Introduction
Over the past few decades, herbicides have been widely used to enhance global agricultural productivity and ensure food security to meet the demands of a growing population (Tilman et al. Reference Tilman, Cassman, Matson, Naylor and Polasky2002). However, the extensive use of herbicides has also led to considerable environmental challenges. China, as the largest user of herbicides worldwide, applies quantities that far exceed the global average (Sharma et al. Reference Sharma, Kumar, Shahzad, Tanveer, Sidhu, Handa, Kohli, Yadav, Bali, Parihar, Dar, Singh, Jasrotia, Bakshi and Ramakrishnan2019). In the agricultural ecosystem, herbicides do not degrade completely over time, leaving substantial residues in the soil (Froger et al. Reference Froger, Jolivet, Budzinski, Pierdet, Caria, Saby, Arrouays and Bispo2023; Zhang et al. Reference Zhang, Yan, Jones, Jiao, Sun, Liu, Zhu, Zhang, Zhai, Shen and Chen2022). These residues are frequently mobilized by rainfall, leading to their dispersion through environmental pathways, including atmospheric deposition, surface runoff, and leaching into groundwater, ultimately accumulating in freshwater ecosystems (He et al. Reference He, Li, Gao, He and Hao2023). The persistence and widespread presence of herbicides in aquatic ecosystems pose significant risks to the health and functioning of these systems. Non-target organisms, including microorganisms, aquatic plants, animals, and even humans, are vulnerable to herbicide exposure, which may disrupt ecological processes and biodiversity (Parrón et al. Reference Parrón, Requena, Hernández and Alarcón2011). Specifically, herbicides can severely impair the germination and growth of aquatic plant seeds, threatening the vital ecological functions they provide (Zhang et al. Reference Zhang, Hu and Li2024).
Submerged aquatic plants play a fundamental role in maintaining the ecological balance of lake ecosystems, particularly in shallow lakes. As primary producers, submerged plants enhance dissolved oxygen levels by releasing oxygen through photosynthesis, which is essential for sustaining aquatic life (Pedersen et al. Reference Pedersen, Colmer and Sand-Jensen2013). Additionally, submerged plants possess excellent nutrient uptake ability, enabling them to accumulate heavy metals and degrade organic pollutants from lake water and sediment, thereby underscoring their high ecological value and widespread use in ecological restoration projects (Ersoy et al. Reference Ersoy, Scharfenberger, Baho, Bucak, Feldmann, Hejzlar, Levi, Mahdy, Noges, Papastergiadou, Stefanidis, Sorf, Sondergaard, Trigal, Jeppesen and Beklioglu2020; Jiang et al. Reference Jiang, Li, Chang, Li and Yin2012; Sand-Jensen and Borum Reference Sand-Jensen and Borum1991). Moreover, submerged plant seeds are indispensable for maintaining plant populations and ensuring their resilience to environmental stressors, further amplifying their importance in lake ecosystems (O’Donnell et al. Reference O’Donnell, Fryirs and Leishman2016; Zhao et al. Reference Zhao, Zhang, Liu, Yin, Wang and Li2017).
Butachlor, an acetanilide herbicide, is extensively used in rice (Oryza sativa L.) cultivation in China due to its high efficiency and cost-effectiveness (Lin et al. Reference Lin, Pang, Zhou, Wu, Bhatt and Chen2021). Despite its agricultural benefits, butachlor’s strong hydrophobicity and low leaching potential result in its adsorption to soil particles, but it can still accumulate in water bodies through surface runoff or sediment resuspension. This characteristic, combined with its toxicity to non-target organisms, raises environmental concerns (Jolodar et al. Reference Jolodar, Karimi, Bouteh, Balist and Prosser2021). Recent studies have shown that herbicides can induce hormetic effects in plants, stimulating growth at low concentrations and inhibiting it at higher doses (Belz and Duke Reference Belz and Duke2014). These responses may affect plant physiology and species interactions, especially in variable aquatic environments. Chloroacetamide herbicides like butachlor are often applied with safeners such as fenclorim to reduce crop toxicity by enhancing detoxification via glutathione (GSH) S-transferases (Scarponi et al. Reference Scarponi, Quagliarini and Del Buono2006). However, the ecological effects of such safener–herbicide combinations on non-target aquatic plants remain poorly understood. Previous studies have highlighted the highly toxic mechanisms of butachlor toward various organisms (Abigail et al. Reference Abigail, Samuel and Ramalingam2015), yet its specific impacts on the germination and early growth of aquatic plants remain insufficiently understood. Given the ecological importance of aquatic plants in agro-aquatic systems, understanding how butachlor influences their germination is critical for assessing its broader environmental impact and informing sustainable herbicide management strategies.
