Green-synthesized ZnO nanoparticles combined with fulvic and ascorbic acid strengthen antioxidant responses in sweet corn under salinity stress

Soil salinity raises reactive oxygen species and limits crop performance. Green-synthesized zinc oxide nanoparticles (ZnO NPs) may help, but whether combining them with fulvic acid or ascorbic acid adds real benefit is unclear. We measured oxidative stress and antioxidant responses in sweet corn under non-saline and saline conditions across nine treatments: an untreated control, foliar or irrigation-applied ZnO NPs, ZnO NPs combined with fulvic acid or ascorbic acid, and each organic acid applied alone. We tracked leaf hydrogen peroxide (H 2 O 2), superoxide dismutase (SOD), catalase (CAT), and peroxidase (POD). Salinity, treatment, and their interaction significantly affected H 2 O 2, SOD, and CAT (p < 0.001). POD showed significant treatment and interaction effects (p < 0.001) but no significant salinity main effect (p = 0.135). Salinity raised H 2 O 2 in the untreated control by 68.5%. Under salinity, irrigation-applied ZnO NPs with ascorbic acid cut H 2 O 2 by 66.2%, and the foliar combinations with fulvic acid and ascorbic acid cut it by 65.6% and 64.8%. Foliar ZnO NPs with ascorbic acid raised SOD by 41% and CAT by 132.6% relative to the saline control, giving the strongest combined SOD-CAT response. Foliar ZnO NPs combined with ascorbic acid produced high SOD and CAT responses and low H 2 O 2 accumulation. Foliar ZnO NPs combined with fulvic acid and irrigation-applied ZnO NPs combined with ascorbic acid also produced significant biochemical responses under salinity. These responses depended on formulation and application method.

Salinity reduces soil water potential, thereby restricting water uptake by plant roots. As Na + and Cl - build up in the plant, they disrupt K + retention, enzyme function, membrane integrity, and photosynthesis. These disturbances increase the production of reactive oxygen species (ROS), including superoxide and H 2 O 2. At low levels, ROS act as stress signals. Once the plant can no longer keep up with detoxifying them, they damage lipids, proteins, pigments, and DNA (Mittler, 2002; Sachdev et al., 2021; Singh et al., 2022). Plants regulate ROS through coordinated enzymatic and non-enzymatic antioxidant systems. SOD turns superoxide into H 2 O 2. CAT, ascorbate peroxidase, and other peroxidases then break that H 2 O 2 down or redirect it (Gill and Tuteja, 2010). Ascorbate, glutathione, phenolics, and related compounds contribute to antioxidant redox regulation, while the ascorbate–glutathione cycle links redox state with stress signaling (Foyer and Noctor, 2011). Looking at just one enzyme can be misleading. Evaluating H 2 O 2 together with several enzyme assays provides a broader characterization of leaf antioxidant biochemistry without extending the interpretation to unmeasured physiological or agronomic outcomes. Zinc plays a structural role in proteins, membranes, gene transcription, and Cu/Zn-SOD. It is also commonly deficient in agricultural soils, which is why cereal breeders and agronomists already pursue Zn biofortification (Cakmak and Kutman, 2018). ZnO NPs can serve as a highly reactive zinc source and may shift stress signaling and nutrient status as well. Their effects still depend heavily on particle size, surface chemistry, dissolution, dose, crop species, and delivery route, and mechanistic reviews stress that zinc-based nanoparticles need formulation-specific dose tuning to avoid phytotoxicity (Khan et al., 2023). In salt-stressed common bean, foliar spray, nano-priming, and soil application each produced a different physiological and ion-balance response, with nano-priming the most consistent (Gupta et al., 2024). Previous work reported changes in antioxidant status after foliar application of biosynthesized ZnO NPs in faba bean (Mogazy and Hanafy, 2022), and in chilli, 100 ppm biogenic ZnO NPs lowered H 2 O 2 by 33.1% and malondialdehyde by 54.4% under salinity (Adnan et al., 2025). These reported biochemical responses also indicate the need for formulation- and route-specific testing. Ascorbic acid and fulvic acid may influence ZnO NP formulations through different biochemical and physicochemical pathways. Ascorbate scavenges ROS directly and keeps the ascorbate–glutathione cycle running (Broad et al., 2020). Previous studies reported changes in antioxidant activity, osmotic adjustment, and ion balance after external application in salt-stressed barley and pea (Hassan et al., 2021; Kanwal et al., 2024), and recent quinoa work supports a similar role in salinity acclimation (Shah et al., 2025). Fulvic acid works differently: it chelates nutrients, changes their mobility, supports osmotic adjustment, and shapes rhizosphere activity. In salt-stressed citrus, potassium fulvic acid improved pigment status, antioxidant-related traits, soil enzyme activity, and rhizosphere microbial composition (Zhang et al., 2023).

