Resulting Key Physiological Changes in Triticum aestivum L. Plants Under Drought Conditions After Priming the Seeds with Conventional Fertilizer and Greenly Synthesized Zinc Oxide Nanoparticles from Corn Wastes

Abstract

This research study investigated the production and properties of zinc oxide (ZnO) nanoparticles derived from corn husks and their priming effects on wheat plant proliferation and antioxidant mechanisms compared to the nutri-priming technique under regular irrigation and drought-stressed conditions. Transmission and scanning electron microscopy (TEM and SEM), energy-dispersive X-ray spectroscopy (EDAX), and X-ray diffraction confirmed the nanoparticles’ hexagonal morphology and typical dimensions of 51 nm. The size and stability of these nanoparticles were assessed through the size distribution and zeta potential analysis, indicating reasonable stability. Fourier-transform infrared spectroscopy (FTIR) detected the newly formed functional groups. This study emphasized the role of reactive oxygen species (ROS) and phenolic compounds in plant responses to nanoparticle treatment, particularly in detoxifying harmful radicals. The research also examined the activity of antioxidant enzymes, including peroxidase (POX), catalase (CAT), and glutathione reductase (GR), in alleviating stress caused by oxidation while subjected to various treatments, including micronutrient seed priming with DR GREEN fertilizer. Some biochemical compounds, such as total phenolics (TPCs), total flavonoids (TFCs), and total hydrolysable sugars, were estimated as well to show the effect of the different treatments on the wheat plants. The findings suggested that ZnO nanoparticles can enhance antioxidant enzyme activity under certain conditions while posing phytotoxic risks, underscoring the complexity of plant–nanoparticle interactions and the potential for improving crop resilience through targeted micronutrient applications.

1. Introduction

Rising temperatures from global warming accelerate the water cycle and cause more intense hydrological events. Drought is one of the worst calamities worldwide because it diminishes water availability and increases water resource vulnerability [1]. Drought is an unavoidable stressor that affects crop growth and output [2]. Drought stress alters plant morphology, physiology, and biochemistry [3]. Climate change, human development, and urbanization are making droughts more frequent and destructive. Its economic, natural, and social effects on agriculture are significant. Drought stress at critical growth stages reduces crop productivity worldwide [4]. The annual agricultural losses are about USD 6 billion and rising [5].

This risk is significant, as the European Union (EU) is one of the world’s largest cereal producers, and wheat (Triticum aestivum L.) cereal stands out in terms of cultivated area [6]. Wheat is one of humanity’s staple foods because it is a primary source of calories and proteins [7]. Throughout the meiotic stage, rice and wheat suffered a 35–75% reduction in grain yield due to water deficiency [8]. Wheat yield is affected by drought stress at all growth stages, but the reproductive stage and grain formation are the most critical stages [9]. Brief periods of intense drought stress during blossoming can lead to significant termination of pollen grains and ovules, resulting in irreversible reductions in fertility and grain number per spike [10]. Severe drought stress can result in complete crop failure by abruptly terminating the crop’s life cycle [11], altering protein biosynthesis and other plant metabolites, and altering gene expression [12]. Drought can significantly impair cellular metabolism, ion buildup, the stability of membranes, and the fundamental makeup of proteins [13]. Drought stress causes plants’ molecular, cellular, and physiological disorders by promoting reactive oxygen species (ROS) generation, which leads to loss of cell membrane stability without any protective mechanism [14]. Many technologies, including nanotechnology and seed priming, have arisen to mitigate the detrimental impacts of environmental stress on crops, raise water use efficiency, avert plant diseases, secure food supply, diminish environmental pollution, and promote sustainability [15].

Before planting seeds, seed priming (SP) entails soaking the seeds in water, comprising organic or inorganic salts if desired, under regulated conditions. After that, the seeds are allowed to dry in the shade. Priming, which is highly similar to the process of hydrating seeds, helps plants establish themselves more effectively by reducing the impact of environmental challenges, which eventually results in better crop outputs. Seed priming employs many different techniques to mitigate the detrimental impacts of drought stress. These encompass the first recruitment of seed reserves for nutrition, the enlargement of embryonic cells, and the degradation of endosperm [16]. According to Chen and Arora (2011), these processes boost the metabolic activities that occur prior to germination, which ultimately results in high and consistent germination percentages [17]. Diverse organic and inorganic compounds are applied to seed priming, which benefits crop growth. As a consequence, the SP technology has been categorized into the following categories: water-priming, osmo-priming, oxidation/reduction-dependent priming, chemical priming, hormone priming, nutritional priming, biopriming, and nano-priming [18]. These categories are based on the features and functions of the SP chemicals.

Nanoparticles (NPs) enhance plant protection by augmenting metabolic antioxidant activity to mitigate oxidative damage [19]. NPs modulate salt tolerance in various plant species by regulating hormone levels, antioxidant enzyme activities, ionic homeostasis, and regulation of genes and systems of defense [20]. NPs can readily penetrate plant tissues, enhancing plant function and enabling efficient tolerance to drought and salt stress [21]. NPs improve the membrane’s stability and intake of nutrients and preserve the plant’s apparatus for photosynthesis from damage caused by salinity and drying out stress [22,23]. It was hypothesized that incorporating ZnO-NPs into plants’ seeds with elevated Zn concentrations would enhance germination rates and other growth characteristics. Zinc nanoparticles could potentially influence the activity of antioxidant enzymes [24]. ZnO–nutrient nanoparticles significantly affect plant shape, biological functioning, and biochemistry, enhancing reproduction and boosting crop yields, particularly in drought and saline conditions [25]. Wheat showed noticeable improvement in growth characteristics following the ingestion of ZnO-NPs at specified concentrations [26]. It demonstrated a substantial enhancement in biological attributes and the content of chlorophyll, carotenoids, carbohydrates, and protein contents. Enzymes that fight oxidation, such as peroxidase (POX), superoxide dismutase (SOD), catalase (CAT), and cumulative flavonoid content, also confirmed the effect of nutrition on wheat [27]. The synthesis of nanomaterials through green methods has received significant attention because it is environmentally friendly and sustainable [3].