The use of butachlor poses significant risks to freshwater vegetation. Even at minimal concentrations (as low as 0.0001 mg L−1), butachlor has been shown to severely inhibit the growth of submerged plant species such as coontail (Ceratophyllum demersum L.), western waterweed [Elodea nuttallii (Planch.) H. St. John], and Asian tape grass [Vallisneria natans (Lour.) H. Hara] (Pan et al. Reference Pan, Li, Xu and Gao2009). At higher concentrations (>1.06 mg L−1), butachlor becomes highly toxic, negatively affecting the dry matter accumulation and growth of floating aquatic plants such as water hyacinth [Eichhornia crassipes (Mart.) Solms] and water lettuce (Pistia stratiotes L.) (Huang et al. Reference Huang, Zhao, Xian, Li, Zhou and Sun2024). Additionally, butachlor exposure reduces chlorophyll content in V. natans and E. nuttallii, severely impairing their photosynthetic efficiency (Abigail et al. Reference Abigail, Samuel and Ramalingam2015). This herbicide also inhibits enzymatic activities crucial for plant stress response, such as peroxidase activity in common duckweed (Lemna minor L.), which compromises the plant’s ability to detoxify reactive oxygen species (ROS) under stress (Gomes et al. Reference Gomes, Kitamura, Marques, Barbato and Zamocky2022). These findings illustrate the pervasive and detrimental impacts of butachlor on submerged plant physiology, raising concerns about its broader ecological implications in lake ecosystems.
Herbicides are known to affect non-target plant germination and growth, but their impacts vary depending on concentration, exposure duration, and species sensitivity. In aquatic systems, compounds like glyphosate have been shown to suppress seed germination and elevate oxidative stress markers such as malondialdehyde (MDA) and proline (Pro), while reducing antioxidant enzyme activities (Huang et al. Reference Huang, Deng, Li, Ye, Wei, Sang and Xie2020). In contrast, some terrestrial studies report stimulatory effects under certain conditions, reflecting species- and compound-specific responses (Shen et al. Reference Shen, Zhang, Peng, Zhang and He2022; Yin et al. Reference Yin, Yan, Zhuang, Fang and Gu2023).
However, most research has focused on terrestrial plants or non-target aquatic organisms, with limited attention to submerged macrophytes such as V. natans (Zhang et al. Reference Zhang, Hu and Li2024). As a keystone species in shallow freshwater ecosystems, V. natans helps stabilize sediments, cycle nutrients, and purify water due to its strong adaptability and ability to absorb suspended particles and nutrients. It is widely distributed and plays an important role in reducing eutrophication. Despite its ecological value, little is known about how herbicides affect V. natans during seed germination. Butachlor, a common amide herbicide used in rice farming, is persistent in the environment (30 to 60 d half-life) and toxic to aquatic organisms (Abigail et al. Reference Abigail, Samuel and Ramalingam2015), but its impact on the antioxidant system of V. natans seeds remain poorly understood. Given that V. natans seeds have a narrow obovoid shape and glandular hairs, they may exhibit distinct responses to butachlor-induced stress, highlighting the need for further investigation. This study aims to fill this gap by addressing the following research questions: (1) Does butachlor reduce the germination rate of V. natans seeds? (2) How do varying concentrations of butachlor influence the activities of oxidative enzymes, antioxidants, and oxidative products in these seeds? (3) Is there a correlation between butachlor’s impact on the antioxidant system and changes in germination rates? This work seeks to clarify the ecological impacts of butachlor in aquatic ecosystems, offering insights to support sustainable agricultural practices and strengthen environmental protection, ultimately promoting both ecological and agricultural sustainability.