Although ZnO nanoparticles (ZnO NPs), fulvic acid, and ascorbic acid have been studied separately under abiotic stress, their combined effects are less clear. Few studies have compared these treatments under the same crop, salinity level, ZnO NP dose, and sampling conditions, especially in sweet corn. Ascorbic acid may directly support ROS scavenging, while fulvic acid may affect Zn complexation, nutrient mobility, nanoparticle behavior, and root-zone processes. The application route may also influence these responses. Foliar application involves leaf retention and penetration, whereas irrigation involves soil adsorption, nanoparticle transformation, rhizosphere interactions, and root uptake. This study directly compared ZnO NPs alone, ZnO NPs with fulvic acid, ZnO NPs with ascorbic acid, and the available organic-acid-alone treatments under saline and non-saline conditions. Both foliar and irrigation applications were evaluated. The study focused on changes in H 2 O 2, SOD, CAT, and POD. The design allows formulation and application dependent responses to be identified, but it does not provide a formal test of biochemical synergy.

We hypothesized that ZnO NPs combined with fulvic acid or ascorbic acid would reduce leaf H 2 O 2 and alter SOD, CAT, and POD responses, particularly under salinity. These responses were expected to differ from the untreated control and from the individual ZnO NP and organic acid treatments. The ascorbic acid formulation was expected to favor H 2 O 2 removal because ascorbate directly participates in ROS scavenging and the ascorbate–glutathione cycle. In contrast, the fulvic acid formulation was expected to produce a different antioxidant response by modifying Zn availability, nutrient mobility, nanoparticle dispersion, and root-zone interactions. We also expected foliar and irrigation applications to produce different biochemical responses because they involve different uptake and transformation pathways.

Plant material, growth conditions, and experimental design

The pot experiment was conducted at the Organic Garden at Florida International University. Sweet corn (Zea mays L.) cultivar ‘Golden Bantam’ was grown in 10-inch pots containing sandy loam soil with an initial electrical conductivity of <1 dS m - ¹. The experiment followed a completely randomized factorial design with salinity condition and treatment as fixed factors. Plants were grown under non-saline or saline conditions. For the saline treatment, NaCl was added to the soil before sowing. Soil electrical conductivity (EC) was checked three days later. A 50 mM NaCl solution was then applied at two-week intervals, and EC was checked regularly to maintain a moderate salinity level of 2–3 dS m - ¹. Salt exposure therefore began before sowing and continued until harvest. The total exposure period was three months. Leachate was not monitored. Salt accumulation was assessed only through the regular soil EC measurements. EC was measured using a FieldScout TDR 250 meter.

Green synthesis of ZnO nanoparticles

Green ZnO NPs were synthesized using avocado (Persea americana) seed extract via a co-precipitation method. Avocado seeds were collected from the USDA Agricultural Research Service (USDA-ARS), Subtropical Horticulture Research Station. Zinc acetate Zn(CH 3 COO) 2 analytical grade, 99% purity; Sigma-Aldrich) was used as the zinc precursor. Sodium hydroxide (NaOH, pellets), ascorbic acid, and fulvic acid were purchased from Fisher Scientific (USA). A 0.01 M Zn(CH 3 COO) 2 solution was prepared in 90 mL of deionized (DI) water and heated to 70 °C under continuous stirring. Subsequently, 10 mL of avocado seed extract was added dropwise to the Zn(CH 3 COO) 2 solution. A 0.1 M NaOH solution was then added gradually until the pH reached ~11.0. The resulting white zinc hydroxide precipitate was maintained at 70 °C under continuous stirring for 5 h to convert into ZnO NPs. The synthesized NPs were collected by centrifugation at 13,000 rpm for 20 min and washed repeatedly with DI water to remove residual reactants and impurities.

