE). During analysis, about 50 µL of CFH was mixed along with the reaction mixture and the sample absorbance was measured at 412 nm using a UV-Visible spectrophotometer (Shimadzu 1800, Japan). The obtained results were expressed as moles mg^− 1 min^− 1 x 10⁶.

2.3.9 Data analysis

All statistical analysis was performed using MS office 365 Excel (Microsoft-USA). The raw data obtained from each experiment was analyzed, processed and expressed finally as mean and standard deviation. Student t-test was performed to test the statistical differences between control and treatment for samples collected on day-to-day basis. One-way ANOVA followed by post-hoc analysis using Tukey HSD was performed. Differences were considered statistically significant when p value was < 0.05. In addition, the biomarker data sets were subjected to principal component analysis (PCA) to ascertain the relationship between treatments (unexposed and exposed to continuous chlorination) in different tissues (gills, mantle, digestive gland and foot) and their response to different biomarkers (SOD, CAT, ROS and H₂O₂) over the exposure period.

3.1 Mortality

As the concentration of actibromide increases, the survival rate decreases more rapidly, indicating the toxic effects of the biocide. This plot avoids any negative values on the Y-axis, ensuring clarity in survival analysis. The mussels in the control tank experienced a mortality ranging from 3–9% throughout the experiment. In comparison, the mortality rates of green mussels exposed to actibromide® at concentrations of 0.2, 0.5, and 1.0 mg L^-1 are gradually increasing towards 12, 7 and 4 days respectively (Fig. 2). Whereas in treatment tanks the onset of mortality was observed from the 3rd day of exposure and increasing to 38% by the 8th day, and 100% mortality occurring on the 12th day at residual concentration of 0.2 mg L^-1. Mortality significantly (one-way ANOVA, p < 0.05), increased with increasing biocide concentration and exposure time. Exposure to 0.5 mg L^-1 of actibromide® resulted in 12% mortality on the third day, escalating to 100% on the seventh day. In contrast, mussels subjected to 1.0 mg L^-1 of actibromide® showed 13% mortality on the second day, reaching 100% mortality by the fourth day.

3.2 Condition index (CI)

In the control tanks, the condition index remained relatively constant during the experiment. Figure 3 illustrates the CI (%) of adult green mussels in both control and treatment tanks which were exposed to actibromide®. In contrast to the control group, mussels exposed to 0.2 mg L^-1 of actibromide® showing a significant (p < 0.05) 48% decrease in their CI. Moreover, continuous actibromide® with a residual concentration of 0.5 mg L^-1 led to a significant reduction (57%, p < 0.05) in CI. Mussels treated with 1.0 mg L^-1 displayed a notably extremely significant (p < 0.001) reduction in CI, with a 92% decrease compared to tested sub lethal concentrations (0.2 & 0.5 mg L^-1). One-way ANOVA followed by Tukey HSD indicated significant differences (p < 0.05) compared to control.

3.3 Ammonia

Green mussels exposed to actibromide® induced ammonia excretion, instead of pseudofaeces production (Fig. 4). In control tanks there was pseudofaeces production and ammonia levels in these tanks ranged from 1.6–6.1 µmol L^-1 from 0 to 12 days of experiments. Whereas ammonia levels in experimental tanks showed an increase with time at respective concentration − 0.2 mg L^-1 (4.2–31.3 µmol L^-1), 0.5 mg L-1 (5.7–32.8 µmol L^-1) and 1.0 mg L^-1 (12.2–30.1 µmol L^-1). A significant (p < 0.001) difference was observed with biocidal concentrations versus control.

3.4 ROS generation

The total ROS production in gill, mantle, digestive gland, and foot tissue samples averaged between 212 and 7431 arbitrary units (A.U) in all tested biocide concentrations (0.2, 0.5, and 1.0 mg L^-1). Meanwhile, green mussels in the control tanks exhibited a ROS production of 36.54 ± 2.1 A.U. As shown in Fig. 5, it is evident that even the lowest actibromide® concentration (0.2 mg L^-1) led to a significant (p < 0.05) increase in intracellular ROS production in all the tissue samples tested. ROS peaked its production across the all-tested concentrations within the third day. Of the four tissues analyzed, the foot showed the lowest ROS levels as well as the mantle tissue following closely behind. In contrast, the digestive gland exhibited more pronounced ROS production, followed by the gill tissue showing the next highest levels, compared to those in control. There was a significant dose-dependent increase (p < 0.05) in ROS generation observed in all tissues at 0.5 and 1.0 mg L^-1 concentrations.

3.5 Hydrogen peroxide production (H₂O₂)

Among the tissues analyzed, the digestive gland showed significant (p < 0.05) highest H₂O₂ activity across all concentrations, followed by the gill tissue, while the foot tissue exhibited the lowest activity, with the mantle tissue slightly higher. As shown in Fig. 6, H₂O₂ levels were elevated in treated tanks compared to the control. H₂O₂ generation in all the tissues peaked on the third day of exposure and gradually declined as the experiment continued. In the mantle tissue, H₂O₂ levels were higher in tanks with 0.5 and 1.0 mg L^-1 actibromide® residuals compared to those with 0.2 mg L^-1. In the control tanks, H₂O₂ levels in the gill, mantle, digestive gland, and foot tissues varied between 13.2 and 23.8 µmole g^-1 fw. In contrast, the treated tanks showed elevated H₂O₂ levels in the digestive gland, with averages ranging from 25.2 to 36.8 µmole g^-1 fw.