Materials and Methods
The experiments were conducted using dormancy-broken seeds of V. natans. The seeds were procured in April 2023 from Xin Daye Yemucao Seed Industry, located in Hubei Province, China.
Germination Experiment
The experiment was conducted under controlled laboratory conditions in May 2023. Before the experiment, the V. natans seeds were surface sterilized using a 5% sodium hypochlorite solution for 5 min, followed by thorough rinsing with deionized water (4 to 5 times) to remove residual disinfectant. For the germination assay, 50 V. natans seeds were immersed in 10 ml of prepared butachlor solutions at concentrations of 2, 20, 200, and 2,000 μg L−1, with a control treatment (0 μg L−1) using deionized water. These concentrations represent a gradient of environmentally and toxicologically relevant exposure levels: 20 μg L−1 reflects typical butachlor concentrations detected in paddy field drainage water (Alhassan et al. Reference Alhassan, Ashong, Ababio and Kwaansa-Ansah2025); 200 μg L−1 simulates peak runoff concentrations during intense rainfall events (Imaizumi et al. Reference Imaizumi, Suzuki, Shiraishi, Nakajima, Serizawa, Sakurai and Shiraishi2018); and 2,000 μg L−1 represents an extreme pollution scenario used in threshold-effect assessments, consistent with concentrations applied in worst-case ecotoxicological studies (Onwona-Kwakye et al. Reference Onwona-Kwakye, Hogarh and Van den Brink2020). The germination experiment was carried out according to the method applied by Zhao et al. (Reference Zhao, Zhang, Liu, Yin, Wang and Li2017). The seeds and the solutions were added to a 6-well transparent microplate (Corning Inc., Corning, NY, USA), which was placed in a constant-temperature incubator set to 25 ± 1 C with a 12-h:12-h light/dark cycle. Each treatment was replicated three times. Seed germination was recorded daily at 10:00 AM, and the experiment was terminated when no additional seed germination was observed for 3 consecutive days. In this study, germination ability was evaluated by germination percentage and germination index. Germination percentage was calculated using the following equation:
$${\rm{Germination\;percentage}} = {{n}\over{N}} \times 100\% $$
where n is the total number of germinated seeds, and N is the total number of test seeds. Germination index was calculated using the following equation:
$${\rm{Germination\;index}} = \sum {{{Gt}}\over{{Dt}}}$$
where Gt is the number of germinated seeds at t days and Dt is the number of days required for germination.
Assessment of the Physiological Effects of Butachlor on Vallisneria natans Seeds
To investigate the physiological effects of butachlor on V. natans seeds, approximately 1.5 g of seeds (≈1,500 seeds) were immersed in 10 ml of butachlor solutions at concentrations of 0 (control), 20, 200, and 2,000 μg L−1. The solutions were prepared using deionized water and distributed in a 6-well transparent microplate (Corning). Each treatment was replicated three times. Additionally, a seed-free control (without seeds) was included to assess butachlor degradation in the absence of plant material. All microplates were placed in a constant-temperature incubator set to 25 ± 1 C with a 12-h:12-h light/dark cycle for 3 d. At 12, 24, 48, and 72 h, microplates corresponding to each treatment were removed from the incubator. A subsample of seeds (approximately 0.2 g) was blotted dry with sterile filter paper and immediately weighed, with three replicates per concentration. The seeds were then transferred to 2-ml cryogenic tubes and stored at −80 C for subsequent biochemical analysis. Additionally, 5 ml of the culture solution was collected from each well, including those from the seed-free control group, for immediate quantification of butachlor concentration. From the preserved samples, one representative set was selected for determining butachlor accumulation within the seeds.
