The peroxidase toxicity assay for the rapid evaluation of municipal effluent quality
0 
Abstract
Rapid and cost-effective tests for the evaluation of industrial and municipal effluents are urgently needed for environmental monitoring. In this context, peroxidase (PER) activity has been proposed as an early-warning biosensor for assessing the water quality of various wastewater discharges and leachates. The peroxidase-toxicity (Perotox) assay includes 0.1 µg/mL PER, albumin, DNA (for the DNA protection index), 0.001% monounsaturated Tween-80, and the substrates luminol and H2O2. The results revealed that an initial burst of luminescence was followed by a steady decrease in luminescence within the first minute, accompanied by periodic (cyclic) changes in the intermediate compound III (CIII) of PER. When urban effluents were added, PER activity was inhibited, with a concomitant increase in lipid peroxidation, indicating oxidative damage. The reduction in PER activity was also associated with the collapse in the periodic formation of CIII, alongside a steady increase in CIII over time. The addition of DNA to the reaction mixture helped mitigate the inhibition of PER by certain effluents, enabling the calculation of a DNA protection index. The levels of polystyrene (PS) in the organic fraction of the effluents were higher in the primary aeration lagoon (36 µg/L) compared to secondary lagoons and membrane filtration
Keywords
INTRODUCTION
In aquatic ecosystems, urban pollution encompasses wastewater from solid waste leachates, and domestic (e.g., hospitals) and industrial activities. Additionally, rainfalls collected by sewer systems carry contaminants from sources such as tires, asphalt, and erosion from infrastructure. These contaminants may enter the sewer drainage systems, where they sometimes mix with raw wastewater before reaching treatment plants[1]. When rainfall events become more frequent and intense due to global warming, wastewater may be directly discharged into temporary holding ponds or released into receiving waters to avoid overwhelming the capacity of wastewater treatment plants[2]. Besides the usual contaminants found in municipal wastewater, such as metals, polyaromatic hydrocarbons (PAHs), detergents, and surfactants, recent studies have identified the presence of emerging pollutants, including products derived from nanotechnology, pharmaceuticals, flame retardants, and plastics[3,4]. Plastics, in particular, pose a growing challenge, as they are ubiquitous in the form of microplastics (ranging from 5 mm to 1 µm) and nanoplastics (1-1,000 nm)[5]. Larger plastic materials not only accumulate in the environment but also gradually degrade into smaller fragments down to the nanoscale, making plastic nanomaterials more readily bioavailable at the sub-cellular level[6]. Nanoplastics have been detected in wastewaters at concentrations ranging from 0.01 to 10 µg/L. Studies have shown that nanoplastics can induce oxidative stress, leading to lipid peroxidation and DNA damage[7,8]. These effects align with the toxic properties of municipal wastewater, which can cause oxidative stress, genotoxicity, and neuroendocrine disruption[9,10]. Given these concerns, there is an urgent need for rapid and cost-effective tools to assess wastewater quality for environmental compliance, especially in the face of climate change and the emergence of new chemicals, such as plastic materials.
Peroxidases (PERs) are heme-containing enzymes involved in the elimination of the toxic reactive oxygen species H2O2, using electron donors (reducing agents) such as ascorbate, thiols, PAHs, and phenols[11-13]. The production of reactive oxygen species is often associated with the initiation of toxicity, and the inhibition of PER activity could serve as an early biochemical event in the manifestation of toxicity[14]. PER activity is involved in the peroxidative catalytic oxidation of H2O2 and requires an electron donor (A): AH2 + H2O2 → AOH + H2O. The formation of AOH indicates the production of oxidized by-products, which are typically less toxic than H2O2. PER enzymes participate in the PER-oxidase reaction and induce oscillations in NADH (electron donor) and O2[15]. These enzymes can oxidize a wide variety of environmental pollutants[16]. Additionally, various environmental pollutants have been shown to inhibit PER activity, leading to oxidative damage, such as lipid peroxidation and ferroptosis[17]. A previous report found that industrial effluents inhibited PER activity, leading to higher H2O2 levels[18]. Interestingly, the inhibition of PER by industrial effluents was linked to fish toxicity (mortality), making this enzyme a potential screening tool for large sample volumes and contributing to a reduction in fish testing for environment monitoring surveys. An interesting variation of the PER inhibition test is the restoration of PER activity following the addition of DNA. In the so-called DNA protection assay, it was revealed that 70% of industrial effluents showing DNA protection were genotoxic, as determined by a bacterial DNA repair (SOS chromotest) assay. A recent study of municipal effluents found that 90% of the tested effluent extracts inhibited PER activity, suggesting potential toxicity[19]. Of these, 60% showed DNA protection of PER activity, further suggesting the presence of genotoxic compounds in these wastewaters. This finding corroborates previous studies indicating that municipal effluents are genotoxic to fish and mussels[20,21]. These studies highlight the usefulness of this rapid assay for screening wastewater quality. In the present study, the PER assay was revisited to better understand PER inhibition by including an unsaturated lipid analog to monitor lipid peroxidation and examine the oscillations of PER intermediates. This approach will provide new information on the mechanisms of complex mixtures and the onset of oxidative damage, in addition to the detection of potential genotoxic compounds. The catalytic intermediates (compound 0, I, II, and III) oscillate over time, with compound III representing a non-catalytic intermediate that binds HOO- to the active center, leading to O2 and H2O when H2O2 is in excess relative to the electron donor molecules[15]. Monitoring the cyclic changes in compound III, a non-catalytic intermediate, was performed to better understand the mechanisms at play during PER inhibition by municipal wastewaters.