This study examined the impact of seed priming with ZnO nanoparticles and DR GREEN fertilizer on various biochemical markers, including TPCs, TFCs, and total sugars, as well as enzymatic activities in wheat plants under both normal irrigation and drought stress conditions by assessing specific antioxidative enzymes such as GR, POX, and CAT. Eco-friendly ZnO nanoparticles synthesized from agricultural waste corn husk were studied using various analytical instruments, including TEM, SEM, XRD, EDAX, size distribution analysis, zeta potential measurement, and FTIR. The environmentally synthesized nanoparticles were advantageous due to their reduced harm, lower toxicity, eco-friendliness, and cost-effectiveness compared to alternative synthesis methods. This study also determined the inquiries concerning the relationship between nanoparticle priming and conventional fertilizer DR GREEN (nutri-priming), as it significantly advanced the objective of sustainable development by mitigating the detrimental effects of drought stress, thereby enhancing crop productivity through a safer and more environmentally friendly methodology.

2. Materials and Methods

2.1. A Sustainable Method for Making Zinc Oxide Nanoparticles Using the Aqueous Extract from the Corn Husks

Twenty grams of dried Zea mays husks were measured and rinsed with distilled water at ambient temperature. The husks were immersed in pure water at roughly 80 to 90 °C for one hour to extract the light, translucent, yellowish tint. Two further filtrations using Whatman No. 1 filter paper were conducted on the dye extract solution to eliminate residual particles. The analytical grade hydrated zinc nitrate, with the formula “Zn(NO₃)₂ 6H₂O (catalog No. A16282, lot No. 10242486, Thermo Fisher Scientific Inc., Waltham, MA, USA), was evaluated as a probable zinc precursor. Furthermore, 1–3 g of zinc nitrate were meticulously mixed with 100 mL of Zea mays dried husk extract solution, stirred for twenty minutes until complete dissolution and color change were achieved, and heated at 100 °C for two hours. Shortly after the occurrence of this phase, a precipitation was seen. The precipitate was repeatedly rinsed with milli-Q water (catalog No. 438793, Thermo Fisher Scientific Inc., Waltham, MA, USA) centrifuged at 6000 rpm for fifteen minutes, and dried in an oven at 100 °C for six hours. To achieve highly crystalline ZnO nanoparticles, the dried precipitate was subjected to annealing in an open furnace at 300 °C for two hours [28,29].

2.2. Investigation of Zinc Oxide Nanoparticles Produced via Eco-Friendly Method

At the Micro Analytical Center, Cairo University, Faculty of Science, Giza, Egypt, the crystallinity of ZnO nanoparticles that were environmentally generated was measured with a Philips PW 3710 apparatus (CuKα radiation, Λ = 1.54056 Å, 50 kV, 40 mA, scanning speed 0.02°/s, dwell duration 1 s). In finely powdered samples, back-packed mounts diminished particle orientation. Calcined ZnO nanoparticles were employed for instrumental broadening to calculate the average crystallite size [21,30]. In Giza, Egypt, at the Micro Analytical Center of Cairo University’s Faculty of Science, FTIR spectra were recorded utilizing a Bruker Vertex 70v apparatus (BRUKER OPTIK GmbH, Rudolf-Plank-Straße 27 D-76275 Ettlingen, Germany) equipped with a Diamond ATR module and DTGS detector, achieving a resolution of 2 cm⁻¹. Moreover, 512 images were averaged to obtain the spectra [21,30]. At a 200 kV accelerator potential, the Micro Analytical Center at Cairo University, Faculty of Science, Giza, Egypt, applied scanning and transmission electron microscopy using an X-FEG electron source on a Thermo Fisher Scientific Inc.-issued FEI Talos F200X electron microscope. An EDAX spectrometer analyzed the materials’ composition using a transmission electron microscope. Transmission electron microscopy and scanning techniques were employed. Aqueous milli-Q dispersion droplets were deposited with a delicate carbon coating over a copper grid and then dried at 60 °C for measurement [31]. Hydrodynamic size and zeta potential of green ZnO nanoparticles in water and complete cell culture medium were investigated using dynamic light scattering. DLS evaluated changes in light scattering intensity caused by Brownian motion to ascertain the distribution of particle velocities. This method determines the hydrodynamic radius or diameter via the Stokes–Einstein equation. The particle’s orthogonality to the light source was delineated. The Zeta sizer Nano-ZS employs laser Doppler velocimetry to find the particles’ zeta potential in a solution. Lasers quantified electrophoretic mobility, i.e., the speed of particles influenced by electric fields. The sample analysis apparatus employs an LDV-specific electric field generator and a 4 mW He-Ne 633 nm laser. ZnO nanoparticles were dispersed in deionized water and agitated vigorously for 5 min to achieve homogeneity. Laser Doppler velocimetry used a Malvern Clear Zeta Potential cell to experiment on 1 mL of the fluid. A 25 µg/mL concentration was used to make the solutions [32].

2.3. Seed Priming

To distinguish the impacts of ZnO nanoparticles and DR GREEN fertilizer (PRIME MAGTRÁGYA; P₂O₅: 250 g/kg, K₂O: 170 g/kg, B: 2.5 g/kg, Cu: 1.75 g/kg, Fe: 35 g/kg, Mn: 30 g/kg, Mo: 0.25 g/kg, Zn: 32.5 g/kg, all micronutrients: 102 g/kg, and all active ingredients: 522 g/kg) (Fabryczna, Chrzanów, Poland), seed priming was conducted. Unprimed seeds (untreated seeds) served as the control group. The wheat variety (Kinachi-97) seeds were collected from the agronomy department at the Hungarian University of Agriculture and Life Sciences, Georgikon Campus, Keszthely, Hungary, and then they were sterilized. Untreated seeds functioned as the control group. The wheat variety seeds were sterilized using 0.1% NaClO for 2 min and subsequently cleansed with deionized water to remove all chloride residues. For seed priming, ZnO nanoparticle suspension was recently generated by suspending the fragments in deionized water in a concentration of 50 mg/250 mL ultrapure water, using ultrasonic agitation for 30 min. The seeds were primed in DR GREEN fertilizer in 10 g/250 mL ultra-pure water. Seeds were immersed in darkness at 25 °C for 24 h, with continuous moderate disturbance. The treated seeds were subsequently rinsed with distilled water four times and dried to restore their natural moisture. Seeds were enclosed in polythene bags and maintained at ambient temperature until additional use. Winter wheat seeds of the Triticum aestivum L. were used for the current experiment. Furthermore, 20 cm black plastic planter pots holding a ratio of soil to peat moss were utilized to sow healthy wheat seeds. The seeds of wheat cultivar were sown in the plastic pots on 11 November 2023 at a depth of 1 cm. Twenty grains were sown in each pot at a distance of 2 cm between the two neighboring grains. Using air circulation and mechanical window-opening systems, the greenhouse was maintained at 25 °C during the day and 20 °C at night, with a comparative humidity of between 60 and 65%. Six treatments were applied: T1: (Control, normal irrigation 100% pot capacity (PC)), T2: (Control, drought stress 50% PC), T3: (Normal irrigation 100% PC + seeds were primed with ZnO-NPs), T4: (Drought stress 50% PC + seeds were primed with ZnO-NPs), T5: (Normal irrigation 100% PC + seeds were primed with DR GREEN fertilizer), and T6: (Drought stress 50% PC+ seeds were primed with DR GREEN fertilizer). The 50% pot capacity irrigation was administered to the soil on the 30th day post-sowing to mitigate stress caused by drought (11 December 2023).