At 12, 24, 48, and 72 h, all samples were collected for the measurement of antioxidant indicators, including ascorbate peroxidase (APX), catalase (CAT), superoxide dismutase (SOD), GSH, Pro, MDA, and hydrogen peroxide (H2O2). Each indicator was analyzed separately. A summary of the single-factor experimental design is presented in Figure 1.
Figure 1. Experimental design.
Determination of Butachlor in Solution and Seed
The concentration of butachlor in the culture solution and V. natans seeds was determined using gas chromatography, following the method of Tsuda et al. (Reference Tsuda, Kojima, Harada, Nakajima and Aoki1997). For solution analysis, 5 ml of the sample was mixed with 5 ml of n-hexane and vortexed for 5 min, and the upper phase was collected. The solvent was evaporated to dryness using a solvent evaporation workstation, reconstituted with 1 ml of n-hexane, vortexed for 30 s, and filtered before being injected into a 2-ml chromatography vial for gas chromatographic analysis. For seed analysis, a 0.2-g seed sample was ground in a mortar with 5 ml of water and transferred into a centrifuge tube, and then extracted with 5 ml of n-hexane by vortexing for 5 min. The upper phase was processed as described for solution samples before gas chromatographic analysis.
Determination of H2O2 and MDA Contents
After exposure to butachlor for 12, 24, 48, and 72 h, 0.2 g of seeds from each treatment were harvested, thoroughly rinsed with deionized water, and blotted dry. To determine H2O2 content, the seeds were homogenized in 2 ml of 1% trichloroacetic acid (TCA) on an ice bath, and the resulting extract was centrifuged at 12,000 × g for 15 min at 4 C. The supernatant was collected and mixed with 0.5 ml of phosphate-buffered saline and 1 ml of potassium iodide. The absorbance at 390 nm was measured using a spectrophotometer, and the H2O2 concentration was calculated by comparing the absorbance values to a standard calibration curve (Velikova et al. Reference Velikova, Yordanov and Edreva2000).
To determine MDA content, a total of 0.2 g seeds were homogenized in a prechilled mortar with 2 ml of 10% TCA on an ice bath to form a uniform homogenate. The homogenate was transferred to a centrifuge tube and centrifuged at 12,000 × g for 10 min at 4 C, and the resulting supernatant was collected as the enzyme extract. For the assay, 1.5 ml of enzyme extract was mixed with 1.5 ml of 0.6% thiobarbituric acid (TBA), thoroughly vortexed, and incubated in a boiling water bath for 15 min. The total MDA content was determined using the TBA method (Heath and Packer Reference Heath and Packer1968).
Determination of Antioxidant Enzymes
To determine the activities of antioxidant enzymes, a total of 0.2 g of seeds was homogenized in a prechilled mortar with 2 ml of ice-cold 50 mmol L−1 sodium phosphate buffer (pH 7.8). The homogenate was then transferred to a centrifuge tube and centrifuged at 12,000 × g for 20 min at 4 C. The supernatant was collected as the enzyme extract. The 0.2 ml of the enzyme extract was used to determine CAT activity by monitoring the changes in H2O2 concentration at 240 nm according to the method outlined by Zhang and Kirkham (Reference Nakano and Asada1996). For the SOD activity, 0.05 ml of enzyme extract was used to determine the enzyme’s capacity to inhibit the photochemical reduction of nitroblue tetrazolium at 560 nm, as described by Zhang and Kirkham (Reference Nakano and Asada1996). To assess APX activity, 0.1 ml of enzyme extract was used, and the change in ascorbic acid (ASA) concentration was measured at 290 nm, following the method of Nakano and Asada (Reference Nakano and Asada1981).
Determination of Pro and GSH Contents
To determine the Pro and GSH contents, a total of 0.2 g of seeds was homogenized in a prechilled mortar with 2 ml of 5% sulfosalicylic acid on an ice bath to obtain a uniform homogenate. The homogenate was transferred to a centrifuge tube and centrifuged at 12,000 × g for 20 min at 4 C. The resulting supernatant was collected as the enzyme extract. For GSH quantification, 0.05 ml of enzyme extract was analyzed based on the reaction of ferric ion, ASA, and bathophenanthroline at 412 nm, following the method of Griffith (Reference Griffith1980). Similarly, Pro content was determined using 0.05 ml of enzyme extract, based on its reaction with triketohydrindene hydrate at 520 nm, following the method of Bates et al. (Reference Bates, Waldren and Teare1973).