The objective of this study was to evaluate the peroxidase-toxicity (Perotox) assay as a rapid tool for assessing wastewater quality in municipal effluents. The Perotox assay involved the addition of unsaturated lipids to investigate the effects of PER inhibition and the sustained levels of H2O2, as indicated by lipid peroxidation. Additionally, the protection of DNA from PER activity was examined by measuring changes in DNA strand breaks. The oscillatory changes in PER intermediate III were also assessed in an attempt to understand the mechanisms behind PER inhibition by municipal wastewater.
MATERIALS AND METHODS
Sample preparation
Horseradish PER, salmon sperm DNA, Tween-80, luminol, and bovine serum albumin were purchased from Sigma Chemicals (Ontario, Canada). Each substance was prepared at a concentration of 0.1 mg/mL in phosphate-buffered saline (PBS: 140 mM NaCl, 1 mM KH2PO4, and 1 mM NaHCO3, pH 7.5) and stored in the dark at 4 °C for no more than one week. The DNA was dissolved in 50 mM NaCl containing 1 mM
Wastewater extract characteristics
Municipal wastewater samples were collected from four cities with varying population sizes as 24-h composite samples over 3 days [Table 1]. A volume of 1 L from each sample was transported to the laboratory, stored at 4 °C in the dark, and subsequently filtered using a 0.8 µm pore cellulose filter. The filtered samples were then stored in dark bottles at -20 °C until analysis. The municipal effluents were fractionated using a C18 solid-phase extraction column. A 200 mL volume was passed through a C18 cartridge containing 200 mg of sorbent, washed with 10 mL of Milli-Q water, and eluted with 1 mL absolute ethanol. The dissolved organic matter (DOC) content in the ethanol extract was quantified at 254 nm as described previously[22], using the equation: DOC (mg/L) = 0.21A + 1.25. The levels of plastic materials in the organic matter matrix (i.e., the C18 extract) were assessed using the copper fluorescence quenching method[23]. After adding 50 µM CuSO4, fluorescence intensities for polypropylene (PP) (excitation: 250 nm, emission: 324 nm) and polyvinyl chloride/polystyrene (PS) (excitation: 295 nm, emission: 411 nm) were measured. Additionally, humic/fulvic acids (HA/FA) were detected at excitation/emission wavelengths of 265 and 463 nm, respectively. The fluorescence difference before and after CuSO4 addition was calculated, and standard solutions of PS (20 nm diameter) and PP (100 nm diameter) were used for calibration. The results were expressed as µg PP or PS/PVC-equivalents/ mg DOC. The levels of PAHs in the ethanol extracts were determined using fixed-wavelength fluorometry[24]. Briefly, 100 µL of the ethanol extracts were placed in a dark 96-well microplate, and fluorescence was measured using a fluorescence microplate reader (Neo-2 Synergy, Biotek Instruments, USA) at the following wavelengths: excitation at 290 nm/emission at 340 nm (for light PAHs: naphthalene), excitation at 325 nm/emission at 370 nm (for medium PAHs: pyrene), and excitation at 385 nm/emission at 440 nm [for heavy PAHs: benzo(a)pyrene]. Standard solutions of PAHs were used for calibration, and the recovery rates for each PAH size class ranged between 80% and 95% during C18 extraction columns and ethanol elution.