3.6 Superoxide dismutase

Super-oxide dismutase (SOD), activity in gill, mantle, digestive gland, and foot tissues was detected in both control and treatment tanks (Fig. 7). SOD activity across all tissue samples ranged from 7.9 to 11.8 U mg^-1 protein in the untreated tanks. SOD levels in all tissue samples showed a significant, concentration-dependent increase (one-way ANOVA, p < 0.05) following the different residue of actibromide®. The digestive gland exhibited the highest activity, while gill, mantle, and foot tissues showed the lowest activity compared to control across all concentrations of actibromide®. At a concentration of 0.2 mg L^-1, SOD activity in the digestive gland peaked on the sixth day, ranging from 11.76 to 49.46 U mg^-1 protein. Where in, the peak SOD activity in the gill, mantle, and foot tissues occurred on the third day of exposure and subsequently declined throughout the experiment. At 0.5 mg L^-1, SOD activity significantly increased (p < 0.05) by the third day of exposure but slowly reduced by the sixth day. The lethal concentration of actibromide® (0.1 mg L^-1) significantly differed (p < 0.05) from the other concentrations (0.2 and 0.5 mg L^-1), with effects observed by the third day of exposure. Overall, SOD activity was higher in 0.5 and 1.0 mg L^-1 compared to 0.2 mg L^-1 across all tissues.

3.7 Catalase

The catalase (CAT) activity was detected in all the tissue samples of green mussels. In comparison CAT activity of mussels in the control tanks remained relatively consistent over the entire duration of the experiment (Fig. 8). Initially, CAT activity showed an increase up to the 3rd day of exposure. After the 6th day, CAT activity in all tissues remained more or less steady throughout the period, with an average of 28.19 ± 3.07 U mg^-1 protein. Mussels in the 0.2 mg L^-1 treatment tanks displayed similar trends from the 3rd day onwards. However, in the digestive gland, mantle, and foot tissues, CAT activity increased by approximately 2.0-fold in the 0.5 and 1.0 mg L^-1 treatments. Similar to SOD activity, CAT activity was higher in the digestive gland, ranging from 32.47 to 50.80 U mg^-1 protein, whereas gill tissue exhibited the lowest CAT activity across all concentrations. The CAT activity observed in digestive gland was almost the same in 0.5 and 1.0 mg L^-1 actibromide® residuals. However, CAT activity significantly (p < 0.05) increased at higher biocide concentration (1.0 mg L^-1) compared to 0.2 and 0.5 mg L^-1. No significant differences (p > 0.05) were observed in tissue samples treated at concentrations of 0.5 and 1.0 mg L^-1 of actibromide® residuals.

3.8 Total Glutathione (GSH)

Total glutathione was determined, in the gill, mantle, digestive gland and foot tissues (Fig. 9). Actibromide® exposed mussels showed a significant difference (p < 0.05) of GSH activity at all the tested concentrations (0.2, 0.5 and 1.0 mg L^-1) compared to controls. The digestive gland tissue showed the highest level of GSH, followed by gill, mantle and foot tissue. GSH levels increased significantly (p < 0.05) with increase in concentration of actibromide® (0.5 and 1.0 mg L^-1), compared to 0.2 mg L^-1. No change in glutathione levels was recorded in the mussel in the control tanks. Same as SOD and CAT activity GSH level also picked significantly (one-way ANOVA, p < 0.05) in the initial three days of exposure.

3.9 DNA damage

In tanks treated with actibromide®, the percentage tail DNA (% tail DNA) increased with the increase in concentration of actibromide® as well as increase in exposure time (Fig. 10). Particularly, in tanks treated with 0.2 mg L^-1, the % tail DNA exhibited a gradual rise towards the experiment's conclusion in comparison to the control. The mussels treated with 0.2 mg L^-1 initially showed 3.0 ± 2.0% tail DNA which significantly increased 13 ± 5.0% on the last day of the experiment. In contrast the mussels exposed to 0.5 mg L^-1 showed 34 ± 3.0% on the 7th day. Similarly, the mussels exposed to 1.0 mg L^-1 actibromide®, showed 3.0 ± 2.0% tail DNA in the first day of exposure, which significantly increased to 37% in the 3rd day of exposure. Student t-test performed between control and treatment samples showed significant increase (t-test, p < 0.001) in the length of DNA migration in green mussel haemocytes exposed to actibromide®. The length of migration appeared to plateau, while the extent of DNA damage in cells exposed to higher concentrations was too great to permit an accurate measurement of the migration pattern (Fig. 11). At each concentration of actibromide®, a homogeneous response, in the extent of DNA migration among cells was observed.

3.10 AChE activity

The gill tissue of green mussels exhibited lower AChE activity compared to haemolymph, which showed higher activity. Consequently, haemolymph was chosen for further analysis due to its elevated AChE activity. Following 3 days of exposure to actibromide®, all tested concentrations showed a considerable (3–5 folds) decrease in AChE enzyme activity (Fig. 12). Significant inhibition (P = 0.0005) was observed in each of the tested concentration compared to controls. Even at sub lethal concentrations (0.2 mg L^-1), a 78% inhibition was observed, while it was 84% and 93% with 0.5 and 1.0 mg L-1, respectively. Moreover, mussels treated with 0.5 and 1.0 mg L^-1 exhibited greater inhibition in AChE enzyme activity compared to those exposed to 0.2 mg L^-1.