Data Analyses
Statistical analyses were conducted in IBM SPSS Statistics (Armonk, NY, USA). A one-way ANOVA was used to assess the differences in butachlor accumulation in V. natans seeds, residual butachlor concentration in solution, seed germination parameters, and physiological indicators among different butachlor concentrations (0, 20, 200, and 2,000 μg L−1) at each time point (12, 24, 48, and 72 h). A repeated-measures ANOVA was conducted to evaluate the temporal variations in butachlor accumulation, residual butachlor concentration, germination parameters, and physiological indicators under different butachlor treatments (0, 20, 200, and 2,000 μg L−1) throughout the experimental period. Additionally, Pearson correlation analysis was performed to examine the relationships among butachlor accumulation in seeds, residual butachlor concentration in solution, seed germination parameters, and physiological indicators. The confidence limits were set at 95%.
Results and Discussion
Changes in Germination Ability of Vallisneria natans Seeds
Germination percentages dropped to 23%, 23%, 17%, and 14% at 0, 20, 200, and 2,000 μg butachlor L−1, respectively (Figure 2A). The germination index also decreased to 1.6, 1.7, 1.2, and 0.9 at the same herbicide concentrations (Figure 2B). The greatest reductions in these parameters compared with the control were observed at 200 and 2,000 μg L−1. The seed germination percentage was lower than that of the control (0 μg L−1) at butachlor concentrations of 200 and 2,000 μg L−1. Similarly, the seed germination index at butachlor concentrations of 2,000 μg L−1 was lower than that of the control.
Figure 2. The germination ability of seeds of Vallisneria natans under different concentrations of butachlor after 14 d: (A) germination percentage and (B) germination index. Significant differences from the control are indicated by * (P < 0.05) and ** (P < 0.01). n = 3. Error bars represent the standard deviation.
Changes in Butachlor Content in Vallisneria natans Seeds and in the Solution
Butachlor concentrations in V. natans seeds increased to 109, 330, and 267 µg kg−1 after 72-h exposure to 20, 200, and 2,000 μg L−1, respectively (Figure 3A). Residual butachlor in the solution decreased to 2.3, 5.5, and 39 μg L−1 at the same exposure levels (Figure 3B). The highest accumulation in seed tissue and corresponding decline in aqueous butachlor levels were observed at 200 and 2,000 μg L−1. Seed butachlor content was markedly higher than in the control (0 μg L−1) at 200 and 2,000 μg L−1, while the residual concentrations in solution were lower than initial levels.
Figure 3. The butachlor content in Vallisneria natans seeds and in the solution under different butachlor concentrations: (A) butachlor content in seeds (μg kg−1) and (B) butachlor concentration in solution (μg L−1). Significant differences from the control are shown by * (P < 0.05) level compared with control; ** (P < 0.01). n = 3. Error bars represent the standard deviation.
Changes in H2O2 and MDA Contents in Vallisneria natans Seeds
After 12-h exposure, H2O2 content in seeds was 0.6, 0.5, 0.6, and 0.7 µmol g−1 FW at 0, 20, 200, and 2,000 μg butachlor L⁻¹, respectively (Figure 4A). MDA content after 24 h exposure was 6.6, 7.6, 6.7, and 12.3 nmol g−1 FW at the same concentrations (Figure 4B). The greatest increases in both parameters compared with the control were observed at 2,000 μg L−1. H2O2 content at 2,000 μg L−1 was 0.1 µmol g−1 FW higher than the control. Similarly, MDA content under 2,000 μg L−1 was 5.7 nmol g−1 FW higher than that of the control.