Preparation and application of ZnO NPs

ZnO NP suspensions were prepared at a concentration of 200 mg L - ¹. Combined formulations contained either 0.01% fulvic acid or 0.01% ascorbic acid. All plant-treatment formulations were adjusted to pH 8.0. The ZnO NP suspensions were sonicated for 2 min and applied immediately after preparation. The suspensions were not stored before application, and no visible settling was observed during the preparation and application period. Treatments were applied three times at 3-week intervals after germination with cumulative dose of 1mg per plant. The treatments were designated as follows: ZT1, untreated control; ZT2, foliar application of green-synthesized ZnO NPs; ZT3, irrigation application of green-synthesized ZnO NPs; ZT4, foliar application of green-synthesized ZnO NPs combined with fulvic acid; ZT5, irrigation application of green-synthesized ZnO NPs combined with fulvic acid; ZT6, foliar application of green-synthesized ZnO NPs combined with ascorbic acid; ZT7, irrigation application of green-synthesized ZnO NPs combined with ascorbic acid; ZT8, irrigation application of 0.01% fulvic acid and ZT9 irrigation application of 0.01% ascorbic acid (Table 1). Each experimental unit consisted of one pot containing one plant. Four independent pots were assigned to each treatment × salinity combination, providing four biological replicates (n = 4 pots) per treatment × salinity combination. With nine treatments and two salinity conditions, the experiment comprised a total of 72 pots. Where two measurements were obtained from an individual biological replicate, the measurements were averaged before statistical analysis; therefore, inferential analyses were based on four independent biological replicates per treatment x salinity combination. Leaf samples were collected before harvest, washed, placed on ice, and processed under cold conditions, as shown in Table 2. Pots were arranged according to a completely randomized design and repositioned regularly throughout the experiment to minimize potential positional effects.

Hydrogen peroxide content

H 2 O 2 content was measured with the potassium iodide method of Velikova et al. (2000). Fresh leaf tissue (0.50 g) was homogenized in 5 mL of ice-cold 0.1% trichloroacetic acid. The homogenate was centrifuged at 12,000 × g for 15 min. The reaction contained 0.50 mL of supernatant, 0.50 mL of 10 mM potassium phosphate buffer at pH 7.0, and 1.0 mL of 1 M potassium iodide. The mixture was kept in darkness before absorbance was measured at 390 nm. Reagent and sample blanks were used. H 2 O 2 was quantified from a 0–50 µM standard curve and expressed as µmol g - ¹ fresh weight.

Superoxide dismutase activity

SOD activity was measured by the NBT photoreduction assay of Beauchamp and Fridovich (1971). Fresh leaf tissue was extracted in 50 mM phosphate buffer at pH 7.8 containing 0.1 mM EDTA and 1% polyvinylpyrrolidone. The reaction mixture contained phosphate buffer, methionine, EDTA, NBT, riboflavin, and enzyme extract. The reaction was initiated by adding riboflavin and exposing the mixture to fluorescent light for 10 min. Absorbance was measured at 560 nm. A light control without enzyme extract and a dark control were included. SOD activity was reported as the percentage inhibition of NBT photoreduction.

Catalase activity

CAT activity was determined with a molybdate endpoint assay adapted from Aebi (1984). Fresh leaf tissue was homogenized in 50 mM phosphate buffer at pH 7.0 containing 1% polyvinylpyrrolidone. The extract was centrifuged at 12,000 × g for 15 min at 4 °C. The reaction contained phosphate buffer, 10 mM H 2 O 2, and enzyme extract. Sodium molybdate was added after 1 min to stop the reaction and react with residual H 2 O 2. Absorbance was measured at 405 nm. CAT activity was expressed as the percentage of H 2 O 2 degraded relative to the no-enzyme control.

Peroxidase activity

POD activity was measured with guaiacol as the substrate (Chance and Maehly, 1955). Fresh leaf tissue was homogenized in 50 mM phosphate buffer containing 1% polyvinylpyrrolidone. The extract was centrifuged at 12,000 × g for 15 min at 4 °C. The reaction contained phosphate buffer, 20 mM guaiacol, 40 mM H 2 O 2, and enzyme extract. Absorbance at 470 nm was recorded at 9, 63, 117, and 171 s, corresponding to 0.15, 1.05, 1.95, and 2.85 min. POD activity was calculated as the linear regression slope and expressed as ΔA 470 min - ¹. One operational unit represented an absorbance increase of 0.01 min - ¹ under the assay conditions. All time-course regressions met the predefined linearity criterion of R² ≥ 0.95. Enzyme responses were expressed using assay-based activity indices because protein concentration was not determined for the extracts. Therefore, SOD, CAT, and POD values represent relative assay responses under standardized extraction and reaction conditions rather than protein-normalized specific enzyme activities. Comparisons were restricted to samples processed using the same tissue mass, extraction procedure, and assay conditions.