Physio-chemical characteristics of wastewaters
| Wastewater treatment plant | Population | DOC | PS (µg/L) | PAHslight (µg/L) | PAHsmedium (µg/L) | PAHshigh (µg/L) |
| Lagoons | 123,182 | 1.18 ± 0.01a | 38 ± 0.5a | 51 ± 2a | 8 ± 0.5a | 0.06 ± 0.01a |
| SecA1 | 656,205 | 1.23 ± 0.02b | 16 ± 0.2b | 33 ± 13b | 7.2 ± 0.7a | 0.47 ± 0.06b |
| SecA2 | 589,748 | 1.14 ± 0.02a | 11 ± 0.3b | 22 ± 2b | 3.8 ± 0.3b | 0.19 ± 0.07c |
| SeCM | 523,000 | 1.095 ± 0.012c | 10 ± 0.2b | 30 ± 3b | 3.8 ± 0.8b | 0.15 ± 0.07c |
Perotox assay
The Perotox assay was based on a previous methodology for toxicity screening of industrial effluents[18,19]. The assay involves a reaction mixture containing 0.1 µg/mL PER, albumin, and 0.0001% Tween-80 in PBS (an unsaturated surfactant serving as a proxy for unsaturated lipids), combined with 0.1% H2O2 -0.1 mM luminol mix. DNA (0.1 µg/mL) was also added to duplicate wells to assess recovery of PER activity (DNA protection assay). The reaction media was added to the effluent extract equivalent to 10× of the C18 extract and mixed for 5 min in a clear-bottom black 96-well microplate (Synergy-4, Biotek Instruments, CA, USA). Afterward, luminol and H2O2 were added to initiate the reaction, which was maintained at 25 °C for 30 min. Luminescence readings and measurements at 418 nm (indicative of the PER compound III intermediate) were taken every 2 min[25]. The same procedure was repeated using ethanol extracts pre-incubated with 1 µg/mL DNA for 5 min to evaluate the influence of DNA on PER reaction rates. The DNA protection index was calculated as: PER activity with DNA/Per activity. At the end of the incubation period, 5 µL of ice-cold TCA was added to the reaction mixture and centrifuged at 10,000 × g for 5 min at 4 °C. The supernatant (containing DNA strands) was mixed with 10 µg/mL Hoechst dye in 100 mM KH2PO4 buffer (pH 8), and fluorescence was measured at 360 nm excitation/460 nm in dark microplates as described above. Standard solutions of salmon sperm DNA were used for calibration. For Tween-80 peroxidation, malonaldehyde levels were determined using thiobarbituric acid reagent methodology[26]. Briefly, 10 µL of the reaction mixture at 30 min was mixed with 90 µL of water and 50 µL of TCA 1% was added. Then, 50 µL of thiobarbituric acid (0.67%) was added, and the mixture was heated at 70 °C for 5 min. Fluorescence at
Data analysis
The in vitro exposure experiments to the various influents/effluents from four cities representing different wastewater treatments were repeated three times in the absence and presence of added DNA to evaluate DNA protection. The population sizes of the cities (anonymous) and the types of wastewater treatments employed were as follows [Table 1]: aerated lagoon (Lag), two secondary activated sludge effluents (SecA), and a secondary membrane bioreactor (SecM). This study aimed to assess the impact of treatment processes in cities with different population sizes. To account for the influence of population size, an analysis of covariance (ANCOVA) was performed, with the treatment process as the main variable and population size (log-transformed) as a covariate. Critical differences between wastewater treatments from the four cities and the controls (Milli-Q water used as an operational control at the effluent containers and during C18 extraction steps) were identified using the least significant difference (LSD) test, with significance set at α ≤ 0.05. The relationships between the measured endpoints were examined using Pearson’s moment correlation test, with results summarized in Table 2. All statistical analyses were conducted using the StatSoft software package (USA).
Correlation analysis
| Population | PS | OC | PAHsl | PAHsmed | PAHsHigh | Perox | PeroxDNA | DNAP | TBARS | TBARSDNA | C3rate | C3DNA rate | DNAS | |
| PS | -0.56 | 1 | ||||||||||||
| OC | -0.14 | 0.19 | 1 | |||||||||||
| PAHsl | -0.06 | -0.23 | 0.81 | 1 | ||||||||||
| PAHsmed | -0.09 | -0.18 | 0.79 | 0.96 | 1 | |||||||||
| PAHsHigh | -0.17 | 0.26 | 0.83 | 0.68 | 0.80 | 1 | ||||||||
| Perox | -0.34 | 0.18 | 0.42 | 0.5 | 0.59 | 0.59 | 1 | |||||||
| PeroxDNA | -0.19 | 0.15 | 0.16 | 0.33 | 0.41 | 0.46 | 0.38 | 1 | ||||||
| DNAP | 0.08 | -0.03 | -0.04 | 0.10 | 0.10 | 0.09 | -0.29 | 0.75 | 1 | |||||
| TBARS | 0.77 | -0.46 | 0.02 | -0.02 | -0.03 | -0.05 | -0.52 | -0.26 | 0.16 | 1 | ||||
| TBARSDNA | 0.62 | -0.46 | -0.31 | -0.26 | -0.32 | -0.43 | -0.67 | -0.59 | -0.07 | 0.65 | 1 | |||
| C3rate | -0.08 | 0.45 | 0.44 | 0.26 | 0.23 | 0.32 | 0.30 | -0.23 | -0.45 | -0.15 | -0.26 | 1 | ||
| C3DNA rate | 0.76 | -0.47 | 0.03 | -0.02 | -0.04 | -0.07 | -0.32 | -0.55 | -0.34 | 0.57 | 0.58 | 0.22 | 1 | |
| DNAs | 0.31 | -0.46 | 0.46 | 0.68 | 0.50 | 0.1 | -0.09 | -0.15 | 0.07 | 0.31 | 0.40 | 0.10 | 0.25 | 1 |
RESULTS
Optimization of Perotox assay
The Perotox assay includes the addition of an unsaturated detergent (Tween-80) as a proxy for unsaturated lipids to evaluate lipid peroxidation [Figure 1]. This assay assesses the effects of PER inhibition and sustained H2O2 levels on protein, PER intermediate analysis (418 nm), and DNA protection. The consequences of sustained H2O2 levels due to PER inhibition were measured by lipid peroxidation and DNA strand break formation. Moreover, the assay enables real-time monitoring of PER states by tracking absorbance changes at 403 and 418 nm, corresponding to the native and component III (CIII) intermediate states of PER, respectively. CIII is a non-catalytic intermediate involved in the (spontaneous) transformation of 2H2O2 → O2 and 2H2O when the reducing substrate is deleted relative to H2O2. This new Perotox “version 2.0” biosensor offers a more detailed assessment of PER activity under oxidative stress at the protein/enzyme, DNA, and lipid levels.