2.4. Design, Preparations, and Equipment for Wheat Planting

The pot foundation was coated with 1 kg of pre-washed and dried stones and a plastic net. A perforated mechanism produced three 8 mm apertures at the pot’s base for irrigation and aeration. An empty pot was filled with a soil–peat mixture (1:1 by weight). Black peat and Eutric Cambisol soil samples were collected from the “A” horizon of the research farm area at the Hungarian University of Agriculture and Life Sciences, Georgikon Campus. A 4 mm sieve was used to sift soil and peat for enhanced growth carefully. Furthermore, 1.5 kg of soil and 1.5 kg of peat were homogenized in a cement mixer to generate a potted growth medium. The soil–peat mixture’s moisture content and water-holding capacity were quantified gravimetrically to regulate irrigation in control and treatment pots [21,33]. When properly air-dried, soil–peat moss 1:1 always has moisture. This moisture must be incorporated while determining soil pot capacity water. The initial step involved quantifying the gravimetric water content of the potting soil.

The requisite oven-dried earth was in each pot. Air-dried soil has 41.99% moisture (0.4199 g/g), necessitating 2112.82 g of oven-dried soil. The volume of water required to saturate potting soil to its capacity was measured, allowing for the quantity of air-dried soil necessary to produce 2112.82 g (2.113 kg) of oven-dried soil. The pot’s moisture retention capacity was 66.67%. That quantity was the soil’s 100% pot capacity water needs. Each pot was allocated an equal mass of air-dried soil comparable to oven-dried soil. The moisture content in air-dried soil was evaluated. Notwithstanding its seeming insignificance, that moisture must be taken into account. The soil moisture required to achieve pot capacity was 24.68% (0.2468 g/g). Each gram of soil requires 0.2468 g of liquid for pot capacity. The soil sample that was air-dried contained 0.4199% moisture.

The amount of water needed to accomplish 100% pot capacity (PC) in air-dried soil equals the PC moisture. Air-dried soil moisture content: 66.67% − 41.99% = 24.68% (0.2468 g water/g oven-dried soil). Furthermore, 3 kg of oven-dried soil necessitates 521.44 g (521.44 mL) (0.521 L) to achieve a 100% plasticity coefficient (PC). Water is more readily added by volume than by weight. It should be recalled that the density of water is 1 g/cm³. We know how much water to add to 3 kg of air-dried potting soil to meet pot capacity (100%). Moreover, 50% pot capacity stresses plants, while 100% pot capacity maintains them well-watered. The experiment’s weekly water requirements for plant irrigation were determined using the Time Domain Reflectometer (TDR) calibration curve equation (y = 0.6803x + 0.3041). Substitute the device value into the equation. The result is subtracted from 0.66, pot capacity moisture. Based on pot capacity, the moisture required for potting soil is multiplied by 2112.82, the weight of oven-dried soil. The standard quantity of water required was recorded. The experiment commenced on 11 November 2023 and concluded on 8 March 2024.

2.5. Calculation of Total Sugars That Can Be Hydrolyzed

We extracted and assessed total soluble sugars via a substantially improved technique from the collected dried leaves (100 mg) at the end of the experiment (8 March 2024) at the ripening stage. In a boiling water bath, immersing dry wheat leaves in 5 mL of 2.5 N HCl for 3 h generated soluble sugars with agitation. The cooled boiling tube containing the digested material used solid sodium carbonate to neutralize the solution until the effervescence stopped. The sample was centrifuged at 5000 revolutions per minute. The supernatant was evaporated and mixed with filtered water to assess soluble carbohydrates. The extract was subjected to a boiling water bath for 10 min after cooling to determine total soluble sugars, and 3.0 mL of newly made anthrone was added. Genesys Instruments Ltd. (PG Instruments Ltd., T 80, Lutterworth, UK). T80 spectrophotometer readings showed absorptions at 630 nm [34].

2.6. Calculating Total Phenolic Content (TPC)

The Folin–Ciocalteu reagent method was used to quantify the total phenolic content. The leaves were collected and dried at the end of the experiment at the ripening stage (8 March 2024). Initially, 100 μL of wheat leaves aqueous extract with a concentration of 2.5 g/100 mL were placed into multiple test tubes. Adding 1 mL of double-distilled water brought the volume down to 1 mL. This process was repeated to create several stocks of the plant extract, each with a concentration of 2500 ppm. Moreover, 500 μL of Folin–Ciocalteu reagent with a concentration of 1 normal (1 N) was introduced into the solution, and then 2 mL of sodium carbonate were added with a concentration of 20%. After that, the concoction was sealed in a dark environment at room temperature for 90 min. The determinations were conducted in quadruplicate. The spectrophotometer (Genesys Instruments Ltd. T80) was used to measure the absorbance of the generated blue color at a wavelength of 650 nm relative to the reagent blank. Gallic acid was employed as a reference substance at a concentration ranging from 10 to 100 parts per million (ppm). In micrograms of Gallic acid equivalent (μg of GAE/g of extract), the total phenolic content was measured [21,35].

2.7. Calculating Total Flavonoid Content (TFC)

The aluminum chloride colorimetric technique evaluated the concentrations of flavonoids in aqueous-based wheat extracts. The leaves were collected and dried at the end of the experiment (8 March 2024) at the ripening stage. At first, 500 μL of wheat leaves aqueous extract with a concentration of 2.5 g/100 mL was placed into many test tubes. The volume was then adjusted to 1 mL by adding pure water twice distilled. Subsequently, many plant extract stocks were generated, each with a concentration of 12.5 ppt. The test tubes were thoroughly mixed by vortexing. Subsequently, 300 μL of a 5% sodium nitrite solution was added to every one of the test tubes. The test tubes were vortexed again and left undisturbed for five minutes at room temperature. Each test tube was supplemented with 300 μL of a 10% aluminum chloride solution. The contents were thoroughly mixed with a vortex mixer and left undisturbed for 5 min at room temperature. Furthermore, 2 mL of a 1 molar sodium hydroxide solution were added to each test tube. The contents of the test tubes were mixed thoroughly and then allowed to incubate in the dark at room temperature for 15 min. Ultimately, the spectrophotometer (Genesys Instruments Ltd. T80) was used to measure the absorbance of the pink color that resulted from the reaction against the reagent blank. A wavelength of 510 nm was used for the measurement. Quercetin was employed as a reference substance at a concentration ranging from 100 to 1000 parts per million (ppm). Micrograms of quercetin equivalent (μg of QE/g of extract) were used to measure the total flavonoid content [21,35].