Statistical analysis

Data are presented as mean ± standard deviation. The effects of salinity, treatment, and their interaction were tested using Type II two-way analysis of variance, as shown in Table 2. Separate one-way analyses were conducted within each salinity condition, followed by Tukey’s honestly significant difference test at α = 0.05. Tukey groups were calculated separately for saline and non-saline conditions. Means sharing at least one letter were not significantly different.

Residual normality and variance homogeneity were evaluated using the Shapiro–Wilk test and median-centered Levene test, respectively. Outlier and influence diagnostics were used for quality control screening. No observations were excluded. The analysis included four biological replicates per treatment × salinity combination. The full factorial model had 54 residual degrees of freedom. All tables, figures, and statistical tests were generated from the same replicate-level dataset archived in Zenodo. Percentage changes were calculated relative to the untreated control within the same salinity condition. Biological interpretation considered the means, standard deviations, percentage changes, and Tukey groups together. Numerical differences were not considered biologically meaningful when treatments shared a Tukey group or when the differences were small relative to the observed variability. Analyses were performed in Python version 3.12.13 using SciPy version 1.16.3 and statsmodels version 0.14.6. Data were managed using pandas version 2.2.2, and figures were generated using Matplotlib version 3.10.0. For Figures 1–4, bar heights and error bars represent the replicate-level mean and standard deviation, respectively, and the displayed letters are the compact-letter representation of the condition-specific Tukey HSD pairwise comparisons.

Characterization of ZnO nanoparticles

The structural and morphological characteristics of the synthesized ZnO NPs were investigated using powder X-ray diffraction (PXRD; Bruker D6 PHASER) and high-resolution transmission electron microscopy (HR-TEM; JEOL JEM-2100Plus), as shown in Figure 1. The PXRD pattern confirmed the formation of crystalline hexagonal wurtzite ZnO, with characteristic diffraction planes indexed to (100), (002), (101), (102), (110), (103), (200), (112), (201), (004), and (202), consistent with the standard JCPDS card No. 80-0075 (space group P6 3 mc). The average crystallite size, calculated using the Scherrer equation, was ~18 nm. In contrast, HR-TEM images revealed quasi-spherical particles with an approximate apparent diameter of ~100 nm estimated visually from a representative TEM micrograph. The difference between the PXRD crystallite size and the TEM observed particle size suggests that the individual particles may consist of multiple crystalline domains and/or that particle aggregation contributed to the larger particle size observed by TEM.

Hydrogen peroxide content

Salinity increased H 2 O 2 concentration in the untreated control (ZT1) by 68.5%, from 0.169 ± 0.002 to 0.284 ± 0.003 µmol g - ¹ FW. All treatments significantly reduced H 2 O 2 relative to ZT1 under the corresponding salinity condition (p < 0.05; Figure 2). Under non-saline conditions, foliar ZnO NPs combined with fulvic acid (ZT4) and foliar ZnO NPs combined with ascorbic acid (ZT6) produced the largest reductions, 71.8% and 70.3%, respectively, and formed the lowest statistical group. Under salinity, irrigation-applied ZnO NPs combined with ascorbic acid (ZT7), ZT4, and ZT6 reduced H 2 O 2 by 66.2%, 65.6%, and 64.8%, respectively. Their concentrations ranged from 0.096 to 0.100 µmol g - ¹ FW and did not differ significantly. Ascorbic acid alone (ZT9) was the least effective treatment under salinity, with an H 2 O 2 concentration of 0.151 ± 0.001 µmol g - ¹ FW, although it remained lower than ZT1.