Figure 1. Principle of the Perotox assay. The Perotox enzyme assay is based on the production of oxygen radicals during the PER reaction of H2O2 and a reducing substrate. Both H2O2 and the oxidized products could, in turn, oxidize various proxy substrates in addition to luminol, such as an unsaturated lipid (Tween-80), protein (albumin), and DNA. The oxidation of unsaturated carbons of Tween-80 detergent leads to peroxidation (aldehyde formation), while interaction with DNA leads to DNA strand breaks. The addition of DNA during the PER reaction with H2O2 and lumino/fluorogenic substrate (luminol or dihydrofluorescein) could also protect PER against inhibition by the xenobiotics, the so-called DNA protection assay. Perotox: Peroxidase-toxicity; PER: peroxidase.
Functional characteristics of the Perotox assay
Following the addition of H2O2 and luminol, a rapid increase in luminescence is observed, followed by a decrease over time [Figure 2A]. During the reaction, increasing the amount of unsaturated detergent Tween-80 results in the formation of malonaldehyde (hydrolysis of peroxides) [Figure 2B]. This rise in malonaldehyde levels was also accompanied by an increased formation rate of PER compound III [Figure 2C], suggesting that not all H2O2 is oxidized by PER and luminol. Instead, PER compound III followed a secondary pathway, leading to the degradation of H2O2 to H2O and O2 when the luminol concentration is lower than that of H2O2. In the presence of DNA, luminescence decreases over time, similar to the pattern observed with PER alone [Figure 2D]. The levels of DNA strand breaks, as determined by the TCA-precipitation method, also increased as the DNA concentration rose [Figure 2E]. As with PER alone, the formation of compound III over time increased with the addition of DNA, further suggesting that H2O2 was somewhat in excess relative to luminol when DNA or Tween-20 was present [Figure 2F].
Figure 2. PER reaction characteristics in the presence of Tween-80 and DNA. Perotox assay in control (absence of toxicants) conditions. Change in luminescence (A), production of malonaldehyde (B), and formation of Complex III (C) in the absence of exogenous DNA. Change in luminescence (D), acid-soluble DNA (E), and formation of Complex III (F) in the presence of exogenous DNA. PER: Peroxidase; Perotox: peroxidase-toxicity.
Intermediates of PER activity
The native form of the PER enzyme and its intermediate compound III (CIII) were evaluated during the PER reaction in the presence of DNA and municipal effluent extracts [Figure 3]. During this reaction, the PER enzyme transitioned through various intermediates, from its native form (characterized by an absorbance at 403 nm) to various intermediate compounds such as C0, CI, CII and CIII (characterized by an absorbance at 418 nm for CIII intermediate). The native form acts as a link between the peroxidation (C0, CI, CII) and oxidation (CIII) reactions: CIII
Figure 3. Oscillation of compound III over time. The change in compound III (418 nm) intermediate was monitored over time with PER alone, PER with DNA in the controls, and in the presence of municipal effluent samples. PER: Peroxidase.
Application of the Perotox assay on municipal wastewaters
The Perotox assay was tested on municipal wastewaters subjected to different treatments [Table 1]: lagoons (Lag), Secondary aerated lagoons (SecA1 and A2), and secondary membrane filtration (SecM). All these effluents underwent primary treatment involving particle sedimentation. The wastewater effluents were characterized by measuring dissolved organic carbon (DOC), PS materials, and light, medium, and heavy PAHs [Table 1]. The wastewater originated from Canadian cities with different population sizes, all of which utilized aeration lagoons followed by secondary aeration or membrane filtration. A strong correlation was observed between population size and effluent flow rates (r = 0.94; P < 0.001). DOC levels were higher in the Lag and SecA1 treatments compared to SecA2 and SecM treatments. However, DOC levels showed no significant relationship with population size. The concentrations of PS materials in the organic matrix were also evaluated. PS levels were significantly higher in the Lag treatment compared to the other lagoons employing a secondary treatment. Moreover, PS levels were negatively correlated with population size (r =
The impact of municipal effluents (at 10× concentration) from different wastewater treatment types (Lag, SecA1, SecA2, and SecM) was examined using the Perotox assay [Figure 4]. PER activity was significantly reduced by all wastewater treatment types, with SecA2 causing the most severe inhibition [Figure 4A]. PER activity was negatively correlated with the population size (r = -0.9), suggesting that water quality impacts were stronger with larger populations. All treatment types significantly increased the peroxidation of the unsaturated detergent, with SecA2 treatment resulting in the greatest damage [Figure 4B]. The levels of detergent peroxidation were significantly correlated with population size (r = 0.57). The formation rate of CIII was significantly reduced only with SecM treatment, with no effect observed from population size.