2.8. Determination of the Activities of Glutathione Reductase, Peroxidase, and Catalase Enzymes

Utilizing the slightly altered technique established by Venisse et al. (2001), 0.5 g of fresh leaf tissue was homogenized in a 3 mL 50 mM phosphate buffer at a pH of 7.0. The fresh samples were collected for enzyme determinations on 7 February 2024 at the stem extension stage (Boot). The buffer comprised 1% polyvinylpyrrolidone, 1 mM polyethyleneglycol, 1 mM phenylmethylsulfonyl fluoride, 8% (w/v) polyvinylpyrrolidone, and 0.01% (v/v) Triton X-100. The mixtures were centrifuged at 11,500 rpm for ten minutes at four degrees Celsius, and the resulting supernatant was used to assess enzyme activity, including AsPOX, GR, GST, and POX [36].

2.8.1. Assessment of the Peroxidase (POX) (EC 1.11.1.7) Activity

In 20 s at pH 6.0 and 20 °C, one milligram of purpurogallin can be produced from one unit of peroxidase and pyrogallol. This unit is the same as approximately 18 µM units per minute at 25 °C. Absorbance: 420 nm, formation of perpurogalline. For 2 min, the absorbance was measured at 420 nm every 20 s to track the change [37]. Reaction Mixture: 50 μL leaf extract + 2 mL reaction mix (100 mM phosphate buffer (pH 6), 5% pyrogallol, and 0.5% H₂O₂).

$$\mathrm{units}/\mathrm{mL} \mathrm{enzyme}={\displaystyle \frac{(\mathsf{\Delta }\mathrm{A} 420 \mathrm{n}\mathrm{m}/20 \mathrm{s} \mathrm{T}\mathrm{e}\mathrm{s}\mathrm{t}-\mathsf{\Delta }\mathrm{A} 420 \mathrm{n}\mathrm{m}/20 \mathrm{s} \mathrm{B}\mathrm{l}\mathrm{a}\mathrm{n}\mathrm{k})\times (\mathrm{S}\mathrm{a}\mathrm{m}\mathrm{p}\mathrm{l}\mathrm{e} \mathrm{v}\mathrm{o}\mathrm{l}\mathrm{u}\mathrm{m}\mathrm{e} \mathrm{i}\mathrm{n} \mathrm{m}\mathrm{L})\left(\mathrm{d}\mathrm{f}\right)}{\left(12\right) \times \mathrm{V}\mathrm{o}\mathrm{l}\mathrm{u}\mathrm{m}\mathrm{e} (\mathrm{i}\mathrm{n} \mathrm{m}\mathrm{L}) \mathrm{o}\mathrm{f} \mathrm{e}\mathrm{n}\mathrm{z}\mathrm{y}\mathrm{m}\mathrm{e} \mathrm{u}\mathrm{s}\mathrm{e}\mathrm{d}}}$$

df = Dilution factor.

12 = Extinction coefficient of 1 mg/mL purpurogallin.

2.8.2. Assessment of the Glutathione Reductase (GR) (EC 1.6.4.2) Activity

When NADPH is oxidized, the absorbance reaches 340 nm. Here is the reaction mixture: 50 μL of leaf extract plus 1 mL of the following: 0.2 M Tris/HCl buffer (pH 7.8), 3 mM EDTA, 0.2 mM NADPH, and 0.5 mM oxidized glutathione [36]. The enzyme activity is expressed as units/mL enzyme.

$$\mathrm{units}/\mathrm{mL} \mathrm{enzyme}={\displaystyle \frac{(\mathsf{\Delta }\mathrm{A} 340 \mathrm{n}\mathrm{m}/\mathrm{m}\mathrm{i}\mathrm{n} \mathrm{T}\mathrm{e}\mathrm{s}\mathrm{t}-\mathsf{\Delta }\mathrm{A} 340 \mathrm{n}\mathrm{m}/\mathrm{m}\mathrm{i}\mathrm{n} \mathrm{B}\mathrm{l}\mathrm{a}\mathrm{n}\mathrm{k})\left(\mathrm{d}\mathrm{f}\right)}{\left(6.22\right) \times \mathrm{V}\mathrm{o}\mathrm{l}\mathrm{u}\mathrm{m}\mathrm{e} (\mathrm{i}\mathrm{n} \mathrm{m}\mathrm{L}) \mathrm{o}\mathrm{f} \mathrm{e}\mathrm{n}\mathrm{z}\mathrm{y}\mathrm{m}\mathrm{e} \mathrm{u}\mathrm{s}\mathrm{e}\mathrm{d} \times (\mathrm{S}\mathrm{a}\mathrm{m}\mathrm{p}\mathrm{l}\mathrm{e} \mathrm{v}\mathrm{o}\mathrm{l}\mathrm{u}\mathrm{m}\mathrm{e} \mathrm{i}\mathrm{n} \mathrm{m}\mathrm{L})}}$$

df = Dilution factor.

6.22 = The ß-NADPH millimolar extinction coefficient at 340 nm.

2.8.3. Assessment of Catalase (CAT) Activity (EC 1.11.1.6)

Catalase is an enzyme that breaks down hydrogen peroxide into water and oxygen. The enzyme’s assay depends on estimating residual H₂O₂ by titration with KMnO₄ [38]. A conical flask contains 5 mL distilled water, 1 mL H₂O, 1 mL 0.1 M H₂O₂, and 1 mL enzyme extract. Incubate the mixture at 28 ± 1°C for 15 min. Finally, adding 5 mL of 10% H₂SO₄ to the reaction mixture stops enzyme activity. Titration against KMnO₄ (0.05 N) solution estimates residual H₂O₂ in the mixture. Repeat each set three times. A control set titration is also performed without enzyme in the reaction mixture (6 mL dist. water and 1 mL H₂O₂ substrate). Incubate the set as before and halt the reaction with 5 mL of 10% H₂SO₄ before titration against KMnO₄ (0.05 N) solution. This set is repeated three times. The enzyme’s H₂O₂ decomposition is measured by the difference between the control set (without enzyme) and reaction mixture (with enzyme) titration readings. Titration readings are taken when a faint pink tint lasts at least 10 s. mg H₂O₂ decomposed/gm. of fresh tissue/1 mL of 0.5 N KMnO₄ = 17 mg of H₂O₂.