3.11 One-way ANOVA, including Tukey’s HSD

A pair-wise comparison was performed between the different biomarker assay systems by performing One-way ANOVA followed by Post-hoc Tukey HSD, mainly to compare the effect of different tissues (gill, mantle, digestive gland and foot) exposed to varying actibromide® concentrations. The results from ANOVA clearly indicate statistically significant differences (p < 0.001) between the different biomarker assay systems, in response to different actibromide® concentrations. However, upon subjecting to post-hoc analysis using Tukey’s HSD test, the pair-wise data revealed that the biomarker assays were dependent on tissue-specific responses exposed to varying actibromide® concentrations. In case of gill, mantle and digestive gland tissues, exposed to 0.2 mg L^− 1 of actibromide®, pronounced ROS activity was found to exhibit extremely significant difference (p < 0.0001) compared to other assay systems. Results showed that even the lowest concentration, elicited ROS activity in mussels indicating the oxidative capacity of the biocide. Further it also indicated the triggering of the related enzymatic quenching systems in due course of time to nullify the effect of the biocide. However, in case of the foot tissues exposed to 0.2 mg L^− 1 of actibromide®, almost all of the enzyme systems were activated, mainly due to the fact that the mussels open their valves more frequently for feeding and also for sensing the surrounding environments, wherein the biocide gains access to the inner component of the mussels. The mussel tissues exposed to 0.5 mg L^− 1 of actibromide® also exhibited highly significant differences in biomarker assay systems. However, upon performing a pair-wise comparison subjected to post-hoc analysis, the results revealed that the gill and foot to be highly affected by the biocide, thereby eliciting a number of enzyme markers throughout the experimental duration. The sequence of events that triggered during the assays started with ROS generation followed by H₂O₂ generation leading to generation of other quenching enzyme systems such as SOD, CAT and GSH activities. Extremely significant differences (p < 0.0001) were noted with ROS production to the generation of other quenching enzyme systems. Mantle and digestive gland did not show much significant difference in between the enzyme assays. Pair-wise comparison of the assays performed between the different tissue samples exposed to 1.0 mg L^− 1 of actibromide® reveals high ROS activity in all tissues tested. Pronounced ROS activity with extremely significant differences compared to other assays were observed in the foot tissue.

3.12 Insights from multivariate analysis:

Correlation matrix analysis was carried out to assess the correlation efficiency among different biomarkers evaluated and the response of different experimental tissues. As represented in Fig. 13, there was a strong correlation among the indexes. Principal component analysis (PCA) was carried out with the measured values to understand the effects of actibromide® concentrations and experimental durations on the oxidative stress biomarkers and their quenching enzymes viz., ROS, H₂O₂, SOD, CAT, and GSH of different organ systems (Gill, mantle, digestive gland, and foot). The correlations between the parameters studied were based on the directions of the vector i.e., if the vectors are close and forming a small angle then they represent positive correlation, if they meet at 90^◦, they are not likely to correlate with each other, and when they diverge and meet at 180^◦, they are negatively correlated (Chakraborty et al., 2023; Chakraborty et al., 2024). The representation of principal component analysis is illustrated as a biplot in Fig. 13.