Figure 4. Influence of municipal wastewater extracts on the PER assay. The Perotox assay was used to determine the potential toxic properties of municipal influents and effluents (A). The formation rate of CIII (B; 418 nm) and the levels of aldehydes (C; TBARS) were determined. The control represents the PER assay in the presence of the vehicle (ethanol). The data are expressed as the mean with the standard error. The star symbol * indicates significance relative to controls. PER: Peroxidase; Perotox: peroxidase-toxicity; TBARS: thiobarbituric acid reactants (malonaldehyde).
Influence of DNA addition to the Perotox assay
A small decrease in PER activity was also observed in the presence of exogenous DNA, though the decrease was less pronounced compared to PER alone [Figure 5A]. The levels of thiobarbituric acid reactants were also elevated across the various treatment processes, with SecA2 showing the highest levels of detergent (lipid) peroxidation [Figure 5B]. The formation rate of the CIII intermediate (which occurs when the co-substrate luminol decreases relative to H2O2) was elevated in the SecA2 treatment, indicating that excess
Figure 5. Influence of municipal wastewaters on the peroxidase toxicity in the presence of DNA. The Perotox assay was tested on municipal influents and effluents in the presence of DNA (A) with detergent peroxidation (B). The formation rate of CIII (C; 418 nm), the DNA protection index (D), and the formation of acid-soluble DNA (E) were determined. The control represents the PER assay in the presence of the vehicle (ethanol). The data are expressed as the mean with the standard error. The star symbol * indicates significance relative to the controls. Perotox: Peroxidase-toxicity; PER: peroxidase.
DISCUSSION
Mode of action of PER inhibition
The Perotox assay is a simple biosensor designed to evaluate water quality in complex mixtures, such as effluents. The principle behind the assay is based on the observation that PER inhibition, which reduces the rate of H2O2 elimination, leads to increased oxidative damage, such as lipid peroxidation (Tween-80) and DNA breaks. In a previous study[18], the Perotox assay successfully identified toxic effluents in rainbow trout when PER activity was reduced, highlighting its potential as an alternative method to reduce the need for fish in toxicity testing. Furthermore, inhibition of PER reaction by primary- and secondary-treated municipal wastewaters (Singapore) has also been observed, further supporting its applicability for screening municipal wastewaters[30]. In the absence of wastewater extracts, PER intermediates oscillate between compounds 0, I, II, and III, eventually returning to compound 0, as shown in Figure 2. CIII formation occurs when the reducing substrate concentration is lower than that of H2O2. leading to non-catalytic formation of O2 and H2O, similar to catalase activity[15]. However, unlike catalase, this step is non-catalytic and does not substantially enhance the production rates of O2 and H2O. In the presence of DNA, which can interact with luminol (3-aminophthalate) via H bonds at the amino group, the formation rate of CIII was inversely related to DNA-Per activity (r = -0.55), supporting the view that CIII formation was involved with PER activity inhibition. When DNA was present, CIII levels increased, and the cyclic changes in CIII were maintained, though with lower amplitudes. The FD of the PER reaction decreased, as expected, with the addition of DNA (FD = 1.215) compared to PER reaction alone (FD = 1.344), suggesting that the increased complexity of the reaction environment leads to limited substrate availability for PER. The addition of effluent samples and DNA further decreased the FD down to FD = 1.174. PER activity was somewhat lowered in the presence of DNA, indicating that luminol and H2O2 reacted less efficiently. This finding aligns with previous observations that DNA strands reduce the chemoluminescence reaction rate of luminol with H2O2[31].
The influence of municipal wastewaters on PER
Following the addition of municipal effluents with or without DNA, CIII oscillations are maintained at lower amplitudes, with a steady increase in CIII formation over time. This suggests that PER activity can be inhibited by various chemicals in effluents, potentially due to a reduction in the spatial domain of the enzyme (FD, especially in the presence of complex macromolecules such as DNA) and the formation of non-catalytic CIII over time. Ferroptosis, a form of programmed cell death, occurs when Fe, H2O2, and decreased PER activity lead to the accumulation of lipid hydroperoxides[32]. This mechanism could represent a fundamental basis for cell death induced by Fe release (from hemoproteins such as PERs) and the formation of lipid hydroperoxides due to persistent H2O2 in cells. Heme proteins have been shown to bind various metals, releasing Fe during heme breakdown (catabolism) via the heme oxygenase pathway[33]. Several chemicals, including herbicides (e.g., paraquat), detergents (e.g., Brij-96 and Tween-20), phenol, and heavy metals such as Hg, Co, and Ni, have been reported to inhibit PER activity[13]. Conversely, some pro-oxidant compounds such as Cd, Mg, and Zn have been observed to increase PER activity, which could deplete cellular antioxidants over the long term. In algae, PER activity was commensurate with copper and lead levels in stormwater, suggesting that certain chemicals can induce PER activity[11]. While reduced PER activity in control samples was associated with lower biomass, increases in PER activity above the normal range were also linked to reduced biomass (algal growth). This suggests that deviations in PER activity from the normal range may be harmful due to the accumulation of either H2O2 or oxidized substrates.