Superoxide dismutase activity

Most treatments significantly increased SOD activity relative to the untreated control (ZT1) under the corresponding salinity condition (p < 0.05; Figure 3). Foliar ZnO NPs combined with ascorbic acid (ZT6) produced the highest numerical activity under both conditions. Under non-saline conditions, ZT6 increased SOD activity by 33.7% and was statistically comparable with irrigation-applied ZnO NPs combined with fulvic acid (ZT5), irrigation-applied ZnO NPs combined with ascorbic acid (ZT7), fulvic acid alone (ZT8), and ascorbic acid alone (ZT9). Under salinity, ZT6 increased SOD activity by 41.0%, reaching 90.57 ± 1.21% inhibition, and was statistically comparable with irrigation-applied ZnO NPs alone (ZT3), ZT5, and ZT7. Fulvic acid (ZT8) and ascorbic acid (ZT9) applied alone were less effective under salinity than most ZnO NP combinations. ZT9 maintained significantly higher SOD activity than ZT1, whereas ZT8 did not differ significantly from ZT1. For SOD, the effects of treatment [F(8,54) = 78.68], salinity [F(1,54) = 39.07], and their interaction [F(8,54) = 24.33] were significant (p < 0.001).

Catalase activity

CAT activity showed the largest relative response to treatment (Figure 4). Under non-saline conditions, ZT6 produced the highest CAT activity (51.13 ± 3.41% H 2 O 2 degraded), compared with 2.59 ± 1.36% in ZT1, and significantly exceeded the other treatments. Foliar ZnO NPs combined with fulvic acid (ZT4) produced the second-highest activity, whereas foliar ZnO NPs (ZT2), irrigation-applied ZnO NPs (ZT3), and ascorbic acid alone (ZT9) did not differ significantly from ZT1. Under salinity, ZT6, ZT4, and irrigation-applied ZnO NPs combined with ascorbic acid (ZT7) formed the highest Tukey group, increasing CAT activity by 132.6%, 124.3%, and 122.6%, respectively, relative to the saline ZT1. ZT9 remained statistically comparable with the saline ZT1.

Peroxidase activity

POD activity showed a pronounced treatment-specific response (Figure 5). Under non-saline conditions, ascorbic acid alone (ZT9) and irrigation-applied ZnO NPs combined with fulvic acid (ZT5) produced the highest activities, reaching 0.351 ± 0.006 and 0.332 ± 0.013 ΔA 470 min - ¹, respectively, and did not differ significantly. Foliar ZnO NPs combined with ascorbic acid (ZT6) produced 0.195 ± 0.017 ΔA 470 min - ¹ and significantly exceeded foliar ZnO NPs combined with fulvic acid (ZT4; 0.149 ± 0.014 ΔA 470 min - ¹). Fulvic acid alone (ZT8; 0.179 ± 0.020 ΔA 470 min - ¹) was intermediate and did not differ significantly from either ZT6 or ZT4. Foliar ZnO NPs alone (ZT2), irrigation-applied ZnO NPs alone (ZT3), and irrigation-applied ZnO NPs combined with ascorbic acid (ZT7) formed a lower statistical group, with activities ranging from 0.081 to 0.109 ΔA 470 min - ¹. The untreated control (ZT1) had the lowest activity, at 0.033 ± 0.003 ΔA 470 min - ¹.

Under saline conditions, ZT9 maintained the highest POD activity (0.323 ± 0.034 ΔA 470 min - ¹). ZT4 was statistically comparable with ZT9, with an activity of 0.277 ± 0.007 ΔA 470 min - ¹. ZT5 produced 0.261 ± 0.028 ΔA 470 min - ¹ and was statistically comparable with ZT4. ZT7 showed an intermediate response of 0.210 ± 0.043 ΔA 470 min - ¹. ZT2 produced 0.098 ± 0.017 ΔA 470 min - ¹ and did not differ significantly from ZT1 (0.054 ± 0.008 ΔA 470 min - ¹). ZT3, ZT6, and ZT8 produced 0.079 ± 0.015, 0.066 ± 0.027, and 0.084 ± 0.016 ΔA 470 min - ¹, respectively, and did not differ significantly from ZT1. The significant salinity × treatment interaction primarily reflected increased POD activity in ZT4 and ZT7 and decreased activity in ZT5, ZT6, and ZT8, while ZT9 remained consistently high across both salinity conditions.