Figure 4. The hydrogen peroxide (H2O2) and malondialdehyde (MDA) contents in Vallisneria natans seeds under different concentrations of butachlor and exposure time: (A) H2O2 content (μmol g−1 FW) and (B) MDA content (nmol g−1 FW). Significant differences from the control are shown by * (P < 0.05) level compared with control; ** (P < 0.01). n = 3. Error bars represent the standard deviation.
Changes in Activities of Antioxidant Enzymes in Vallisneria natans Seeds
The activity of CAT in V. natans seeds after 12 h was 31, 31, 33 and 9.7 U min−1 g−1 FW at 0, 20, 200, and 2,000 μg butachlor L⁻¹, respectively (Figure 5A). The activity of APX in V. natans seeds also changed to 48, 47, 58, and 16 U min−1g−1 FW under the same treatment conditions (Figure 5B). The greatest reductions in both enzyme activities compared with the control were observed at 2,000 μg L−1. In contrast, SOD activity showed no differences across butachlor concentrations (Figure 5C).
Figure 5. Activities of catalase (CAT), ascorbate peroxidase (APX), and superoxide dismutase (SOD) in Vallisneria natans seeds under different concentrations of butachlor: (A) CAT activity (U min−1g−1 FW); (B) APX activity (U min−1g−1 FW); and (C) SOD activity (U min−1g−1 FW)). Significant differences from the control are shown by * (P < 0.05) level compared with control; ** (P < 0.01). n = 3. Error bars represent the standard deviation.
Changes in GSH and Pro Contents in Vallisneria natans Seeds
The GSH content in seeds at the conditions of 0, 20, 200 and 2,000 μg L−1 after 72 hours was 14, 21, 17 and 15 µmol g−1 FW respectively (Figure 6A). The greatest increase compared with the control was observed at 20 μg L−1, where GSH content was 6.5 µmol g−1 FW higher. There was no difference in the Pro content of V. natans seeds in response to different concentrations of butachlor (Figure 6B).
Figure 6. The glutathione (GSH) and proline (Pro) contents in Vallisneria natans seeds under different concentrations of butachlor: (A) GSH content (μmol g−1 FW) and (B) Pro content (mg g−1 FW). Significant differences from the control are shown by * (P < 0.05) level compared with control; ** (P < 0.01). n = 3. Error bars represent the standard deviation.
Relationship between Physiological Indexes and Germination Ability in Vallisneria natans Seeds
Positive relationships were observed between APX activity and V. natans seed germination percentage (Figure 7A), as well as between APX activity and germination index (Figure 7B). The strongest positive relationships were observed at higher APX activity levels. MDA content was positively correlated with butachlor concentration in the seeds (Figure 7C) and the solution (Figure 7D). In contrast, a negative correlation was observed between H2O2 content and CAT activity in V. natans seeds (Figure 7E).
Figure 7. Relationship between physiological indices and germination ability in Vallisneria natans seeds: (A) ascorbate peroxidase (APX) activity vs. germination percentage; (B) APX activity vs. germination index; (C) malondialdehyde (MDA) content vs. butachlor content in seeds; (D) MDA content vs. butachlor content in solution; and (E) hydrogen peroxide (H2O2) content vs. catalase (CAT) activity in seeds.
