Optimizing Nanopriming Duration: Chitosan‐Stabilized Iron Oxide Enhances Lettuce Seedling Performance and Cost‐Effectiveness

ABSTRACT

Climate change and nutrient limitations increasingly hinder seed germination and early seedling establishment, highlighting the need for sustainable enhancement strategies. This study evaluates a green nanopriming approach using chitosan‐stabilized iron oxide nanoparticles synthesized via an aqueous, low‐energy co‐precipitation process that employs biodegradable chitosan and avoids hazardous reagents, fully aligning with green chemistry principles. Lettuce seeds were primed in a 1 ppm nanoparticle suspension for 2, 6, 8, 16, and 24 h to identify the optimal priming duration. Chitosan markedly improved nanoparticle dispersion and colloidal stability, producing a strongly positive surface charge (+35.5 mV) that enhanced nanoparticle–seed interactions. The 8‐h priming treatment yielded the greatest improvements, achieving a germination rate of 96.67% alongside the highest shoot and root elongation and biomass accumulation. Longer exposures (16 and 24 h) reduced performance, suggesting potential overstimulation or early metabolic imbalance. These enhancements likely arise from improved iron bioavailability, nanoparticle stability, and stimulation of early metabolic activation. A pilot‐scale cost assessment demonstrated excellent economic feasibility, with production costs of only $4.40 per 10‐L batch, supporting scalability for agricultural use. Overall, optimizing nanopriming duration using chitosan‐stabilized iron oxide nanoparticles offers a sustainable, economical, and environmentally responsible approach to strengthen crop establishment under stress.

Chitosan‐stabilized Fe₃O₄ nanoparticles synthesized via an aqueous green route exhibit high colloidal stability and are employed for seed nano‐priming. Systematic variation of priming duration (224 h) identifies an optimal exposure time (8 h) that maximizes seed germination and early seedling growth. The graphical abstract illustrates the relationship between nanoparticle stability, priming duration, and biological response, highlighting the critical role of time‐dependent nanoseed interactions in enhancing plant performance while avoiding overexposure effects.

1. Introduction

Global agriculture is at a critical juncture, facing multiple challenges that threaten food security, crop productivity, and economic stability. Climate change manifested through rising temperatures, drought, and unpredictable weather patterns has significantly impacted global crop yields. For example, the International Grains Council recently revised its global wheat production forecast for 2024–2025 downward by 2 million metric tons, predominantly due to adverse climatic events in Europe [1]. Such climatic stresses directly diminish seed vigor, a critical determinant of successful crop establishment. Seed vigor encompasses rapid germination, robust seedling growth, and stress tolerance attributes frequently compromised by elevated temperatures during seed development, thereby reducing overall crop performance [2, 3].

In response, seed priming technologies have emerged as pre‐sowing strategies that enhance seed vigor and germination under challenging environmental conditions. Traditional priming techniques such as hydropriming and osmopriming use controlled hydration to activate seed metabolism and improve germination rates [4, 5, 6, 7]. Extending this concept, nanopriming integrates engineered nanoparticles (NPs) to further enhance seed physiological responses, including antioxidant enzyme activities, hormonal signaling, and nutrient uptake efficiency [8, 9]. Recent studies have demonstrated that nanopriming with iron oxide NPs (FeNPs) significantly improves biomass and yields in crops such as wheat by enhancing photosynthetic efficiency and nutrient bioavailability [10, 11].

Iron, a vital micronutrient involved in chlorophyll biosynthesis and enzymatic processes, often presents limited bioavailability in agricultural soils, necessitating effective delivery approaches [12]. While FeNPs offer an attractive solution owing to their high surface reactivity and large surface area, their practical application is frequently impeded by physicochemical instability, particularly particle agglomeration. This aggregation, primarily driven by van der Waals and magnetic dipole–dipole interactions, significantly reduces NP dispersion and bioavailability [13]. To mitigate this, surface stabilization strategies such as coating with biodegradable polymers are critical. Chitosan, a naturally occurring polysaccharide, has emerged as an excellent stabilizer due to its ability to enhance NP colloidal stability, reduce aggregation, and promote bioactivity, making it highly suitable for agricultural applications [14, 15].

Despite nanopriming's promise, conventional agriculture remains heavily reliant on chemical fertilizers, contributing to environmental pollution and diminishing returns in productivity due to inefficient nutrient use and uneven seed nutrient absorption [16, 17]. In addition, practical implementation of new technologies depends significantly on their economic viability. Thus, it is crucial to optimize not only the biological performance of nanopriming but also its economic feasibility for large‐scale agricultural use. Surprisingly, the effects of priming duration critical for balancing effective metabolic activation with economic practicality remain poorly understood, creating a significant knowledge gap.

Building upon our preliminary investigation with uncoated FeNPs [10], the present study introduces a systematic approach to optimize lettuce (Lactuca sativa) seed nanopriming using chitosan‐stabilized iron oxide NPs (CS‐FeNPs). Unlike previous work, this study specifically aims to: (i) improve FeNP colloidal stability through precise chitosan stabilization, (ii) identify an optimal priming duration that maximizes seed germination and seedling vigor, and (iii) assess the economic feasibility of CS‐FeNP nanopriming at pilot scale to evaluate practical scalability for agricultural adoption. It is hypothesized that careful optimization of priming duration and NP stabilization will synergistically enhance bioavailability, biological activity, and economic sustainability. Overall, this study provides significant interdisciplinary insights combining bioinorganic chemistry, nanobiotechnology, and sustainable agriculture by offering novel, economically viable nanopriming approaches. This contributes toward addressing critical agricultural challenges posed by climate stress and nutrient limitations, ultimately supporting global food security and sustainable agricultural practices.