Experimental biomarkers like ROS, H₂O₂, SOD, CAT, and GSH exhibited a strong correlation. From the analysis of gill tissue biomarkers and experimental duration (Fig. 14a), it is observed that the components, PC1 and PC2 collectively sum up to 96.21% (PC1 85.43% and PC2 10.78%) of data variability. The experimental biomarkers like ROS, CAT, and H₂O₂ all fall in component 2 of the scatter plot. From the scatter plot it is clear that with an increase in these parameters in component 2, the toxicity increases. Actibromide® treatment of day 6; 0.5 mg L^− 1, day 3; 0.2 mg L^− 1, day 6; 0.2 mg L^− 1, day 12; 0.2 mg L^− 1, and day 9; 0.2 mg L^− 1 all fall in the segment of components 2 also indicating maximum toxicity. GSH and SOD enzyme activity is the only parameter that falls in component 1 along with the day 3; 1.0 mg L^− 1 and day 3; 0.5 mg L^− 1 concentrations. This suggests that these treatment groups can be directly correlated with GSH and SOD activity. The other experimental durations like day 0; control, day 3; control, day 6; control, day 12; control, day 0; 0.2 mg L-1, day 2; 0.2 mg L^− 1, day 0; 0.5 mg L^− 1, and day 0; 1 mg L^− 1 were clubbed and all fall together in the other domain devoid of any parameters. This suggests the lower concentration of day 0; 0.2 mg L^− 1, day 2; 0.2 mg L^− 1, day 0; 0.5 mg L^− 1, day 0; 1 mg L^− 1 and the above-mentioned control group duration expressed lower toxicity as they all fall in the scatter plot opposite to the five main biomarkers. It suggests that they comprise a separate cluster that is in negative correlation with the others thereby indicating the least toxicity compared to other experimental groups like day 6; 0.5 mg L^− 1, day 3; 0.2 mg L^− 1, day 6; 0.2 mg L^− 1, day 12; 0.2 mg L^− 1, day 9; 0.2 day 3; 1 mg L^− 1 and day 3; 0.5 mg L^− 1 conc. Accordingly, from the analysis of mantle tissue, biomarkers, and experimental duration (Fig. 14b), it is represented that the components, PC1 and PC2 collectively sum up to 97.8% (PC1 91.39% and PC2 6.41%) of data variability. In addition, the experimental biomarkers like ROS and GSH both fall in component 2 of the scatter plot. From the scatter plot it is clear that with the increase in these parameters in component 2, the toxicity increases. The actibromide® treatment of day 12; 0.2 mg L^− 1, day 9; 0.2 mg L^− 1, day 6; 0.5 mg L^− 1, and day 6; 0.2 mg L^− 1 all fall in the segment of component 2 indicating also maximum toxicity. Consequently, the H₂O₂, CAT, and SOD enzyme activity are the other parameters that fall in component 1 along with day 3; 0.2 mg L^− 1, day 3; 0.5 mg L^− 1, and day 3; 1.0 mg L^− 1 conc. This suggests, that these treatment groups can be directly correlated with H₂O₂, CAT, and SOD enzyme activity. The other experimental durations like day 0; 0.5 mg L^− 1 and day 0; 1.0 mg L^− 1 fall together in the other domain devoid of any parameters. This suggests those durations and concentrations expressed lower toxicity as they all fall in the scatter plot opposite to the five main biomarkers. On the other hand, experimental durations in control (day 0, 3, 6, 9, and 12); day 0 of 0.2 mg L^− 1 were all clubbed together and fell into other domains devoid of any experimental biomarkers. It suggests that they comprise a separate cluster that is in negative correlation with the others thereby indicating the normal response and least toxicity compared to other experimental groups like day 12; 0.2 mg L^− 1, day 9; 0.2 mg L^− 1, day 6; 0.5 mg L^− 1, day 6; 0.2 mg L^− 1, day 3; 0.2 mg L^− 1, day 3; 0.5 mg L^− 1, and day 3; 1.0 mg L^− 1 conc. In addition, from the analysis of digestive gland tissue biomarkers and experimental duration (Fig. 14c), it is observed that the components, PC1 and PC2 collectively sum up to 95.64% (PC1 88.09% and PC2 7.55%) of data variability. The experimental biomarkers like H₂O₂, GSH, and CAT all fall in component 2 of the scatter plot. actibromide® treatment of day 12; 0.2 mg L^− 1, day 9; 0.2 mg L^− 1, day 6; 0.5 mg L^− 1, and day 6; 0.2 mg L^− 1 all fall in the segment of component 2 indicating also maximum toxicity. From the scatter plot it is clear, that with the increase of activity in these parameters in component 2, the toxicity increases. Accordingly, the ROS, and SOD enzyme activity are the other parameters that fall in component 1 along with day 6; 0.2 mg L^− 1, day 6; 0.5 mg L^− 1, day 9; 0.2 mg L^− 1, and day 12; 0.2 mg L^− 1 conc. This suggests that these treatment groups can be directly correlated with ROS, and SOD enzyme activity. Similarly, the other experimental durations like day 0; 0.2 mg L^− 1, day 0; 0.5 mg L^− 1, day 0; 1.0 mg L^− 1 along with control day and day 6 all fall and clubbed together in the other domain devoid of any parameters. This suggests those durations and concentrations expressed lower toxicity as they all fall in the scatter plot opposite to the five main biomarkers. On the other hand, experimental durations like control (day 0, 3, and 12), all pointed together and fell into other domains devoid of any experimental biomarkers. It suggests that they comprise a separate cluster that is in negative correlation with the others thereby indicating the normal response and least toxicity compared to other experimental groups like those fall in PC1 and PC2 along with response biomarkers. Thereafter, from analysis of foot tissue biomarkers and experimental duration (Fig. 14d), it is observed that the components, PC1 and PC2 collectively sum up to 95.61% (PC1 88.14% and PC2 7.47%) of data variability. The experimental biomarkers like H₂O₂ and ROS all fall in component 2 of the scatter plot. The actibromide® treatment of Day 12; 0.2 mg L^− 1, Day 9; 0.2 mg L^− 1, Day 6; 0.2 mg L^− 1, and Day 3; 0.2 mg L^− 1 all fall in the segment of component 2 indicating also maximum toxicity. From the scatter plot it is clear that with the increase of activity in these parameters in component 2, the toxicity increases. Concurrently, the SOD, CAT, and GSH enzyme activity are the other parameters that fall in component 1 along with day 3; 0.5 mg L^− 1, day 3; 1mg L^− 1, and day 6; 0.5 mg L^− 1 conc. This suggests that these treatment groups can be directly correlated with SOD, CAT, and GSH enzyme activity. However, the other experimental durations like all control groups (day 0, 3, 6, 9, and 12) and day 0; 0.2 mg L^− 1, day 0; 0.5 mg L^− 1, day 0; 1.0 mg L^− 1 all fall and clubbed together in the other domain devoid of any parameters. This suggests day 0; 0.2 mg L^− 1, day 0; 0.5 mg L^− 1, and day 0; 1.0 mg L^− 1 durations and concentrations expressed lower toxicity and negative correlation as they all fall in the scatter plot opposite to the five main biomarkers. On the other hand, control groups comprise a separate cluster with the opposite direction of biomarkers thereby indicating the normal response compared to other experimental groups like those fall in PC1 and PC2 along with response biomarkers. Thus, PCA can provide information on the correlation between the individual and binary mixture toxicity using chemometric methodology.