2.9. Statistical Analysis

This study used experimental methods to examine all subjects four times and report the average value and standard error. The WASP program was used for statistical analysis. This study examined group differences using one method to analyze variance (ANOVA). As a statistical tool, the LSD test was used with 1% and 5% significance [39,40].

3. Results

3.1. An Investigation of Green ZnO-NPs

X-ray diffraction (XRD) peak lists of zinc oxide nanoparticles that were manufactured in an environmentally responsible manner are displayed in Figure 1. These peak lists include residual sodium (Na) and oxygen (O). The prominent peaks for ZnO nanoparticles could be seen in red when the diffraction of the particles was studied. Figure 2, Figure 3, Figure 4 and Figure 5 demonstrate TEM, SEM, and energy-dispersive X-ray spectroscopy of ZnO-NPs generated using the green method. Figure 2’s TEM image shows ZnO-NPs’ excellent quality due to their nearly hexagonal shape. Images obtained using SEM of ZnO nanoparticles are depicted in Figure 3 and Figure 4, respectively. These images illustrate the particles’ uniformity in terms of both their shape and their size. On closer inspection, it is clear that ZnO nanoparticles are present in powder form. The ZnO sample primarily consists of zinc (Zn) and oxygen (O), as seen in Figure 5, with both elements uniformly distributed across the ZnO-NPs’ surface. There were traces of carbon elements and sulfurous compounds present. The size distribution of the ZnO-NPs manufactured using the green approach is depicted in Figure 6. These nanoparticles display a monomodal distribution with a half-width of approximately 51 nanometers. An investigation into the zeta potential was conducted to determine the surface charges of zinc oxide nanoparticles. The results of this study illuminated the stability of the colloidal ZnO nanoparticles. There is a correlation between the magnitude of zeta potential and the potential stability of colloids. In that particular investigation, the nanoparticles that were produced displayed a remarkable level of stability, as evidenced by a zeta potential value of −19.12 millivolts. Water was used as a dispersion, and the zeta potential of the zinc oxide nanoparticles that were produced in an environmentally responsible manner was evaluated to determine their potential. It is illustrated in Figure 7 that the zeta potential that was obtained was −19.12 millivolts. The FTIR spectra of ZnO-NPs that were manufactured in an environmentally responsible manner are depicted in Figure 8. These spectra were collected within the wavelength range of 4000 to 500 cm⁻¹. Some peaks were connected with six different functional groups in the spectrum of zinc oxide nanoparticles. These peaks were observed at 3500, 3100, 1690.50, 1450.14, 546.40, and 490.20 cm⁻¹.

3.2. Biochemical and Stress Marker Analysis of Wheat Leaves Subjected to Various Treatments

Table 1. The effects of seed priming with ZnO-NPs and DR GREEN fertilizer on the total phenolic content (TPC) in wheat plants under normal irrigation and drought stress conditions.

Treatments	TPC (mg/g)
T1 (Control, normal irrigation 100% PC)	19.24 ± 0.04 a
T2 (Control, drought stress 50% PC)	17.64 ± 0.20 b
T3 (Normal irrigation 100% PC+ seeds were primed with ZnO-NPs)	17.76 ± 0.23 b
T4 (Drought stress 50% PC + seeds were primed with ZnO-NPs)	16.21 ± 0.20 d
T5 (Normal irrigation 100% PC+ seeds were primed with DR GREEN fertilizer)	16.97 ± 0.13 c
T6 (Drought stress 50% PC+ seeds were primed with DR GREEN fertilizer)	17.36 ± 0.21 bc
LSD (0.01)	0.784
LSD (0.05)	0.559
Coefficient of variation	1.793

Each value represents the mean ± SE. Values with the same letter are not significantly different at (p ≤ 0.05), and the comparison is performed according to different treatments in the main column.

shows that the total phenolic content (TPC) decreased under drought stress in the plants exposed to drought stress (T2) and the plants primed with ZnO-NPs under drought stress (T4) compared to the control plants and plants primed with ZnO-NPs (T1 and T3) that were non-stressed or non-stressed. However, the seeds were previously primed with ZnO-NPs, while DR GREEN–seed-primed plants showed a significant increase in the plants primed with DR GREEN under drought stress (T6) compared to the plants primed with DR GREEN (T5), which were not stressed. The highest TPC was obtained from the control plants in regular 100% PC irrigation (19.24 mg/g), and the lowest content was noticed in drought-stressed plants in 50% PC irrigation while the seeds were primed with ZnO-NPs (16.21 mg/g).

Table 2. The effects of seed priming with ZnO-NPs and DR GREEN fertilizer on the total flavonoid content (TFC) in wheat plants under normal irrigation and drought stress conditions.

Treatments	TFC (mg/g)
T1 (Control, normal irrigation 100% PC)	40.26 ± 1.29 ns
T2 (Control, drought stress 50% PC)	35.72 ± 3.45 ns
T3 (Normal irrigation 100% PC+ seeds were primed with ZnO-NPs)	34.95 ± 4.07 ns
T4 (Drought stress 50% PC + seeds were primed with ZnO-NPs)	28.29 ± 1.90 ns
T5 (Normal irrigation 100% PC+ seeds were primed with DR GREEN fertilizer)	30.95 ± 2.55 ns
T6 (Drought stress 50% PC+ seeds were primed with DR GREEN fertilizer)	29.92 ± 0.56 ns
LSD (0.01)	ND
LSD (0.05)	ND
Coefficient of variation	13.503

Each value represents the mean ± SE. ^ns refers to non-significant, and ND refers to non-determined. The comparison is performed according to different treatments in the main column.

shows the total flavonoid content (TFC), and there were no significant differences among all the treatments, whether they were stressed or not.

Table 3. The effects of seed priming with ZnO-NPs and DR GREEN fertilizer on the total hydrolysable sugar content in wheat plants under normal irrigation and drought stress conditions.