2. Materials and Methods

2.1. Materials Used for NP Synthesis and Seed Preparation

This study utilized lettuce (L. sativa L.) seeds, selected for their rapid germination, widespread use in agricultural research, and sensitivity to priming techniques [10]. The seeds were sourced from a certified supplier (Crop Power Sdn. Bhd., Malaysia) to ensure uniformity and viability. Chitosan with a molecular weight of 1000 g/mol, iron(III) chloride hexahydrate (FeCl₃·6H₂O, ≥ 98%), and iron(II) chloride tetrahydrate (FeCl₂·4H₂O, 98%) were used as the primary materials for NP synthesis. Chitosan was obtained from Biolution Resources Sdn. Bhd., Malaysia, while all other chemical reagents were purchased from Sigma‐Aldrich, MA, USA. Analytical‐grade ethanol and deionized water, produced using a Barnstead Smart2Pure Water Purification System (Thermo Fisher Scientific, MA, USA), were used throughout the synthesis and washing steps to ensure consistency and high purity. A Vibra‐Cell ultrasonic probe (Sonics & Materials Inc., CT, USA) was employed to achieve uniform dispersion of the NPs, and a 3 M ammonia solution was used as the precipitating agent.

2.2. Synthesis of CS‐FeNPs

CS‐FeNPs were synthesized using a co‐precipitation method adapted from established protocols [18]. Specifically, 0.1 g of chitosan (1 kDa) was dissolved in 100 mL of 1% (w/v) acetic acid, then heated to 70°C until fully solubilized. In parallel, 2.7 g of FeCl₃·6H₂O and 1.0 g of FeCl₂·4H₂O were dissolved in 100 mL deionized water, maintaining a 2:1 molar ratio to yield magnetite (Fe₃O₄) [19]. The iron solution was then purged with nitrogen gas at a flow rate of 100 mL/min for 30 min in a 250 mL three‐necked round‐bottom flask to minimize Fe²⁺ oxidation. Following this, the iron solution was slowly added to the chitosan solution under vigorous magnetic stirring (600 rpm). A 3 M ammonia solution was added dropwise (approximately 10 mL total) until the pH reached ∼11, triggering NP precipitation. The mixture was kept at 70°C for 1 h to ensure nucleation and stabilization. The NPs were then washed repeatedly with deionized water and ethanol, followed by centrifugation at 10 000 rpm for 20 min until neutral pH was achieved. The resulting CS‐FeNPs were oven‐dried at 60°C to constant weight and stored for further use.

2.3. Characterization of NPs

To analyze the morphological and colloidal properties of the synthesized NPs, transmission electron microscopy (TEM) and dynamic light scattering (DLS) techniques were employed. For TEM analysis, 1 mg of dried NP samples—both FeNP and CS‐FeNP—was dispersed in 10 mL of analytical‐grade acetone to form a suspension [20]. The suspension was ultrasonicated for 5 min at a frequency of 50 kHz to ensure uniform dispersion and minimize aggregation. A droplet of the suspension was then placed on a carbon‐coated copper grid and air‐dried under clean, dust‐free conditions. TEM imaging was carried out using a JEM‐1400Plus Electron Microscope (JEOL Ltd., Japan) operating at 120 kV, providing high‐resolution images for assessing particle morphology and size distribution.

For FESEM and EDS analysis, NP powders of FeNP and CS‐FeNP (1k) were sputter‐coated with platinum and analyzed using a JSM‐7610F (JEOL Ltd.). FESEM was used to observe the surface morphology of NPs, while EDS mapping was conducted to evaluate the elemental distribution and confirm the presence of Fe, C, O, and N. This analysis was particularly useful for validating the successful coating of FeNPs with chitosan, with nitrogen (N) serving as a distinct signature of the polymer.

To evaluate the suspension stability of FeNP and CS‐FeNP over time, a sedimentation kinetics study was performed by diluting NP stock solutions to 10 ppm in deionized water and transferring the suspensions into standard cuvettes. The absorbance at 532 nm was recorded at regular time intervals using a UV–Vis spectrophotometer (Shimadzu UV‐1800, Japan). The decline in absorbance was used to estimate the relative suspension stability and sedimentation rate of NPs, allowing for quantitative comparison between bare and chitosan‐coated formulations.

For DLS analysis, NP suspensions of FeNP and CS‐FeNP were prepared at a concentration of 1 ppm (equivalent to 0.2 mg in 20 mL deionized water) to evaluate hydrodynamic diameter and zeta potential [21]. The suspensions were ultrasonicated for 2 min before measurement using a Malvern Zetasizer Nano‐ZS ZEN 3600 (Malvern Panalytical, UK). These analyses enabled the assessment of chitosan's stabilizing effect on FeNPs, confirming the importance of uniform dispersion and colloidal stability for effective nanopriming applications.