Salinity increased leaf H 2 O 2 in untreated plants by 68.5%, indicating disruption of cellular redox balance. Salt stress can increase ROS production by disturbing water relations, ion balance, and energy metabolism (Mittler, 2002; Bose et al., 2014). Excess ROS can damage membranes, proteins, pigments, and nucleic acids (Sachdev et al., 2021; Singh et al., 2023). However, H 2 O 2 also functions as a signaling molecule. Its concentration should therefore be interpreted together with antioxidant enzyme responses. SOD converts superoxide to H 2 O 2, while CAT and peroxidases contribute to H 2 O 2 removal (Gill and Tuteja, 2010). Under salinity, ZT4, ZT6, and ZT7 reduced H 2 O 2 by 64.8–66.2% relative to the untreated control. Their H 2 O 2 concentrations were statistically comparable. These treatments also produced high SOD or CAT responses. The combined results indicate altered antioxidant regulation rather than superiority of one formulation. Zinc may contribute to these responses through its functions in membrane stability, enzyme activity, and Cu/Zn-SOD (Cakmak and Kutman, 2018). ZnO NP effects also depend on dose, formulation, dissolution, aggregation, and application route (Khan et al., 2023). In salt-stressed maize, foliar ZnO NPs affected antioxidant activity, Na + and K + accumulation, photosynthesis, growth, and yield (Seleiman et al., 2023). In common bean, foliar application, soil application, and seed priming produced different antioxidant, growth, and nutrient responses (Gupta et al., 2024). Biosynthesized ZnO NPs also modified antioxidant and physiological responses in faba bean (Mogazy and Hanafy, 2022). Similar responses were reported in pea and rice, where ZnO NP treatments affected antioxidant activity, photosynthetic traits, Zn uptake, and growth under salinity (Mustafa et al., 2024; Dang et al., 2024). These studies provide physiological context for the present biochemical findings. However, the corresponding physiological responses were not measured here.

The ZnO NP ascorbic acid treatments produced low H 2 O 2 concentrations and high SOD or CAT responses under salinity. Ascorbic acid can scavenge ROS and participate in the ascorbate–glutathione cycle (Foyer and Noctor, 2011; Broad et al., 2020). This mechanism may have contributed to the observed response. However, the present design cannot separate direct ascorbate effects from changes in Zn availability or formulation behavior. It also does not establish synergy between ZnO NPs and ascorbic acid. Fulvic acid produced different responses with foliar and irrigation applications. ZT4 combined low H 2 O 2 with high CAT and POD responses. ZT5 maintained high SOD and POD responses. Fulvic substances can affect nutrient mobility, osmotic regulation, and rhizosphere processes (Zhang et al., 2023). Foliar application exposes leave directly to the formulation. Irrigation introduces the formulation into soil, where minerals, organic matter, root exudates, and microorganisms may alter aggregation, dissolution, and Zn availability. These pathways may explain the contrasting responses of ZT4 and ZT5. Similar application-dependent effects have been reported for ZnO NPs in common bean (Gupta et al., 2024). Studies of green-synthesized lignin nanoparticles also show the value of physiological and structural validation. These nanoparticles altered antioxidant metabolism and reduced salinity-associated ultrastructural damage in maize (Javaid et al., 2024). The present experiment did not assess cellular ultrastructure. Therefore, the measured antioxidant responses cannot be used to infer changes in cellular structure. POD showed a different pattern from H 2 O 2, SOD, and CAT. The salinity main effect was not significant (p = 0.135), but the salinity × treatment interaction was significant (p < 0.001). POD increased under salinity in ZT4 and ZT7 but decreased in ZT5, ZT6, and ZT8. ZT9 remained high under both conditions. Guaiacol peroxidase can participate in H 2 O 2 metabolism, phenolic oxidation, and cell-wall processes. A high POD response alone does not indicate stronger antioxidant protection. It should be interpreted with H 2 O 2, SOD, and CAT.

Several limitations restrict the mechanistic interpretation. Tissue Zn, Na + /K + balance, oxidative damage, nanoparticle uptake, dissolution, localization, photosynthesis, biomass, and yield were not measured. The responses therefore cannot be attributed exclusively to nanoparticulate ZnO. Contributions from dissolved Zn and residual seed-derived compounds cannot be excluded. The results support formulation-, salinity-, and application-dependent changes in biochemical antioxidant responses. The interpretation is restricted to the measured antioxidant variables and does not extend to growth, yield, photosynthesis, or other unmeasured plant responses. Field applicability was not evaluated because the study used controlled pot conditions and did not include field-level validation.

Green-synthesized ZnO NP formulations altered H 2 O 2 accumulation and antioxidant assay responses in sweet corn under salinity. Several combined treatments produced low H 2 O 2 and high SOD or CAT responses. POD followed a distinct treatment-dependent pattern. These results show formulation- and application-dependent biochemical responses. The conclusions are restricted to H 2 O 2, SOD, CAT, and POD responses. Growth, yield, photosynthesis, and nanoparticle uptake require separate measurement.