Wastewater contaminants and PER activity
This study found an inverse relationship between PER activity and population size (and corresponding flow rates), with plastic materials being more common in smaller cities. Smaller wastewater treatment plants are perhaps ill-equipped to reduce plastic pollutant loads in effluents[34]. The presence of plastics was linked to increased CIII formation rates in the absence of DNA (r = 0.45), indicating that plastics could also reduce PER activity by enhancing CIII formation rates. Exposure of Vicia faba L seedlings to microplastics led to reduced PER activity and root sugar content[35]. Moreover, the addition of Cd with microplastics also resulted in decreased PER activity. In mouse primary hepatocyte cells, 20 nm nanoplastics demonstrated a pronounced ability to induce lipid peroxidation, due to the reduced activity of antioxidant enzymes such as superoxide dismutase, catalase, and perhaps glutathione PER[36]. However, in this study, PER activity showed a negative correlation with thiobarbituric acid levels, both with and without added DNA, suggesting that decreased enzyme activity (and consequently reduced removal of H2O2) leads to the oxidation of unsaturated Tween-80 in the reaction mixture. In wastewater samples, there was no direct indication that plastics inhibited PER activity, as no correlation was observed. However, recent findings suggest that PER enzyme can adsorb onto the surfaces of amino-coated (i.e., positively charged) PS nanoparticles[37]. This adsorption forms a protein corona on the plastic nanoparticles, reducing their Zeta potential, surface roughness, and PER activity. These results imply that plastic nanoparticles could induce oxidative stress and cellular damage by inhibiting PER activity. Further research is needed to elucidate the role of plastic size and shape in modulating PER activity. Interestingly, the negative correlation between PS plastics and lipid hydroperoxide formation (in the absence or presence of DNA) suggests that PS plastics could in some way adsorb other chemicals, thereby preventing inhibition of PER activity. Plastics are well known to act as chemical vectors, binding various substances on their hydrophobic surfaces[38]. For example, plastics can adsorb chemicals with logKow values between 3.3 and 9, such as perfluoroalkyl substances (PAFSs), hexachlorocyclohexanes (HCHs), and polycyclic aromatic hydrocarbons (PAHs). During weathering, the formation of C-O and N-H groups on plastic surfaces enhances their capacity to adsorb metals such as Cr, Pb, and Co. It is, therefore, possible that plastic materials in effluents could act as chemical sinks and as vectors for pollutant transport. Notably, municipal effluents often contain unsaturated surfactants and detergents, which can also contribute to peroxidation[39]. This hypothesis could be tested by directly applying the Perotox assay to wastewater samples, without the addition of Tween-80, to measure lipid peroxidation levels.
CONCLUSIONS
The Perotox assay is proposed as a rapid screening tool for complex liquid mixtures, such as effluents and leachates. An unsaturated detergent was included in the reaction mix to evaluate the impact of PER inhibition (indicated by lower H2O2 removal rates), which serves as a proxy for the formation of lipid hydroperoxides from polyunsaturated lipids. The assay revealed that inhibition of PER activity is associated with decreased amplitudes in CIII oscillations, an increased rate of non-catalytic intermediate CIII formation, reduced spatial dimensions, and the formation of lipid peroxidation products. These findings form the toxicological basis for understanding the effects of PER inhibition in organisms exposed to various environmental pollutants. The addition of exogenous DNA during the reaction did not result in significant changes to PER activity, suggesting it could be used to identify chemical interactions that prevent PER inhibition. This simple biosensor represents a convenient and cost-effective tool for screening various complex mixtures and emerging contaminants that compromise water quality and toxicity.
DECLARATIONS
Authors’ contributions
Extraction of municipal wasterwaters for the Perotox assay, data analysis, and revision of the draft manuscript: André, C.
Collection and preparation of municipal wastewatetrs, supervision, funding acquisition, and revision of the draft manuscript: Smyth, S.A.
Funding acquisition, data acquisition, data analysis and preparation of the draft and final manuscript:
Availability of data and materials
The data that support the findings of this study are available from the corresponding author upon reasonable request.
Financial support and sponsorship
This work was funded by the Chemical Management Plan for wastewater monitoring of Environment and Climate Change Canada.
Conflicts of interest
Gagné, F. is an Editorial Board Member of the journal Water Emerging Contaminants & Nanoplastics.
Ethical approval and consent to participate
Not applicable.
Consent for publication
Not applicable.
Copyright
© The Author(s) 2025.
REFERENCES
1. Shafi, M.; Lodh, A.; Khajuria, M.; et al. Are we underestimating stormwater? Stormwater as a significant source of microplastics in surface waters. J. Hazard. Mater. 2024, 465, 133445.