2.4. Seed Priming Procedure and Germination Assay

Seeds of L. sativa L. were purchased from Crop Power Sdn. Bhd. (Malaysia) through the online platform in March 2024. Species identity was confirmed visually based on supplier information and morphological characteristics by the research team. The material examined corresponds to commercially distributed seed stock; therefore, no herbarium voucher was deposited. All experimental procedures used this same batch of seeds to ensure consistency throughout the study. To determine the optimal priming duration for lettuce seeds, seeds were treated with a 1 ppm suspension of CS‐FeNPs across varying durations (2, 6, 8, 16, and 24 h) [22]. This concentration was selected based on prior optimization trials to balance effective NP bioavailability without inducing phytotoxicity [23]. No range comparison was done in this study, but 1 ppm was found to yield consistent germination improvement. A total of 180 seeds were used, with three biological replicates of 30 seeds per treatment. Each replicate represented an independent batch of seeds subjected to the same priming and germination procedure, ensuring biological independence and reproducibility. Before priming, all seeds were surface‐sterilized using 3% hydrogen peroxide for 10 min and rinsed three times with deionized water to remove residual oxidants.

Sterilized seeds were immersed in the CS‐FeNP suspension contained in sterile 50 mL Falcon tubes, with gentle orbital shaking at 100 rpm to ensure uniform exposure and prevent NP settling. After each priming duration, seeds were removed and air‐dried on sterile filter paper at room temperature for 24 h, allowing them to return to their initial moisture content before sowing. This step minimized the risk of premature germination or radicle protrusion during storage.

After drying, the seeds were sown in 90 mm Petri dishes lined with Whatman No. 1 filter paper moistened with 5 mL of deionized water. The germination assay was conducted in a controlled growth chamber at 25°C under a 12‐h light/dark photoperiod for 14 days [24]. Parameters evaluated included germination percentage, root length, shoot length, and fresh weight of seedlings. Control treatments included (i) seeds primed with deionized water and (ii) seeds primed with FeNPs without chitosan coating, to distinguish the effects of chitosan stabilization.

2.5. Statistical Analysis

All experimental data were subjected to statistical analysis to determine the significance of observed differences among treatments. Seed germination and growth parameters were analyzed using one‐way analysis of variance (ANOVA), followed by Fisher's least significant difference (LSD) post hoc test. Each treatment was conducted in triplicate, and the results were expressed as mean ± standard deviation (SD). Statistical analyses were performed using IBM SPSS software, with a significance level set at p < 0.05.

3. Results and Discussion

3.1. Physicochemical Characterization of NPs

In Figure 1A, uncoated FeNPs aggregate into dense, irregular clusters. This clustering arises from two fundamental forces at the nanoscale: (i) magnetic dipole–dipole interactions, wherein each superparamagnetic Fe₃O₄ core transiently magnetizes and attracts its neighbors, and (ii) van der Waals forces, which become increasingly significant as interparticle distances shrink (< 10 nm) [25]. The result is a highly polydisperse ensemble in the dry state with limited interstitial separation.

FIGURE 1.

TEM images of nanoparticles. (A) FeNP showing clustered aggregation due to magnetic interactions and absence of stabilizing agents. (B) CS‐FeNP exhibit visible chitosan corona contributing to increased core size, more defined particles with improved dispersion due to steric stabilization by the chitosan layer.

By contrast, Figure 1B reveals that CS‐FeNPs exist as more discrete, uniformly sized cores enveloped by a thin chitosan layer. The measured core diameter (∼102.8 nm) includes both the iron oxide nucleus and its polysaccharide sheath. Fundamentally, chitosan's rigid polymer chains extend into the surrounding medium, creating a steric barrier that physically prevents cores from approaching each other closer than the polymer thickness. Concurrently, protonation of chitosan's primary amines at the working pH imparts a positive surface charge, generating an electrostatic double layer that further repels adjacent particles. This dual (“electrosteric”) stabilization mechanism dramatically reduces the effective aggregation energy barrier.

When these stabilized particles are dispersed in water, the electrostatic repulsion (zeta potential +35.5 mV; Table 1) and steric hindrance work in concert to maintain a much narrower size distribution (∼112.7 nm hydrodynamic diameter) compared to bare FeNPs (∼234.2 nm) [26]. The TEM images thus visually confirm that chitosan not only coats but also functionally transforms FeNPs into a stable colloid; an essential prerequisite for reproducible, uniform seed nanopriming.

TABLE 1.

Hydrodynamic size and zeta potential of FeNP and CS‐FeNP as measured by DLS. CS‐FeNP shows a smaller hydrodynamic size and significantly higher zeta potential compared to uncoated FeNP, indicating improved colloidal stability.

Nanoparticles treatment	Hydrodynamic size (nm)	SD (nm)	Zeta potential (mV)	SD (mV)
FeNP	234.2	98.76	13.4	3.46
CS‐FeNP	112.7	22.52	35.5	4.82

In Figure 2A, bare FeNPs appear as angular, polyhedral crystals that cluster into dense agglomerates. This morphology reflects the inherently high surface energy of freshly precipitated magnetite, which without any surface modifier drives particles to minimize total surface area via sintering‐like contacts and fusion at grain boundaries. The irregular shapes and sharp edges further promote mechanical interlocking once dried, making these aggregates particularly resistant to redispersion.

FIGURE 2.