Oxidizing biocides (e.g. chlorine) have been mostly used to combat biofouling in cooling water systems worldwide. Among them chlorine has been the most preferred biocide of choice due to its low cost, ease of handling and known breakdown products (Rajagopal et al., 1995; Nair et al., 1997; Venkatnarayanan et al., 2017). Kalpakkam a rocky shore area, located on the South East coast of India, is a high biofouling potential site with heavy fouling by green mussels observed in the cooling water systems (Rajagopal et al., 1997). The green mussel Perna viridis has been the extensively studied organism with respect to fouling control using oxidizing biocides (Rajagopal et al., 2003a; Chavan et al., 2018), mortality and valve movement response (Rajagopal et al., 1996), filtration activity, byssogenesis (Rajagopal et al., 2006), antioxidant enzyme activity (López-Galindo et al., 2010), genotoxicity (Chavan et al., 2016), and physical health conditions (Masilamoni et al., 2002). The power station abstracts coastal seawater at the rate of 33 m³ sec^− 1 with a velocity of ~ 3 m sec^− 1 through a sub seabed tunnel, which has been heavily infested by green mussels (Perna viridis) with loading of 211 kg m^− 2 observed in the vertical shaft of the seabed tunnel (Rajagopal et al., 1996). The station practices a low dose continuous chlorination regime of 0.2 ± 0.1 mg L^− 1, which is otherwise called as “exomotive chlorination”, essentially meant to drive out mussel larval spat without settling inside the CWS. However, under practical circumstances in large cooling water systems, proper reach of the biocide near the wall is questionable to the varying geometries and sizes of the conduits, which results in establishment of biofouling communities. Continuous low dose chlorination with residuals of 0.2 ± 0.1 mg L^− 1, has been found to be ineffective against settled adult green mussels (Masilamoni et al., 2002). Green mussels have been shown to detect chlorine concentrations as low as 0.15 mg L^− 1 and complete valve closure occurs above 0.55 mg L^− 1 (Masilamoni et al., 2002). It has been demonstrated using the Mossel Monitor® device, that valve closure in Perna viridis was initiated at residuals of 0.7 mg L^− 1 and 100% valve closure occurs at concentration of 1.0 mg L^− 1 (Venkatnarayanan et al., 2021). To combat mussel fouling in the process seawater heat exchangers (PSWHX) and their conduits and to comply with the discharge regulation and to improve the efficiency of the PSWHX supplementary biocidal addition using actibromide® has been envisaged. It is important to investigate the sub-lethal physiological responses of mussels which offer better indices than lethal responses in planning at a biocide dosing strategy. Quite often, high chlorine residuals, results in denaturation of cell membranes leading to lethal effects, particularly in the gills (Opresko, 1980), but at low residuals, other physiological activities of mussels are affected (Khalanski & Bordet, 1980). Exploring the impact of sub-lethal and lethal concentrations of continuous actibromide® on mortality and physiological changes in adult green mussels would help in fine tuning the biocidal regime to be adopted.

In the present study, the observed time taken for mortality for adult green mussels was found to be 12 days at 0.2 mg L^− 1, 7 days at 0.5 mg L^− 1 and 4 days at 1.0 mg L^− 1 actibromide® residuals. Investigation showed, the 100% mortality appears to be much lower in continuous actibromide® residuals than the previously reported for chlorine for P. viridis (Rajagopal, 1995), Mytilus edulis (James, 1967), Mytilopsis leucophaeata (Rajagopal et al., 2003a), and Dreissena polymorpha (Van Benschoten et al., 1995). It has been reported that at 1.0 mg L^− 1, mussels were able to open their valves to feed, although at a residue rate (White, 1966; Rajagopal et al., 1991b). Such high concentrations cannot be used practically in CWS from an environmental standpoint and biocidal discharges in marine outfall are regulated with an upper threshold limit of continuous discharge of 0.2 ± 0 actibromide®. 1 mg L^− 1 (Jensen, 1982; Venugopalan et al., 2011). In the present study, an elevation in stress biomarkers was noted within the initial three days of exposure to continuous.

Due to continuous actibromide® dosing, mussels were unable to compensate for the reduced food intake, leading to the absence of pseudofaeces production in the treatment tanks, in contrast to observations in the control tanks. Consequently, a notable decrease in growth of mussels is inevitable (Lewis, 1985). The decline in the Condition Index (CI) of P. viridis in the actibromide® treatment tanks indicated to a reduced growth, a phenomenon previously studied in Dreissena polymorpha and Mytilus edulis regarding physiological activities (Rajagopal et al., 2003a), and in P. viridis regarding the continuous effect of low-dose chlorination (Chavan et al., 2018). One possible explanation for this phenomenon could be inadequate feeding resulting from reduced valve opening, a response previously observed in P. viridis exposed to chlorinated environments (Masilamoni et al., 2002; Rajagopal et al., 2003b).

In this study, an increase in ammonia excretion was observed in mussels treated with actibromide® concentrations of 0.2, 0.5, and 1.0 mg L^− 1. Additionally, our personal observations indicated that ammonia production increases with the treatment of various concentrations of oxidizing biocide (Unpublished data). One of the reason could be due to an increase in the protein usage as a substrate during oxidative metabolism. Similar results were observed in Aulacomya ater ribbed mussels exposed to organophosphate pesticides (Führer et al., 2012) and in the bivalve Ruditapes decussates exposed to heavy metals (El-Shenawy, 2004).