2. Ranieri, E.; D’Onghia, G.; Lopopolo, L.; et al. Influence of climate change on wastewater treatment plants performances and energy costs in Apulia, south Italy. Chemosphere 2024, 350, 141087.
3. Holeton, C.; Chambers, P. A.; Grace, L.; Kidd, K. Wastewater release and its impacts on Canadian waters. Can. J. Fish. Aquat. Sci. 2011, 68, 1836-59.
4. Sivarajah, B.; Lapen, D. R.; Gewurtz, S. B.; Smyth, S. A.; Provencher, J. F.; Vermaire, J. C. How many microplastic particles are present in Canadian biosolids? J. Environ. Qual. 2023, 52, 1037-48.
5. Uogintė, I.; Pleskytė, S.; Pauraitė, J.; Lujanienė, G. Seasonal variation and complex analysis of microplastic distribution in different WWTP treatment stages in Lithuania. Environ. Monit. Assess. 2022, 194, 829.
6. Okoffo, E. D.; Thomas, K. V. Mass quantification of nanoplastics at wastewater treatment plants by pyrolysis-gas chromatography-mass spectrometry. Water. Res. 2024, 254, 121397.
7. Acar, Ü.; İnanan, B. E.; Zemheri-Navruz, F. Ecotoxicological effects of polystyrene nanoplastics on common carp: insights into blood parameters, DNA damage, and gene expression. J. Appl. Toxicol. 2024, 44, 1416-25.
8. Li, R.; Nie, J.; Qiu, D.; Li, S.; Sun, Y.; Wang, C. Toxic effect of chronic exposure to polyethylene nano/microplastics on oxidative stress, neurotoxicity and gut microbiota of adult zebrafish (Danio rerio). Chemosphere 2023, 339, 139774.
9. Maranho, L. A.; André, C.; DelValls, T. A.; Gagné, F.; Martín-Díaz, M. L. In situ evaluation of wastewater discharges and the bioavailability of contaminants to marine biota. Sci. Total. Environ. 2015, 538, 876-87.
10. Gagné, F.; Roubeau-Dumont, E.; André, C.; Auclair, J. Micro and nanoplastic contamination and its effects on freshwater mussels caged in an urban area. J. Xenobiot. 2023, 13, 761-74.
11. Wong, M. Y.; Sauser, K. R.; Chung, K. T.; Wong, T. Y.; Liu, J. K. Response of the ascorbate-peroxidase of Selenastrum capricornutum to copper and lead in stormwaters. Environ. Monit. Assess. 2001, 67, 361-78.
12. Busa, L. S. A.; Komatsu, T.; Mohammadi, S.; et al. 3,3’,5,5’-tetramethylbenzidine oxidation on paper devices for horseradish peroxidase-based assays. Anal. Sci. 2016, 32, 815-8.
13. Ilyina, A. D.; Martínez, H. J. L.; López, L. B. H.; Mauricio, B. J. E.; García, J. R.; Martínez, J. R. Water quality monitoring using an enhanced chemiluminescent assay based on peroxidase-catalyzed peroxidation of luminol. ABAB. 2000, 88, 045-58.
14. O'Brien, P. J. Radical formation during the peroxidase catalyzed metabolism of carcinogens and xenobiotics: the reactivity of these radicals with GSH, DNA, and unsaturated lipid. Free. Radic. Biol. Med. 1988, 4, 169-83.
15. Møller, A. C.; Hauser, M. J.; Olsen, L. F. Oscillations in peroxidase-catalyzed reactions and their potential function in vivo. Biophys. Chem. 1998, 72, 63-72.
16. Sellami, K.; Couvert, A.; Nasrallah, N.; Maachi, R.; Abouseoud, M.; Amrane, A. Peroxidase enzymes as green catalysts for bioremediation and biotechnological applications: a review. Sci. Total. Environ. 2022, 806, 150500.
17. Mesmar, F.; Muhsen, M.; Mirchandani, R.; Tourigny, J. P.; Tennessen, J. M.; Bondesson, M. The herbicide acetochlor causes lipid peroxidation by inhibition of glutathione peroxidase activity. Toxicol. Sci. 2024, 202, 302-13.
18. Gagné, F.; Blaise, C. Evaluation of industrial wastewater quality with a chemiluminescent peroxidase activity assay. Environ. Toxicol. Water. Qual. 1997, 12, 315-20.
19. Gagné, F.; Smyth, S. A.; André, C. Plastic contamination and water quality assessment of urban wastewaters. SSRN 2024.
20. Gilroy, È. A. M.; Kleinert, C.; Lacaze, É.; et al. In vitro assessment of the genotoxicity and immunotoxicity of treated and untreated municipal effluents and receiving waters in freshwater organisms. Environ. Sci. Pollut. Res. Int. 2023, 30, 64094-110.
21. Lacaze, E.; Devaux, A.; Bony, S.; et al. Genotoxic impact of a municipal effluent dispersion plume in the freshwater mussel Elliptio complanata: an in situ study. J. Xenobiotics. 2013, 3, 6.