FESEM images of synthesized nanoparticles at 30 000× magnification. (A) Bare FeNPs exhibit dense agglomeration with irregular particle shapes. (B) CS‐FeNPs display a more dispersed morphology with smoother surfaces and improved particle definition, indicating the stabilizing effect of chitosan coating.

By contrast, Figure 2B shows CS‐FeNPs as well‐separated particles with rounded profiles and visibly smoother surfaces. The chitosan layer, adsorbed during synthesis, acts as a physical “cushion” around each core, reducing surface energy and blocking direct Fe₃O₄–Fe₃O₄ contact. Fundamentally, the polymer chains impose an entropic penalty on particle–particle approach: compressing the adsorbed chitosan layers reduces their conformational freedom, creating a repulsive steric force. Simultaneously, protonated amine groups generate electrostatic repulsion. These combined forces prevent grain boundary formation and yield a stable, discrete morphology even in the dry state an essential feature predicting rapid and uniform redispersion in aqueous priming environments [26, 27].

Energy‐dispersive x‐ray spectroscopy (EDS) was employed to validate the surface chemistry modifications imparted by chitosan stabilization (Figure 3). In the uncoated FeNP sample, iron (Fe) and oxygen (O) dominate the elemental composition at approximately 36 and 42 at%, respectively, with carbon (C) contributing the remaining ≈22 at% due to adventitious surface contamination. No nitrogen (N) was detected, confirming the absence of organic stabilizers. Upon chitosan stabilization, a distinct nitrogen signal (∼5 at%) emerges, directly attributable to the primary amine (─NH₂) functionalities of the chitosan backbone. Concurrently, the carbon content increases to ≈25 at%, reflecting the polysaccharide matrix enveloping the iron oxide core. The slight reduction in the detected Fe and O signals (to 35.7 and 37.2 at%, respectively) is best explained by the attenuation of core‐derived x‐ray counts as they traverse the chitosan overlayer. These changes confirm that chitosan forms a conformal, uniform shell around each Fe₃O₄ NP, providing both electrostatic and steric stabilization [26, 27]. Moreover, the presence of surface‐exposed amine groups is likely to enhance bioactive interactions with negatively charged seed coat biomolecules, thereby promoting NP adhesion and facilitating controlled iron delivery during seed nanopriming.

FIGURE 3.

EDS elemental composition (Fe, C, O, and N) for FeNP and CS‐FeNP.

Following the detailed morphological assessment, we probed the colloidal stability of both bare FeNPs and chitosan‐stabilized FeNPs (CS‐FeNPs) via sedimentation kinetics (Figure 4). Suspension capacity, defined as the ratio of instantaneous to initial absorbance (I/I ₀), decays rapidly for uncoated FeNPs: over 50% of the suspension settles within 30 min, and only ∼25% of particles remain dispersed after 600 min (Figure 4A). This precipitous decline reflects the dominance of attractive forces, namely, van der Waals interactions and magnetic dipole–dipole attractions which, in the absence of a stabilizing shell, drive particle aggregation and gravitational sedimentation. By contrast, CS‐FeNPs maintain I/I ₀ > 0.8 even after 60 min and retain ∼60% of their initial suspension after 600 min, evidencing markedly enhanced colloidal stability.

FIGURE 4.

(A) Sedimentation kinetics profiles of FeNP and CS‐FeNP suspensions measured over 600 min, indicating higher colloidal stability for CS‐FeNPs as reflected by slower sedimentation rates. (B) Total suspension stability quantified by area under the curve (AUC) analysis, where CS‐FeNPs exhibit significantly greater AUC values, affirming enhanced suspension retention compared to bare FeNPs. Error bars represent standard deviation (n = 3).

To quantify this improvement, we calculated the area under the sedimentation curve (AUC) for each formulation (Figure 4B). The AUC for CS‐FeNPs is nearly double that of bare FeNPs, highlighting a substantial extension of suspension lifetime and therefore longer effective contact time with seed surfaces during priming. Fundamentally, this enhanced stability arises from chitosan's electrosteric stabilization mechanism: protonated ─NH₃ ⁺ groups on the polysaccharide backbone impart a strong positive surface charge, increasing the electrostatic repulsion component of the DLVO potential barrier, while the polymer chains physically hinder close approach of particles. The resulting combination of increased repulsive energy and steric hindrance elevates the activation energy required for aggregation, thereby retarding both cluster formation and sedimentation under gravity. Such prolonged suspension is critical for uniform NP distribution around seeds, ensuring consistent adsorption, diffusion, and internalization key prerequisites for effective nanopriming.

A derivative analysis of the sedimentation kinetics (Figure 5) further elucidates the dynamic stability of bare FeNPs versus CS‐FeNPs by quantifying the instantaneous rate of absorbance decline (ΔI/Δt) over time. For unmodified FeNPs, the initial sedimentation rate peaks sharply exceeding 0.02 I/I ₀ per minute within the first 10 min reflecting rapid cluster formation as particles overcome repulsive barriers and settle under gravity. The pronounced fluctuations in ΔI/Δt up to 100 min indicate episodic aggregation events, wherein loosely bound clusters periodically detach and re‐settle, consistent with weak interparticle repulsion and dominant van der Waals and magnetic attractions.

FIGURE 5.

Derivative analysis of sedimentation kinetics profiles for FeNP and CS‐FeNP (1k), showing sedimentation rate (ΔI/Δt) over time.