Effects of Butachlor on the Germination of Vallisneria natans Seeds
Seed germination rate is a critical indicator for assessing seed viability and the capacity of seeds to establish under varying environmental conditions (Wang et al. Reference Wang, Wang, Li, Gao, Rao, Song and Li2023). Our germination experiments revealed that seed germination rates declined to 23%, 23%, 17%, and 14%, and the germination index to 1.6, 1.7, 1.2, and 0.9, at 0, 20, 200, and 2,000 μg L−1 butachlor, respectively. Both indicators were inhibited at 200 and 2,000 μg L−1. This dose-dependent suppression indicated that butachlor residues could impair the early developmental stages of aquatic plants. Our findings were consistent with previous studies that have documented similar herbicide-induced inhibitory effects on seed germination in terrestrial plants. For example, Kumar and Jagannath (Reference Kumar and Jagannath2014) observed a reduction in both germination rates and early growth parameters in wheat (Triticum aestivum L.) when exposed to butachlor, with germination percentages decreasing proportionally with increasing herbicide concentrations. Further supporting evidence came from Zhang et al. (Reference Zhang, He, Zhang, Zhang, Zhang and Li2021), who demonstrated that the germination rate, vigor, and vitality index of seeds from perennial ryegrass (Lolium perenne L.), alfalfa (Medicago sativa L.), and sainfoin (Onobrychis viciifolia Scop.) were suppressed following exposure to glufosinate and glyphosate. Our results demonstrated that butachlor can inhibit the germination of V. natans seeds, particularly at higher concentrations (200 and 2,000 μg L−1). This dose-dependent response aligned with previous findings in terrestrial species, indicating that butachlor exerts a broad phytotoxic effect across plant types. The sensitivity observed in V. natans suggested that early developmental stages of submerged macrophytes were especially vulnerable to butachlor exposure. Such inhibition of germination under herbicide stress reduced seedling recruitment and regeneration capacity, ultimately affecting population persistence in aquatic habitats.
Effects of Butachlor on Antioxidant Product and the Antioxidant Defense System in Vallisneria natans Seeds
Plants under stress conditions produce ROS that exhibit dual roles: acting as signaling molecules at low concentrations while causing oxidative damage upon excessive accumulation (Mittler Reference Mittler2002). Our study demonstrated that butachlor exposure increased H2O2 content (0.1 μmol g−1 FW higher than control) and MDA levels (5.7 nmol g−1 FW elevation) in V. natans seeds, indicating oxidative stress as a key mechanism for germination inhibition. These findings align with reports on ROS-mediated seed viability reduction (McDonald Reference McDonald1999), while the observed activation of antioxidant enzymes such as SOD, CAT, and APX can be explained by H2O2-induced lipid peroxidation (Lehner et al. Reference Lehner, Mamadou, Poels, Come, Bailly and Corbineau2008). As a biomarker of lipid peroxidation, MDA accumulation directly reflects the degree of membrane lipid peroxidation in plants (Tripthi et al. Reference Tripthi, Varma, Singh, Sachan, Guerriero, Kushwaha, Bhardwaj, Ramawat, Sharma, Singh, Prasad, Chauhan, Dubey and Sahi2020). Our study revealed that butachlor exposure disrupted redox homeostasis in V. natans seeds through two distinct phases: concentration-dependent effects and temporal dynamics. For concentration-dependent effects, high butachlor concentrations (2,000 μg L−1) rapidly induced H2O2 and MDA accumulation, consistent with findings in gramineous plants (Wang et al. Reference Wang, Li and Lin2013). In contrast, low concentrations (200 μg L−1) transiently activated antioxidant enzymes, leading to H2O2 scavenging during early exposure. For temporal dynamics, prolonged exposure (48 h) overwhelmed the antioxidant defense system, corroborating Jin et al.’s (Reference Jin, Zhang, Ahmed, Rasheed, Naeem, Ye and Zhou2010) report on herbicide-induced enzymatic disruption. However, the observed late-phase MDA decline (Liu et al. Reference Liu, Mu, Zhao and Zhou2015) suggests partial ROS homeostasis recovery capacity.
APX activity in V. natans seeds declined within 12 h of butachlor exposure, indicating that this herbicide inhibits APX and impairs H2O2 metabolism. In contrast, CAT and SOD activities increased with higher butachlor concentrations and prolonged exposure, suggesting a compensatory response to oxidative stress. These enzymes are part of the antioxidant defense system that regulates ROS levels and protects against cellular damage (Jiang et al. Reference Jiang, Qiu, Li, Li and Yin2014; Passardi et al. Reference Passardi, Penel and Dunand2004). Additionally, nonenzymatic antioxidants such as GSH also responded to butachlor stress, particularly at low concentrations, consistent with their role in maintaining redox homeostasis and mitigating oxidative damage (Mostofa et al. Reference Mostofa, Hossain, Fujita and Tran2015; Singh et al. Reference Singh, Srivastava and Prasad2013). These enzymatic and nonenzymatic antioxidants function cooperatively, partly via the ascorbate–GSH cycle, to regulate ROS levels and maintain redox balance (Arias-Baldrich et al. Reference Arias-Baldrich, de la Osa, Bosch, Ruiz-Ballesta, Monreal and Garcia-Maurino2017; Foyer and Noctor Reference Foyer and Noctor2011).