Treatments	Total Sugars (mg/g)
T1 (Control, normal irrigation 100% PC)	191.32 ± 5.09 b
T2 (Control, drought stress 50% PC)	143.06 ± 6.66 c
T3 (Normal irrigation 100% PC+ seeds were primed with ZnO-NPs)	140.83 ± 1.04 c
T4 (Drought stress 50% PC + seeds were primed with ZnO-NPs)	152.43 ± 6.54 c
T5 (Normal irrigation 100% PC+ seeds were primed with DR GREEN fertilizer)	156.32 ± 3.33 c
T6 (Drought stress 50% PC+ seeds were primed with DR GREEN fertilizer)	211.67 ± 7.62 a
LSD (0.01)	23.868
LSD (0.05)	17.024
Coefficient of variation	5.767

Each value represents the mean ± SE. Values with the same letter are not significantly different at (p ≤ 0.05), and the comparison is performed according to different treatments in the main column.

showed that the total hydrolyzable sugar content increased under drought stress in the plants primed with DR GREEN under drought stress (T6) (211.67 mg/g) that was stressed, and the seeds were previously primed with the fertilizer DR GREEN. The plants primed with DR GREEN under drought stress (T6) were followed by the control plants (T1), representing the control plants that were not stressed or primed (191.32 mg/g). There were no significant differences among the other treatments in the total sugar content, whether it was stressed, stressed/primed, or not.

3.3. Determination of the Activities of Glutathione Reductase, Peroxidase, and Catalase Enzymes

Figure 9, Figure 10 and Figure 11 show the released enzymes: glutathione reductase (GR), peroxidase (POX), and catalase (CAT). There was a trend for all released enzymes that the second stressed and non-primed treatment (T2) had the highest concentration of the produced antioxidant enzymes. The seeds not primed with ZnO-NPs or DR GREEN fertilizer and irrigated in 50% PC irrigation had a higher concentration of produced GR, POX, and CAT enzymes than those irrigated in regular irrigation (100% PC). It was also noticed that the control plants (T1), whose seeds were not stressed or primed, had the lowest concentration of the released enzymes compared to the other treatments. In the case of the GR enzyme, there were no significant differences between the plants primed with ZnO-NPs and the plants primed with ZnO-NPs under drought stress (T3 and T4) in the production level. The same was true in the case of the plants primed with DR GREEN and the plants primed with DR GREEN under drought stress (T5 and T6). In the case of the POX enzyme, it was noticed that the plants primed with ZnO-NPs under drought stress (T4) had a higher concentration of the production level compared to the plants primed with ZnO-NPs (T3), and the same was true in comparing the plants primed with DR GREEN (T5) with the plants primed with DR GREEN under drought stress (T6), as the releasing level of POX in T6 (the plants primed with DR GREEN under drought stress) was a little bit higher than in T5 (the plants primed with DR GREEN). However, it is clear that even though the concentration of the enzyme in the plants primed with ZnO-NPs under drought stress and the plants primed with DR GREEN under drought stress was higher than in the plants primed with ZnO-NPs and the plants primed with DR GREEN, it was still less than the concentration of the production level in the case of the plants exposed to drought stress (T2), proving the positive priming effect of ZnO-NPs and DR GREEN fertilizer in decreasing the impact of the drought stress. Regarding the CAT enzyme, it showed the same trend, as although its concentration in the plants primed with ZnO-NPs under drought stress and the plants primed with DR GREEN under drought stress was higher than in the plants primed with ZnO-NPs and the plants primed with DR GREEN, it was still less than the concentration of the production level in the case of the plants exposed to drought stress (T2) that had the highest level of the enzyme concentration.

4. Discussion

The environmentally synthesized ZnO nanoparticles, utilizing the aqueous extract of corn husk as agricultural waste, were examined via transmission electron microscopy (TEM) to ascertain their crystalline properties and dimensions precisely. TEM images of ZnO-NPs confirm the hexagonal morphology of the particles and exhibit limited variation in thickness, aligning with the results of SEM. TEM images revealed that most ZnO nanoparticles (NPs) possess hexagonal morphology, with dimensions nearing 100 nanometers. The surface morphologies generated by ZnO nanoparticles were investigated using scanning electron microscopy (SEM). The scanning electron microscopy (SEM) image of the ZnO nanoparticles demonstrates the existence of spherical formations with closely aggregated particles. The prior results of TEM and STEM investigations aligned with the conclusions presented by Santhoshkumar et al. (2017) [41]. The findings validated that the dimensions of the ZnO-NPs measured 51 nm.

The EDAX analysis confirmed the presence of greenly synthesized ZnO nanoparticles. The elemental analysis of the ZnO-NPs indicated the presence of zinc and oxygen, thereby validating the purity of the synthesized ZnO-NPs. Applying a lacy carbon-coated grid enhances the quality of STEM electron micrographs, which is why the EDAX analysis incorporated carbon. The presence of a sulfur (S) element is contributed to by the structure of the corn husk itself, as the concentration of S in the corn husk reaches 38% [42]. The crystalline structure of ZnO nanoparticles was assessed by X-ray diffraction analysis. The diffraction pattern displayed nine distinct peaks at specific 2θ values. The XRD examination of the environmentally produced ZnO nanoparticles displayed distinctive diffraction patterns. The peaks at 2θ values of 32.6°, 34.56°, 37.36°, 47.5°, 57.45°, 63.5°, 67°, 68.22°, 73.4°, and 77.4° corresponded to the crystallographic planes of ZnO nanoparticles, along with the average particle size of the zinc nanoparticles. No additional diffraction peaks from other phases were detected, which demonstrated the great phase purity of the ZnO-NPs powder, as depicted in Figure 1. The diffraction patterns obtained in this investigation closely paralleled those documented by Ahmed et al. (2024) and Elbahnasawy et al. (2023) [21,35,43].

DLS analysis assesses the hydrodynamic radius [44]. The distribution of ZnO nanoparticles in their suspension and the average size of them were ascertained using DLS analysis. ZnO-NPs were found to have an average of 51 nm, as shown in Figure 6, and those results matched the findings by Kalaba et al. (2024) [44]. The stability of nanostructures is intimately correlated with the zeta potential (surface potential) [45]. Zeta potential analysis was conducted to assess the surface charge of ZnO-NPs, revealing a mean zeta potential of −19.2 mV, which signifies reasonable stability. A comparable zeta potential value (−19.6 and −20.9 mV) is achieved for ZnO-NPs produced from the leaf extract of sea cucumber (Holothuria impatiens) and Cinnamomum tamala [43,46]. Electrostatic repulsions among singly charged nanoparticles during dispersion result in strong positive or negative zeta potential, signifying enhanced physical stability. Conversely, because of the appealing Van der Waals pressures exerted on them, particles with low zeta potential are prone to coagulation or flocculation, likely resulting in inadequate physical stability.