In stark contrast, CS‐FeNPs exhibit a markedly attenuated and smooth sedimentation profile. The initial ΔI/Δt remains below 0.005 and decays gradually to near zero by 300 min, indicating that most particles remain in stable suspension long after the FeNPs have fully settled. This behavior arises from the electrosteric stabilization conferred by the chitosan shell: protonated amine groups generate a robust positive surface potential that elevates the energy barrier for aggregation, while the polymer brush imparts steric constraints that hinder core–core approach. Moreover, the low molecular weight (∼1 kDa) of chitosan used here affords a thin, uniform coating that maximizes surface coverage without excessively increasing hydrodynamic drag, thereby preserving dispersibility. Fundamentally, this dual stabilization mechanism transforms the sedimentation process from a kinetically driven, erratic collapse into a diffusion‐controlled, quasi‐stable suspension. By maintaining NPs in prolonged contact with the seed surface throughout the priming period, CS‐FeNPs ensure consistent bioavailable iron delivery; a prerequisite for uniform metabolic activation and the enhanced germination performance observed. These findings align with prior studies demonstrating that chitosan‐stabilized magnetic NPs exhibit superior hydrodynamic stability compared to their uncoated counterparts, underscoring the critical role of surface engineering in designing effective nanobiotechnologies for agriculture [14, 25, 26].

DLS analysis (Table 1) further elucidated how chitosan stabilization transforms both the hydrodynamic behavior and electrokinetic profile of FeNPs, offering a mechanistic explanation for their enhanced colloidal stability. Bare FeNPs exhibit a broad hydrodynamic diameter of 234.2 ± 98.8 nm substantially larger than the 9.6 nm core measured by TEM indicating extensive aggregation in suspension. Their low surface potential (ζ = +13.4 ± 3.5 mV) lies well below the ±30 mV threshold generally required to generate sufficient electrostatic repulsion in aqueous media [26], allowing van der Waals and magnetic dipolar attractions to dominate and drive cluster formation.

In contrast, CS‐FeNPs display a markedly reduced hydrodynamic size of 112.7 ± 22.5 nm, reflecting two key phenomena: (i) the polymeric chitosan corona mitigates secondary aggregation by physically spacing NPs at length scales exceeding the Debye screening length, and (ii) the presence of a solvation shell around the chitosan chains increases effective particle–solvent interactions, enhancing Brownian motion and opposing gravitational settling. Crucially, the CS‐FeNPs' zeta potential soars to +35.5 ± 4.8 mV above the colloidal stability threshold owing to the protonation of chitosan's ─NH₂ groups in mildly acidic conditions [15, 27]. This high surface charge creates a thick electrostatic double layer; as two particles approach, overlapping diffuse layers generate a strong repulsive force that must be overcome before van der Waals attractions can act, thereby elevating the aggregation energy barrier according to DLVO theory. From a fundamental standpoint, the interplay between steric hindrance and enhanced electrostatic repulsion often termed electrosteric stabilization yields a colloidal regime in which particle–particle collisions are both kinetically and energetically disfavored. The thinness (ca. 5–10 nm) yet uniformity of the chitosan coating ensures that hydrodynamic drag is not unduly increased, preserving rapid diffusion and effective surface coverage. Consequently, CS‐FeNPs maintain a narrow size distribution in solution, ensuring reproducible and uniform interactions with seed surfaces during nanopriming. These combined enhancements in hydrodynamic size reduction and zeta potential elevation provide a solid physicochemical foundation for the observed improvements in seed germination and early seedling growth under CS‐FeNP treatment [28, 29].

3.2. Germination and Seedling Growth Performance Under Different Priming Durations

The germination performance of L. sativa seeds was significantly influenced by the duration of priming with CS‐FeNP. A total of 30 seeds were used for each experimental group, including the control and priming durations of 2, 6, 8, 16, and 24 h. Seeds were dried to original moisture levels post‐priming to eliminate radical protrusion bias. Therefore, the observed improvements are attributed to the treatment effect of CS‐FeNP, rather than prolonged water uptake or overhydration. The use of a shaker during priming further ensured NP‐seed surface interaction.

While prior studies have reported germination effects of FeNPs on lettuce [10], the current work expands the scope by integrating biopolymer stabilization with optimized priming durations. The incorporation of chitosan has been shown to not only enhance dispersion but also introduce beneficial surface charge properties that improve interaction with seeds. Moreover, the optimized 8‐h window revealed a priming “sweet spot,” balancing activation without overexposure. This refinement highlights the importance of tuning both NP surface chemistry and exposure kinetics which a contribution not addressed in earlier works.

Statistical analysis (one‐way ANOVA followed by Fisher's LSD test at p < 0.05) confirmed that priming duration significantly influenced germination outcomes. The 8‐h CS‐FeNP treatment yielded the highest germination percentage (96.67 ± 3.33%), which was significantly greater (p < 0.05) than the control (46.67 ± 5.77%), FeNP (61.67 ± 4.41%), and other duration treatments. Full statistical results, including error bars and group comparisons, are shown in Figure 2. Different letters above bars indicate statistically significant differences based on LSD analysis.