Our findings showed that Pro and GSH levels increased in V. natans seeds in response to butachlor exposure, indicating their role in maintaining redox balance under oxidative stress. Pro levels rose after 12 h of exposure to 20 μg L−1 butachlor, suggesting that Pro contributes to early stress adaptation by stabilizing cellular structures and mitigating ROS-induced damage (Sharma and Dietz Reference Sharma and Dietz2006). Similarly, GSH levels increased at 200 μg L−1 and higher concentrations after 12 h, supporting its role in ROS scavenging and redox regulation (Dresler et al. Reference Dresler, Strzemski, Kovacik, Sawicki, Staniak, Wojciak, Sowa and Hawrylak-Nowak2020; Noctor et al. Reference Noctor, Mhamdi, Chaouch, Han, Neukermans, Marquez-Garcia, Queval and Foyer2012). Together, these responses reflect the involvement of both enzymatic and nonenzymatic antioxidant systems in protecting V. natans seeds from butachlor-induced oxidative stress.
Physiological Mechanisms of Butachlor Affecting Vallisneria natans Seed Germination
Although further studies are needed to confirm the specific signaling pathways involved, our results suggested that butachlor inhibited V. natans seed germination by disrupting redox homeostasis and impairing antioxidant defenses. The initial increase in SOD activity, followed by significant inhibition of APX and CAT activities, indicates an imbalance in ROS detoxification. This sequential response may lead to excess H2O2 accumulation, forming a “H2O2 trap” where ROS production exceeds scavenging capacity.
As oxidative stress intensifies, lipid peroxidation occurs, as evidenced by increased MDA levels. MDA is a marker of membrane damage, and its accumulation suggests compromised cellular integrity during early germination. Antioxidant enzyme inhibition and membrane lipid peroxidation together explain the reduced germination observed under butachlor exposure (Figure 8).
Figure 8. Mechanism of butachlor’s effect on the germination of Vallisneria natans seeds. Abbreviations: APX, ascorbate peroxidase; CAT, catalase; GSH, glutathione; MDA, malondialdehyde; MDHA, monodehydroascorbate; O2 −, superoxide anion; Pro, proline; SOD, superoxide dismutase.
Butachlor disrupts the physiological processes of V. natans seeds by inhibiting key antioxidant enzymes, particularly APX and CAT, leading to the accumulation of H2O2. The excess H2O2 induces oxidative damage through membrane lipid peroxidation, as indicated by increased levels of MDA. Prolonged oxidative stress from butachlor exposure compromises membrane integrity, reduces metabolic activity, and ultimately decreases germination rates. These findings highlight the critical role of the antioxidant defense system in protecting aquatic plants from herbicide-induced oxidative damage. Future research should focus on understanding the long-term ecological impacts of butachlor and other herbicides on aquatic ecosystems, particularly their effects on plant community dynamics and ecosystem stability. Prospective studies could incorporate a more comprehensive dose range to better characterize the dose–response relationship, particularly within the 20 to 200 μg L−1 transition zone, while concurrently employing tetrazolium viability assays and seed histological analysis to assess impacts on seed vigor and internal structures.
Acknowledgments
The authors are grateful to Stephen Maberly for his assistance in improving the language of the article and to Wenmin Huang for her guidance and valuable suggestions on the experiment. Finally, we thank two anonymous reviewers for critical and constructive review of the original draft that helped improve the article.
Funding statement
This research was supported by the National Natural Science Foundation of China (32171575, 42201436).
Competing interests
The authors declare no conflicts of interest.
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