The findings of this study demonstrate a significant elevation in reactive oxygen species (ROS) generation, even at the sub-lethal concentration of actibromide® (0.2 mg L^− 1) tested. Furthermore, higher concentrations of actibromide® (0.5 and 1.0 mg L^− 1) led to escalated levels of ROS in the digestive gland followed by gill. The previous study shows, rise in ROS production as well as increase in DNA damage in isolated tissues of Mytilus galloprovincialis (Dailianis et al., 2005) exposed to cadmium at micromolar concentration. Thermal stress, is one of the abiotic factors which directly affects organismal metabolism, leading to metabolic disorders and the accumulation of reactive oxygen species (ROS) (Bhat and Desai, 1998; Rajagopal et al., 2005). Increased ROS production in the gill tissue of P. viridis has been observed following exposure to both cold and heat stresses (Wang et al., 2018). Similarly, elevated ROS production has been observed in the gill tissue of zebrafish after cold treatment (Wu et al., 2015). Our results are in agreement with other studies that report the induction of oxidative stress after exposure to environmental pollutants and metals (Gómez-Mendikute et al., 2003; Collén et al., 2003; Lee and Shin, 2003; Koutsogiannaki et al., 2006).

ROS generates a variety of non-radicals such as superoxide radical (¹O₂•), hydroxyl radical (OH•) and hydrogen peroxide (H₂O₂•) etc. In the current study only generation of H₂O₂ was observed. Since high H₂O₂ concentration can alter vital functions in marine invertebrates (Abele-Oeschger et al., 1997), we have investigated the effect of actibromide® from sub lethal to lethal concentration. Elevated levels of H₂O₂ were observed in the digestive gland across varying concentrations, surpassing those detected in other assessed tissues. H₂O₂ can alter cell physiology through the formation of OH by Fenton reaction (Cavaletto et al., 2002). A comparable study conducted on Mytilus edulis veliger larvae, subjected to sodium hypochlorite treatment, revealed elevated levels of H₂O₂ upon exposure to concentrations ranging from 0.5 to 0.7 mg L^− 1. Similarly, in the common sole samples, increase in H₂O₂ level with increase in temperature was investigated (Aslan et al., 2018).

The SOD enzyme constitutes a primary defence against oxygen toxicity by catalysing the conservation of super oxide anion to oxygen and hydrogen peroxide, which can be sequentially removed by CAT (Manduzio et al., 2004). The findings of the current study suggest that adult green mussels P. viridis exhibit SOD activity across all tissues, with higher levels observed in the digestive gland followed by gill compared to the other two tissues examined. This observation seems logical, as the digestive gland is the main source of assimilation of feed and gills are regularly exposed to high levels of oxygen due to their respiratory function (Santovito et al., 2005). Similar activity of this enzyme was reported with P. viridis with changes in seasonal variation in the reproductive cycles as well as physico-chemical parameters (Verlecar et al., 2008). This observation is also in good agreement with previous report (Filho et al., 2001). Interestingly, M. edulis exposed to chlorine exhibited elevated activity of a specific isoform of SOD, without impacting its total activity. This suggests that the differential response of isoforms should also be taken into consideration in certain cases. (Manduzio et al., 2004) for assessing toxic responses.

Organisms when experiencing stress conditions, may increase the activity of antioxidant enzymes, such as catalase (CAT), to mitigate the overproduction of reactive oxygen species (ROS), specifically higher rates of H₂O₂ production, and thereby prevent additional cellular damage (Regoli et al., 2014). The increase of CAT activity in the present study signifies the generation of H₂O₂, which also showed a strong positive correlation in the presence of actibromide®. Previous study shows that, after treatment with sub-lethal concentrations of chlorine, M. galloprovincialis showed an increase in CAT activity, which agrees with the results observed with our study (López-Galindo et al., 2010). In contrast, M. galloprovincialis exposed to metal-contaminated areas revealed inhibition of enzyme activities (Vlahogianni et al., 2007). CAT is a significant biomarker for stress induced by oxidizing agents in comparison to SOD (Vlahogianni et al., 2007). Moreover, our findings are in agreement with Verlecar et al., (2008), Chavan et al., (2018) who assessed the seasonal variation and impact of chlorine on the CAT activity in P. viridis. These results endorse the capacity of mussel cells to detoxify superoxide (O₂), which is the precursor of most of the other ROS through the antioxidant enzymatic system.

GSH is a vital molecule that helps the organisms maintain their cellular health by protecting against oxidative stress and aiding in detoxification processes (Yan et al., 1997). Glutathione reductase (GR) which catalyses the reduction of oxidising glutathione (GSSG), is therefore essential for the maintenance of the GSH/GSSG ratio and the cellular redox status, protecting cells against oxidative damage (Box et al., 2007). Unlike CAT, GSH level was more in digestive gland than gill, mantle and foot in sub-lethal as well as lethal concentration of actibromide®. This result could be due to an increased consumption of glutathione required to counterbalance the stress increased due to biocide treatment. A previous study shows that, total glutathione increased slightly in Cyprinus carpio treated with sodium hypochlorite (NaClO), chlorine dioxide (ClO₂) and peracetic acid (PAA) (Elia et al., 2006). In a study of New Zealand P. canaliculus mussels infected with pathogenic Vibrio sp., total glutathione levels were measured. The study found no difference in total glutathione between infected and control mussels, indicating that glutathione levels may not be affected by this particular pathogen (Alfaro and Young, 2016).