The unprimed control group exhibited a germination count of 14 seeds, corresponding to a germination percentage of 46.67%. In contrast, the primed groups showed improved germination percentages of 63.33% (2 h), 80.00% (6 h), 96.67% (8 h), 60.00% (16 h), and 53.33% (24 h). Notably, the 8‐h priming duration yielded the highest germination percentage, with 29 of 30 seeds germinating. This result suggests that an 8‐h priming duration optimizes the benefits of CS‐FeNP, while prolonged durations (16 and 24 h) result in diminished effectiveness. According to Debbarma et al., insufficient priming time may fail to initiate key enzymatic and metabolic repair processes necessary for germination, while excessive priming can lead to premature radicle emergence, increasing susceptibility to desiccation damage during drying [30]. In this study, identifying the optimal priming window ensures metabolic activation without compromising seed viability [6, 31]. Figure 6 illustrates the germination percentages across different priming durations, highlighting the superior performance at 8 h and providing a visual comparison between primed and unprimed seeds. These findings underscore the importance of optimizing priming duration for maximum germination efficiency and demonstrate the potential of CS‐FeNP to enhance seed performance under controlled conditions. In addition, priming duration also influenced subsequent seedling development, including root and shoot lengths and their respective fresh weights.

FIGURE 6.

Effect of priming duration on germination percentage of Lactuca sativa seeds. Seeds were primed for 2, 6, 8, 16, and 24 h and compared to an unprimed control. The highest germination percentage (96.67%) was observed at the 8‐h priming duration, indicating the optimal exposure time for enhancing germination performance.

Root and shoot lengths were also significantly improved at 8 h (3.02 ± 0.18 and 5.18 ± 0.24 cm, respectively), both showing clear statistical separation (p < 0.05) from other treatments. Particularly, the 24‐h priming group exhibited significantly lower values, suggesting potential overexposure effects. Figure 7A,B represents the improvement of plant performance primed with CS‐FeNP in its respective hours. Fresh weight measurements further supported these trends. The highest fresh root (0.0101 g) and shoot weights (0.0281 g) were also recorded at 8 h, indicating superior biomass accumulation. Comparatively, root fresh weights for 2, 6, 16, and 24 h were 0.0035, 0.0062, 0.0042, and 0.0031 g, respectively, while shoot weights were 0.0114, 0.0194, 0.0123, and 0.0081 g. Figure 7C presents the visual appearance of seedlings after 14 days, clearly showing enhanced root and shoot development at the 8‐h priming duration. These results are consistent with the physicochemical characteristics of CS‐FeNP, as shown by the TEM and DLS analyses. The small hydrodynamic size (112.7 nm) and high zeta potential (35.5 mV) of CS‐FeNP suggest effective NP dispersion and uptake during seed priming. The chitosan coating likely enhanced the interaction of NPs with seed surfaces, facilitating improved absorption during the optimal priming period [32]. In conclusion, the findings emphasize the importance of fine‐tuning priming duration when using CS‐FeNP to achieve optimal seedling vigor. These outcomes also highlight the role of physicochemical properties in determining NP bioavailability and uptake efficiency. Further investigation is warranted to explore the underlying mechanisms, including NP internalization pathways and cellular interactions.

FIGURE 7.

Germination performance and early seedling vigor of Lactuca sativa under CS‐FeNP nanopriming at varying durations. (A) Shoot and root lengths, (B) shoot and root fresh weights, and (C) visual representation of seedlings corresponding to each priming duration (2, 6, 8, 16, and 24 h). All seedlings were aligned horizontally with root–shoot junctions placed along the same baseline to enable direct comparison of shoot and root lengths. The most pronounced growth enhancement was observed at 8 h priming duration, indicating optimal exposure. This figure is original and created entirely by the authors.

This study emphasizes germination percentage, shoot/root length, and fresh weight as practical and widely accepted proxies for seed vigor and early growth performance. Although more complex germination indices such as mean germination time or vigor index were not assessed, our selected parameters provide meaningful and interpretable evidence of treatment efficacy in the context of comparing priming durations. The 8‐h CS‐FeNP treatment produced the highest germination percentage (96.67%) and significantly improved seedling growth metrics, reinforcing its potential as an effective nanopriming strategy. This approach aligns with recent works highlighting that germination percentage and early seedling morphology remain key markers for evaluating nanopriming outcomes in field‐relevant species [33, 34, 35].

Compared to conventional priming methods such as hydropriming or osmopriming (e.g., PEG priming), nanopriming offers enhanced benefits through both physical and biochemical mechanisms. While traditional priming facilitates controlled water uptake to activate metabolic processes, it often lacks precise control over micronutrient delivery or oxidative signaling modulation [36]. In contrast, CS‐FeNP‐based nanopriming not only improves colloidal stability and nutrient bioavailability (Fe), but also introduces reactive oxygen species (ROS)‐mediated signaling pathways that promote faster germination and robust seedling vigor [33]. Recent studies have reported that FeNP nanopriming significantly outperforms PEG and hydropriming in promoting early growth, chlorophyll content, and antioxidant enzyme activity in various crops including rice and tomato [37, 38, 39]. Thus, the nanopriming strategy in this study reflects a mechanistically enhanced and potentially better sustainable approach.