Fourier transform infrared spectroscopy (FTIR) analysis revealed an absorption band at 546.40 cm⁻¹, which serves as a definitive indicator of ZnO bonding (fingerprint), establishing that the synthesized materials were indeed ZnO nanoparticles. The bending vibration band of the C-H bond was seen with a frequency of 1450.14 cm⁻¹. The pronounced peak at 1690.50 cm⁻¹ corresponds to the carbonyl (C=O) stretching vibration of the amide I in proteins [43,47]. The peak at 3500 cm⁻¹ corresponds to the stretching vibration of the O–H bond in water, alcohol, and phenols [21,43,48].

While the impact of ZnO-NPs on plant physiology and biochemistry has been established, widespread discussions and uncertainties remain concerning their effects on plant secondary metabolism [49,50]. The impact of nanoparticles on phenolic compounds is significant, as these chemicals are crucial for plant productivity and adaptation in response to biotic and abiotic challenges [49]. Consequently, it is essential to comprehend the comprehensive operation of the plant’s secondary metabolism in response to applying nanoparticles as a potential inducer of oxidative stress.

ZnO nanoparticles have been proven to have both beneficial and phytotoxic effects on the germination and growth of seedlings, as evidenced by prior studies that established their impact on these processes. These impacts have been demonstrated to be both positive and negative. The nature, size, and concentration of nanoparticles, in addition to the genotype of the species that is being assessed, are the primary factors that are responsible for the variability of the response. Possibly responsible for this inhibition are two primary consequences, which are as follows: (1) A chemical toxicity that is dependent on the release of damaging ions, one example of which is the infiltration of nanoscale particles and the dissolution of Zn²⁺ ions from ZnO; (2) A stress or stimulation that is caused by the surface characteristics, size, or shape of the particles [50]. The tolerance law proposed by Shelford is in agreement with the concept of toxicity at increasing doses. The decrease in root length and plant biomass may be the result of toxicity that was caused by the plant’s uptake of nanoparticles from the environment and subsequent accumulation of these particles either intracellularly or intercellularly [51]. Root length and plant biomass may have decreased as a result of this toxicity. Specifically, the connection between reactive oxygen species (ROS) and secondary signaling messengers, which are responsible for the transcriptional regulation of secondary plant metabolism, is of utmost importance, especially when taking into consideration the reactions of plants to nanoparticle treatment [52]. As a result of the truth that micronutrients like manganese, which is a cofactor of phenylalanine ammonia-lyase, are accountable for the transformation of phenolic compounds, nutri-priming has an effect on the secondary metabolism of plants [53].

The content of phenolic compounds can vary greatly [54], and they also play a role as electron donors in the detoxification mechanisms present in organelle structures [55]. Phenolic compounds play a crucial part in the process of ROS detoxification. A study indicates that elevated zinc concentrations can increase phenol content in all Coriandrum sativum L. plant regions, particularly affecting the radicles the most. Besides supplying antioxidants to mitigate excessive ROS production, radicals serve as physical and chemical defenses against biotic and abiotic stresses [56]. Radicals also provide antioxidants. These substances may interact synergistically, elucidating the observed patterns in phenolic compounds and antioxidant activity [57]. Comparable patterns have been identified for Stevia rebaudiana Bertoni [58], the antioxidant activity of Salvia officinalis L. [59], and the radicle of Brassica nigra [60].

Since the TPC decreased under drought stress in the plants exposed to drought stress (T2) and the plants primed with ZnO-NPs under drought stress (T4) compared to the control plants (T1) and the plants primed with ZnO-NPs (T3) that were non-stressed or non-stressed; however, the seeds were previously primed with ZnO-NPs, while DR GREEN–seed-primed plants showed a significant increase under the drought stress in the plants primed with DR GREEN under drought stress (T6) compared to the plants primed with DR GREEN (T5) that were not stressed. Moreover, there were no significant differences among all the treatments, whether they were stressed or not, according to TFC. It means that environmental stress may increase/decrease the synthesis of phenolic chemicals, which detoxify harmful free radicals. Stewart et al. (2000) found that cultivar significantly altered tomato phenolics despite uniform environmental conditions [61]. Water also significantly affects plant phenolic metabolism and fruit composition [62]. The effect of environmental stress on some plants, like tomatoes’ phenolic compounds, is unclear, with studies showing a rise [21,63], a decrease [64], or no change [65]. Some researchers have found that tomato fruits grown in saline have more flavonoids [66], while others have seen a decrease [66].

It was shown in Table 3 that the total hydrolyzable sugar content increased under drought stress in the plants primed with DR GREEN under drought stress (T6) (211.67 mg/g), which occurred after the seeds had been primed with the fertilizer DR GREEN. As was previously observed for other wheat cultivars [67], the increase in hydrolyzable sugars may be connected with the plant’s stress responses throughout its life cycle. Both of these responses have been documented. In addition, it is anticipated that the mature grain will have a significant amount of glucose for glycolysis, which is the first step of respiration that is necessary for germination [68]. During the process of seed priming, the seedling dry weight, leaf area, and leaf CO₂ net assimilation are all improved. Additionally, the photochemical efficiency of photosystem II is optimized, and the activity of α-amylase is increased under abiotic stress [69]. Therefore, the higher amylase activity that occurs during the maturity of primed seeds may result in a stronger oligosaccharide breakdown, which would explain the decrease in the sugars that were described earlier.

Plant ROS metabolism imbalance causes oxidative damage. Long considered the primary line of defense against ROS production, antioxidant enzymes [70,71]. Antioxidant defense enzymes POX, CAT, and GR protect. Plants use these mechanisms to remove H₂O₂, O²⁻, and other ROS. To evaluate how priming treatments affect wheat plant oxidative status, POX, CAT, and GR were examined.

Figure 9, Figure 10 and Figure 11 illustrate the activity of antioxidant enzymes in wheat plants exposed to various priming treatments following 40 days of cultivation. Peroxidase (POX) facilitates the elimination of reactive oxygen species that cause cellular oxidative damage [70,72]. Plant peroxidases serve as biochemical indicators for various abiotic stressors. The current investigation revealed a considerable reduction in POX activity in both ZnO-NPs and DR GREEN-primed plants compared to the plants exposed to drought stress (T2). The activity levels of POX in Zn-NP-primed plants markedly differed from those in DR GREEN-primed plants. ZnO-NPs have shown greater efficacy in decreasing the production level of POX compared to DR GREEN’s fertilizer.