Using the comet assay we have demonstrated a dose-response relationship between the level of DNA strand breaks and the environmentally relevant concertation of actibromide®. In our present study, the mussels which were exposed to continuous mode of actibromide® showed a high % tail DNA compared to control in all the tested concentrations. Biocide addition lead to breakage of DNA strands in the mussels Dreissena polymorpha which have been used as a bioindicator of toxicity in chlorinated drinking water (Bolognesi et al., 2004) environments. Previous study on oysters Crassostrea virginica showed DNA damage in haemolymph exposed to carcinogenic spiked sediments (Nacci et al., 1996). DNA strand breaks increased after one day of exposure to benzo[a]pyrene (B[a]P) (Siua et al., 2004). Previous studies have also suggested that tail DNA content and tail moment are the most satisfactory endpoints to express the recorded DNA damage (Devaux et al., 1998; Hartmann et al., 1997). Rajagopal et al., (2003b) studied, chlorine effects on foot activity, byssus thread production, filtration activity and shell valve movement in Dreissena polymorpha, Mytilopsis leucophaeata and Mytilus edulis. Similarly, (Chavan et al., 2016) have reported genotoxic effect on green mussels P. viridis of in use levels of continuous chlorination. In the present study we used alkaline comet assay method, the most widely used method to study DNA strand break (Singh et al., 1988; Collins, 2004; Frenzilli et al., 2009).

AChE has been initially considered as a specific biomarker for organophosphates; carbamate insecticides (Galgani and Bocquene, 1989; Escartín and Porte, 1997; Canty et al., 2007) and heavy metals (Day and Scott, 1990) in the marine mollusc Mytilus edulis. The enzyme plays an important role in the functioning of neurotransmitter acetylcholinesterase to choline in cholinergic synapses and neuromuscular junctions. In this present study, AChE activity was the most responsive biomarker, showing significantly inhibition levels in haemolymph at both sub-lethal and lethal concentrations (0.2, 0.5 and 1.0 mg L^− 1 TRO) of actibromide®. Studies by Vidal-Liñán et al., (2014) observed inhibition of AChE activity exposed to trace metals in gill tissue. Similar studies using the haemolymph of P. viridis showed strongly inhibition exposed to pharmaceutical drug carbamazepine (CBZ), the plasticizer bisphenol A (BAP) and the herbicide atrazine (ATZ) in a marine bivalve (Juhel et al., 2017). Previous studies reveal high AChE activity was observed in gills of Mytilus galloprovincialis and M. edulis and also in muscle tissue of fish from Mediterrian and Baltic Sea (Zinkl et al., 1987). Also, in M. galloprovincialis, changes in AChE activity were related to agricultural practices in areas where pesticides and biocides were frequently used (Escartín and Porte, 1997). Robillard et al., (2003) found lower AChE levels in the freshwater mussel Anodonta cygnea from a site in an agricultural area than in animals collected downstream from a municipal STP and at the reference site. Most studies assessing AChE levels have reported exposure to heavy metals, pesticides, pharmaceutical drugs etc (Handy et al., 2003). Till date effect of actibromide® on AChE enzyme activity of bivalves has not been investigated and the results indicates, it could serve as a sensitive biomarker of exposure to actibromide®.

The use of actibromide in heat exchangers of cooling water systems is aimed at controlling microbial growth and preventing fouling. Actibromide, a bromine-based biocide, effectively addresses issues like algae, bacteria, and slime that can impair heat exchanger performance and efficiency. By maintaining a clean and biofilm-free surface, actibromide helps ensure optimal heat transfer and prolongs the life of the equipment (Nancharaiah et a., 2000; Satyanarayanan et al., 2008). However, its use requires careful management to minimize environmental impacts, such as the potential release of bromine compounds into the water system. Proper dosing, monitoring, and adherence to environmental regulations are essential to balance effective biocide performance with minimal ecological disruption.

Sublethal cellular and physiological responses of mussels are an effective indicator for assessment of biocides and aid in planning a dosing strategy for their control. Efficacy of continuous actibromide® on adult green mussels was studied at different concentrations (0.2, 0.5, and 1.0 mg L^− 1) to determine mortality rates and cellular effects. Results showed 100% mortality within 12 days (0.2 mg L^− 1), 7 days (0.5 mg L^− 1) and 4 days (1.0 mg L^− 1) respectively exposed to actibromide®. Condition index of exposed mussels deteriorated (48–92%) with increase in concentrations and exposure time which was accompanied by increased ammonia production instead of pseudofaeces excretion. Actibromide® induced toxic effects by generating reactive oxygen species (ROS), inhibiting cellular processes in gills, mantle, digestive gland, and foot tissues. Maximum ROS generation was observed in the digestive gland, along with increased hydrogen peroxide (H₂O₂) production in a dose-dependent manner. Superoxide dismutase (SOD) and catalase (CAT) activity was higher in the digestive gland compared to other tissues. DNA damage, assessed by comet assay, showed significant damage even at the lowest dose (34%) with maximum damage (37%) at the highest concentration. Acetylcholinesterase (AChE) activity, a neurotoxic stress biomarker, was reduced at all tested concentrations (80–91%). As the present study indicates Actibromide® penetration at the cellular level, causing severe damage to gills and digestive gland, reducing feed consumption, and inducing neuronal and genotoxic effects. Previous biomonitoring observations revealed significant settlement and fouling by green mussels in process seawater heat exchangers, despite the consistent application of low-dose chlorination and booster dosing. Hence, for effective biofouling control in the heat exchangers, implementation of supplemental, targeted Actibromide® dosing can be considered