Furthermore, although many nanopriming studies emphasize cereal or horticultural crops, emerging evidence supports significant benefits in leafy vegetables as well. For instance, a recent study on lettuce demonstrated that employing multiple NPs for seed nanorpiming led to increases in biomass, antioxidant enzyme activity, and photosynthetic efficiency, while also mitigating cadmium toxicity stress [40]. This reinforces the potential applicability of nanopriming in leafy species, not just major grains. Moreover, the review by Nile et al. highlights that nanopriming can upregulate aquaporin expression and enhance α‐amylase activity which a mechanisms also observed in nanoprimed leafy seed [33, 37]. By situating our lettuce‐specific findings within this growing body of work, our results contribute to bridging a crucial gap: demonstrating how CS‐FeNPs may realize these mechanistic benefits in lettuce, a globally consumed leafy vegetable.

3.3. Proposed Mechanisms Underlying Enhanced Germination and Growth by Nanopriming

The pronounced enhancement of germination and seedling vigor following CS‐FeNP nanopriming can be understood as a sequence of finely orchestrated physicochemical and biological events (Figure 8). First, chitosan's electrosteric stabilization ensures that FeNPs remain uniformly suspended, maximizing NP–seed encounters. Upon contact, the protonated ─NH₃ ⁺ moieties of chitosan engage in strong electrostatic interactions with the anionic pectins and proteins of the seed coat, effectively “anchoring” NPs at the surface and facilitating their penetration through microscopic apertures in the testa and cuticle [41]. Once adsorbed, the chitosan shell serves as a controlled‐release matrix: under the mildly acidic priming environment, chitosan's amine groups gradually complex with iron ions and undergo slow protonation‐deprotonation cycles that liberate Fe²⁺/Fe³⁺ at rates tuned to seed uptake kinetics [42]. These released iron species diffuse into the aleurone and scutellum, where they act as indispensable cofactors for germination‐related enzymes, including catalase and peroxidase, which decompose inhibitory hydrogen peroxide, and superoxide dismutase, which removes superoxide radicals [43]. This enzymatic activation promotes mobilization of endosperm reserves (starch and proteins), guaranteeing a steady flow of assimilates to the emerging radicle.

FIGURE 8.

Proposed schematic of the nanopriming mechanism using chitosan‐stabilized iron oxide nanoparticles (CS‐FeNPs). Upon soaking, NPs diffuse into the seed coat, enhancing water uptake, Fe availability, and activating α‐amylase and ROS‐scavenging enzymes (SOD, CAT, and POD). These biochemical responses accelerate germination and seedling growth through improved antioxidant defense and metabolic readiness. All elements in this figure were created by the authors.

Simultaneously, trace amounts of ROS generated via Fenton reactions iron‐catalyzed conversion of H₂O₂ to hydroxyl radicals act as key signaling molecules. These ROS, maintained at sublethal levels by the aforementioned antioxidant enzymes, trigger the expression of aquaporin genes and expansin proteins, enhancing water permeability and loosening cell walls to accommodate rapid cell elongation [44, 45]. The net result is accelerated imbibition, faster cell turgor generation, and more vigorous root and shoot protrusion.

Beyond iron delivery, chitosan itself plays a bioactive role: oligomeric fragments released from the polymer can bind to pattern recognition receptors in seed tissues, inducing mild defense‐like responses that upregulate endogenous antioxidant systems and adjust phytohormone balances specifically by attenuating abscisic acid (ABA) and elevating gibberellin (GA) signaling thus favoring germination over dormancy [46].

Crucially, the 8‐h priming window represents the temporal convergence of these processes: it is long enough to allow sufficient NP adhesion, iron release, and signaling activation, yet short enough to avoid radicle desiccation or oxidative damage that might result from prolonged exposure. This harmonious interplay between chitosan‐engineered NP delivery, controlled iron and ROS signaling, enzymatic mobilization of reserves, and hormone‐mediated growth regulation exemplifies a multifaceted mechanism by which CS‐FeNP nanopriming confers superior seed performance that offers a sustainable, tunable strategy for crop establishment under stress [47].

The observed optimum at 8 h of CS‐FeNP priming likely reflects alignment with key physiological stages in early seed germination. Germination is classically divided into three phases: Phase I (rapid water uptake or imbibition), Phase II (plateau phase involving metabolic activation, enzyme repair, and reserve mobilization), and Phase III (radicle protrusion and cell elongation). The 8‐h window may correspond with the transition from Phase I to II, during which seeds become metabolically active but have not yet reached radicle emergence. This timing could allow CS‐FeNPs to efficiently deliver iron while modulating ROS signaling pathways essential for enzymatic activation, without inducing premature growth or cellular stress.

Although we hypothesize that CS‐FeNPs promote seed germination via enhanced ROS signaling, activation of antioxidant enzymes, and improved iron bioavailability, this study did not directly measure biochemical markers such as malondialdehyde (MDA), antioxidant enzyme activities (SOD and CAT), or endogenous ROS levels.

Previous studies support these mechanisms in NP‐based priming systems. For instance, Mazhar et al. showed that seed priming with FeNPs elevated SOD and CAT by ∼13% and 38%, respectively, in wheat [48]. Yang et al. demonstrated nanopriming‐induced upregulation of aquaporin genes and α‐amylase activity in seeds, suggesting enhanced metabolic activation pathways [37]. In addition, Shelar et al. described the concept of an “oxidative window” wherein controlled ROS levels support germination, aligning conceptually with our proposed mechanism [49]. These evidence collectively suggest that prolonged exposure (∼24 h) may lead to oxidative imbalance or cellular damage. Therefore, the 8 h priming duration appears to strike a balance between enabling NP internalization and metabolic activation while limiting cytotoxic stress. In future work, we will validate this timing hypothesis through time‐course biochemical assays and gene expression profiling.