22. Brandstetter, A.; Sletten, R. S.; Mentler, A.; Wenzel, W. W. Estimating dissolved organic carbon in natural waters by UV absorbance (254 nm). J. Plant. Nutrition. Soil. 1996, 159, 605-7.
23. Lee, Y. K.; Hong, S.; Hur, J. Copper-binding properties of microplastic-derived dissolved organic matter revealed by fluorescence spectroscopy and two-dimensional correlation spectroscopy. Water. Res. 2021, 190, 116775.
24. Aas, E.; Beyer, J.; Goksøyr, A. PAH in fish bile detected by fixed wavelength fluorescence. Mar. Environ. Res. 1998, 46, 225-8.
25. Longu, S.; Medda, R.; Padiglia, A.; Pedersen, J. Z.; Floris, G. The reaction mechanism of plant peroxidases. Ital. J. Biochem. 2004, 53, 41-5.
26. Wills, E. D. Evaluation of lipid peroxidation in lipids and biological membranes. In: Snell K, Mullock B, editors. Biochemical toxicology: a practical approach. IRL Press; Washington, DC. 1987; pp 127-52. https://www.abebooks.com/9780947946678/Biochemical-Toxicology-Practical-Approach-APractical-0947946675/plp (accessed 2025-01-21)
27. Jurgelėnė, Ž.; Stankevičiūtė, M.; Kazlauskienė, N.; Baršienė, J.; Jokšas, K.; Markuckas, A. Toxicological potential of cadmium impact on rainbow trout (oncorhynchus mykiss) in early development. Bull. Environ. Contam. Toxicol. 2019, 103, 544-50.
29. Schepers, H. E.; van, B. J. H.; Bassingthwaighte, J. B. Four methods to estimate the fractal dimension from self-affine signals. IEEE. Eng. Med. Biol. Mag. 2002, 11, 57-64.
30. Vdovenko, M. M.; Papper, V.; Marks, R. S.; Sakharov, I. Y. Chemiluminescent assay of phenol in wastewater using HRP-catalysed luminol oxidation with and without enhancers. Anal. Methods. 2014, 6, 8654-9.
31. Huang, Y.; Yue, N.; Fan, A. Cationic liposome-triggered luminol chemiluminescence reaction and its applications. Analyst 2020, 145, 4551-9.
32. Xia, J.; Si, H.; Yao, W.; et al. Research progress on the mechanism of ferroptosis and its clinical application. Exp. Cell. Res. 2021, 409, 112932.
33. Kaminsky, L. The role of trace metals in cytochrome P4501 regulation. Drug. Metab. Rev. 2006, 38, 227-34.
34. Maw, M. M.; Boontanon, N.; Aung, H. K. Z. Z.; et al. Microplastics in wastewater and sludge from centralized and decentralized wastewater treatment plants: effects of treatment systems and microplastic characteristics. Chemosphere 2024, 361, 142536.
35. Wang, H.; Li, Y.; Liu, L.; et al. A study on the growth and physiological toxicity effects of the combined exposure of microplastics and cadmium on the Vicia faba L. Seedlings. Bull. Environ. Contam. Toxicol. 2024, 112, 83.
36. Wang, Y.; Zhao, X.; Tang, H.; et al. The size-dependent effects of nanoplastics in mouse primary hepatocytes from cells to molecules. Environ. Pollut. 2024, 355, 124239.
37. Zhou, J.; Yu, Y.; Luan, Y.; Dai, W. The formation of protein corona by nanoplastics and horseradish peroxidase. Nanomaterials 2022, 12, 4467.
38. Tumwesigye, E.; Felicitas, N. C.; C, A. F.; Siwe, N. X.; William, N. G.; Odume, O. N. Microplastics as vectors of chemical contaminants and biological agents in freshwater ecosystems: current knowledge status and future perspectives. Environ. Pollut. 2023, 330, 121829.
Cite This Article
Open Access
How to Cite
André, C.; Smyth, S. A.; Gagné F. The peroxidase toxicity assay for the rapid evaluation of municipal effluent quality. Water Emerg. Contam. Nanoplastics 2025, 4, 2. http://dx.doi.org/10.20517/wecn.2024.63
Download Citation
Export Citation File:
Type of Import
Tips on Downloading Citation
Citation Manager File Format
Type of Import
Direct Import: When the Direct Import option is selected (the default state), a dialogue box will give you the option to Save or Open the downloaded citation data. Choosing Open will either launch your citation manager or give you a choice of applications with which to use the metadata. The Save option saves the file locally for later use.
Indirect Import: When the Indirect Import option is selected, the metadata is displayed and may be copied and pasted as needed.
About This Article
Copyright
Data & Comments
Data
0
Comments
Comments must be written in English. Spam, offensive content, impersonation, and private information will not be permitted. If any comment is reported and identified as inappropriate content by OAE staff, the comment will be removed without notice. If you have any queries or need any help, please contact us at support@oaepublish.com.