Acknowledgements:

The authors gratefully acknowledge the financial grant from Board of Research in Nuclear sciences, Department of Atomic Energy, Govt of India, funded research project entitled “Development of antifouling technologies against green mussel fouling for process cooling water system of MAPS”, to Dr. D. Inbakandan, Sanction NO 56/14/03/2020-BRNS/36152. The junior research fellowship was awarded to Ms. Bandita Badakumar in the research project. The authors would also like to thank the Head, Water & Steam Chemistry Division for providing necessary facilities to carry out the research work.

Credit author statement:

Bandita Badakumar: Investigation, Data Curation, Methodology, Formal analysis, software, Writing - Original Draft.

Dr. D. Inbakandan: Conceptualization, Methodology, Project administration.

Dr. S. Venkatnarayanan: Methodology, Investigation, manuscript - reviewing

Dr. T.V. Krishna Mohan: Funding and Project administration.

Dr. Y.V. Nanchariah: Review & editing.

Dr. P. Sriyutha Murthy: Conceptualization, Data Curation. Writing - Review & Editing, Supervision.

Conflict of Interest: All authors confirm that there are no known conflicts of interest associated with this publication, and there has been no significant financial support for this work that could have influenced its outcome.

Ethical Approval: This study did not involve human participants or animals, and therefore ethical approval was not required.

Funding: Board of Research in Nuclear sciences, Department of Atomic Energy, Govt of India, funded research project entitled “Development of antifouling technologies against green mussel fouling for process cooling water system of MAPS”, Sanction NO 56/14/03/2020-BRNS/36152.

Authorship: All authors have made significant contributions to the conception, design, execution, or interpretation of the study and are listed in the correct order of authorship. Each author has read and approved the final version of the manuscript.

Authors list:

Bandita Badakumar

Email: [email protected]

D Inbakandan*

Email: [email protected]

S. Venkatnarayanan

Email: [email protected]

T.V. Krishna Mohan

Email: [email protected]

Y.V. Nancharaiah

Email: [email protected]

P. Veeramani

Email: [email protected]

N.K. Pandey

Email: [email protected]

P. Sriyutha Murthy*

Email: [email protected]

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Mitigating Biofouling in Cooling Water System: Actibromide® to Combat Perna viridis Infestation and environmental impact

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Abstract

Figures

1. Introduction

2. Material and Methods

2.1 Collection of green mussels and maintenance of brood stock

2.2 Experimental setup

2.3 Effect of residual actibromide® on physiological metabolism of P. viridis

2.3.1 Physiological response

2.3.2 Ammonia

2.3.3 Sample preparation

2.3.4 Reactive oxygen species

2.3.5 Antioxidant enzymes

2.3.6 Small antioxidant molecule

2.3.7 Genotoxicity - Comet assay

2.3.8 Neurotoxicity - Acetylcholinesterase

2.3.9 Data analysis

3. Results

3.1 Mortality

3.2 Condition index (CI)

3.3 Ammonia

3.4 ROS generation

3.5 Hydrogen peroxide production (H₂O₂)

3.6 Superoxide dismutase

3.7 Catalase

3.8 Total Glutathione (GSH)

3.9 DNA damage

3.10 AChE activity

3.11 One-way ANOVA, including Tukey’s HSD

3.12 Insights from multivariate analysis:

4. Discussion

5. Conclusion

Declarations

References

Additional Declarations

Supplementary Files

Status:

Version 1

Mitigating Biofouling in Cooling Water System: Actibromide® to Combat Perna viridis Infestation and environmental impact

Status:

Version 1

Abstract

Figures

1. Introduction

2. Material and Methods

2.1 Collection of green mussels and maintenance of brood stock

2.2 Experimental setup

2.3 Effect of residual actibromide® on physiological metabolism of P. viridis

2.3.1 Physiological response

2.3.2 Ammonia

2.3.3 Sample preparation

2.3.4 Reactive oxygen species

2.3.5 Antioxidant enzymes

2.3.6 Small antioxidant molecule

2.3.7 Genotoxicity - Comet assay

2.3.8 Neurotoxicity - Acetylcholinesterase

2.3.9 Data analysis

3. Results

3.1 Mortality

3.2 Condition index (CI)

3.3 Ammonia

3.4 ROS generation

3.5 Hydrogen peroxide production (H2O2)

3.6 Superoxide dismutase

3.7 Catalase

3.8 Total Glutathione (GSH)

3.9 DNA damage

3.10 AChE activity

3.11 One-way ANOVA, including Tukey’s HSD

3.12 Insights from multivariate analysis:

4. Discussion

5. Conclusion

Declarations

References

Additional Declarations

Supplementary Files

Status:

Version 1

3.5 Hydrogen peroxide production (H₂O₂)