Similarly, other antioxidant enzymes also played a role in scavenging ROS in plants. CAT protects against oxidative damage to the cell. According to [71], CAT has been suggested to scavenge the bulk H₂O₂, while peroxidases can sequester the remaining H₂O₂. The SOD can convert O₂⁻ to H₂O₂ while CAT decomposes H₂O₂ to H₂O and oxygen molecules [73]. In this study, CAT enzyme activity was significantly increased in the second treatment that was stressed without any priming for its seeds, as well as DR GREEN fertilizer primed, but those seeds that were primed with the fertilizer showed a little increase in the CAT level compared to the second treatment, which meant that although the fertilizer-primed plants had a higher CAT level compared to the control plants (T1), the produced enzyme was still less than the level in the second treatment. Weisany et al. (2012) reported that the increase in the activity of CAT is probably due to the indirect requirement of zinc for the high activity of enzymes involved in H₂O₂ detoxification, such as CAT [74].

Jahantab et al. (2022) proposed that GR activity is regulated and diminished by the application of low concentrations of metal oxide nanoparticles (MO-NPs) and varying levels of drought stress in both Agropyron cristatum L. and Sanguisorba officinalis L. [75]. Nevertheless, elevated concentrations of AgO, FeO, ZnO, and CdO-NPs, specifically, exert detrimental effects and enhance GR activity. GR regulates physiological systems, particularly during drought stress, and will escalate under extreme stress [76]. Consequently, a reduction is observed at low concentrations of MO-NPs. This finding contrasts with prior research that demonstrated that ZnO-NPs (100 and 200 mM) reduced GR activity by 25% and 45%, respectively, in wheat seedlings [77].

The observed reduction in ROS production in ZnO-NP-primed plants can be ascribed to the presence of Zn, which significantly contributes to the attenuation of ROS creation and the protection of cells from ROS damage. Our findings align with the earlier research conducted by Latef et al. (2016), which highlighted the function of nano-Zn in preserving redox homeostasis in plants pre-treated with nano-Zn [78,79].

Therefore, the application of zinc in lower concentrations in nano form likely limited the increase in ROS levels, thereby reducing the activity of antioxidant enzymes such as POX, CAT, and GR. According to Cakmak (2008) and Broadley et al. (2007) [80,81], zinc deficiency can lead to increased ROS production, resulting in plant damage. The observed reduction in ROS production in ZnO NP plants can be ascribed to the presence of Zn, which significantly contributes to the attenuation of ROS creation and the protection of cells from ROS damage. Our findings align with the earlier research conducted by Latef et al. (2016), which indicated the function of nano-Zn in preserving redox homeostasis in plants pre-treated with nano-Zn [78].

Micronutrient seed priming (nutri-priming) has been employed to address micronutrient deficiencies in crops [82,83,84,85,86,87]. Nutri-priming involves soaking seeds in solutions enriched with micronutrients before germination and planting, such as DR GREEN fertilizer. This cost-effective approach can enhance the micronutrient content in the edible parts of plants and raise yield and/or protein levels in several crops, including wheat [83,88,89,90,91,92]. Nonetheless, these advantages are contingent upon the application of appropriate micronutrient dosages, as excessive use may induce cyto- and/or phytotoxicity [93,94,95], resulting from osmotic stress and redox imbalance that alter biochemical compounds including total soluble sugars, amino acids, proteins, phenolics, hormones, and lipids [96].

The ascorbate–glutathione (AsA-GSH) pathway plays a crucial role in detoxifying H₂O₂ and regulating ROS [70,97]. Consequently, activating the AsA-GSH pathway through priming with some chemical solutions like salicylic and ascorbic acids is strongly associated with abiotic stress tolerance. In addition to ROS detoxification, glutathione (GSH) is linked to methylglyoxal (MG) detoxification as drought increases its formation, which aggravates oxidative damage to the next folds [70]. In the current experiment, CAT and POX enzymes—principal enzymes of the AsA-GSH pathway—were significantly decreased in the case of priming with the solution containing the critical micronutrients (DR GREEN), as those enzymes act as primary ROS detoxifiers within the chloroplast, preserving its redox equilibrium [97]. This meant that DR GREEN’s nutrients had contributed to ameliorating the harmful effects of drought stress.

5. Conclusions

The influence of nanoparticles on seed germination, including water absorption, root emergence, and the activation of food mobilization enzymes, necessitates additional research. Nutrient particles improve nutrient absorption in drought-stressed plants; nonetheless, their effects on nutrient channels and ionic transporters necessitate further research. Nutri-priming, like DR GREEN fertilizer, alleviates deficits in crop nutrition. Nutri-priming involves submerging seeds in nutrient-dense liquids. The findings of this study indicated that seed priming with ZnO-NPs and DR GREEN fertilizer exerted distinct influences on the regulation of certain antioxidative enzymes and biochemical indicators. This study investigated the properties of the environmentally synthesized ZnO nanoparticles, which exhibited well-defined shapes, charge, and crystallinity. Their spectra also exhibited some active functional groups that may be attributed to specific features.

The TPCs diminished in plants experiencing drought stress (T2) and in those treated with ZnO-NPs under drought conditions (T4) relative to control plants (T1) and non-stressed plants treated with ZnO-NPs (T3). In contrast, DR GREEN–seed-primed plants showed a notable enhancement. Moreover, no substantial differences were detected across all treatments, irrespective of stress circumstances, as evidenced by TFC. Drought-stressed plants treated with DR GREEN (T6) exhibited elevated levels of total hydrolyzable sugars. This study investigated the POX, CAT, and GR antioxidant enzymes in wheat plants after various priming treatments.

Decreased zinc concentrations in nano form likely attenuated the elevation of reactive oxygen species (ROSs), thereby reducing the activities of peroxidase (POX), catalase (CAT), and glutathione reductase (GR). The concentrations of CAT and POX enzymes markedly diminished upon priming with the critical micronutrients from the DR GREEN solution, suggesting that DR GREEN’s nutrients mitigated drought stress. Green nanoparticles and seed priming promote sustainable agriculture. Their use promotes regenerative agriculture and reduces our environmental impact. This study offers researchers a promising avenue to utilize agricultural waste, such as corn husks, in the synthesis of beneficial metallic oxide nanoparticles, including ZnO-NPs, as well as fertilizers that can alleviate the detrimental effects of drought on commercially valuable crops like wheat. The results may serve as indicators for open-field applications to mitigate one of the most severe abiotic stressors adversely impacting plants: drought stress.