Besides, while the present study included both untreated control seeds and FeNPs without chitosan coating, it did not include a “chitosan‐only” treatment group. This omission limits the ability to fully disentangle whether the observed effects result solely from FeNPs, from chitosan alone, or from a synergistic interaction between the two components. However, the central aim of this study was to investigate the role of chitosan as a stabilizing agent that improves the colloidal behavior of FeNPs and reduces their tendency to agglomerate, thus enhancing nanorpiming efficiency. The chitosan used here was not primarily assessed as an independent biostimulant but rather as a physico‐chemical modifier for FeNP dispersion.

Future studies should include a chitosan‐only control group to clarify its biological contribution and enable a more comprehensive understanding of the independent and combined effects of chitosan and FeNPs. Besides, to further substantiate the mechanistic pathways proposed in this study, future work will incorporate direct biochemical measurements to validate the role of ROS regulation and micronutrient activation during nanopriming. This will include quantifying lipid peroxidation through MDA levels and assessing antioxidant enzyme activities such as superoxide dismutase, catalase, and peroxidase. These markers will help determine whether optimal priming duration aligns with controlled oxidative signaling, while prolonged exposure may induce oxidative stress. Additional analyses, including iron uptake quantification and in situ visualization of ROS using fluorescence‐based probes, will also provide deeper insight into NP–seed interactions. Integrating these biochemical assessments will enable a more definitive evaluation of the underlying mechanisms governing the enhanced seedling performance observed with chitosan‐stabilized iron oxide nanopriming. Despite this limitation, our comparison between FeNP and CS‐FeNP treatments helps to isolate the influence of chitosan on NP performance in seed priming applications.

3.4. Cost and Scalability of CS‐FeNP Nanopriming for Agricultural Application

To assess the real‐world applicability of CS‐FeNP priming, we developed a pilot‐scale process flow diagram and conducted a thorough techno‐economic evaluation (Figure 9; Table 2). The workflow is divided into two core stages: NP synthesis and seed priming. In the synthesis stage, aqueous solutions of iron precursors (FeCl₃·6H₂O and FeSO₄·7H₂O) and chitosan dissolved in acetic acid are separately prepared in stirred reactors. Under an inert nitrogen atmosphere and vigorous stirring, the iron salt solution is introduced to the chitosan solution at 70°C, and the pH is adjusted to ∼11 via ammonia addition to precipitate magnetite cores enveloped by a uniform chitosan shell. This low‐temperature co‐precipitation route avoids energy‐intensive calcination and uses entirely water‐based chemistries, thereby minimizing both capital and operating costs as well as environmental impact [19]. The nascent CS‐FeNPs are immediately subjected to low‐gradient magnetic separation (LGMS) to remove macroscopic aggregates, then oven‐dried at 60°C to produce a free‐flowing powder.

FIGURE 9.

Simulated pilot scale process flow diagram of CS‐FeNP nanopriming.

TABLE 2.

Estimated cost for pilot‐scale CS‐FeNP synthesis and seed nanopriming process (10 L batch).

Description a	Cost ($)/h b
LGMS (wet drum type, $2500 over 15 years)	0.07
Ultrasonic bath (240 W, $200 over 10 years)	0.002
Reaction tank (20 L, FRP, $600 over 10 years)	0.06
Laboratory oven/drying system ($500 over 10 years)	0.05
Power consumption (ultrasound, oven, mixing, 1.5 kW total) c	0.12
Installation (minor plumbing, tubing, drying rack) d	0.30
Labor e	1.00
Raw materials (chitosan, Fe salts, reagents)	2.80
Total ($/h)	4.40

^a Refer supporting documents for technical data, preference for each equipment and raw materials cost showed in Tables S1 and S2.

^b Estimated cost based on batch process (8 h per day, 6 working days per week, 50 working weeks per year).

^c Estimation cost based on Malaysia commercial tariff rate by Tenaga Nasional Berhad (TNB) [52].

^d Installation cost estimated at ∼50% of total equipment purchase cost, based on capital cost estimation methods from Towler and Sinnott and Turton et al. This includes piping, fittings, instrumentation, and small hardware integration required for pilot‐scale batch operation [43, 44].

^e Estimation cost based on 2023–2024 Malaysia's technician labor rate/wage [45].

The dried CS‐FeNP powder is reconstituted in deionized water and held in a buffer tank before seed treatment. Immediately before priming, the suspension undergoes ultrasonication to reestablish the electrosterically stabilized dispersion (hydrodynamic diameter ∼112.7 nm; ζ‐potential +35.5 mV), ensuring consistent NP distribution. Seeds are immersed in this colloid for the optimized 8 h duration, identified in earlier germination assays as the “sweet spot” for maximizing germination percentage (96.7 ± 3.3%) and early seedling vigor without causing premature radicle protrusion or oxidative damage. Following priming, the NP suspension can be recycled up to 80% recovery through a closed‐loop system that returns used suspension to the holding tank for re‐dispersion and reuse, aligning with circular‐economy principles and dramatically reducing material losses.

A comprehensive cost breakdown for a 10‐L batch (Table 2) reveals that raw materials and labor constitute the majority of expenses. Specifically, food‐grade chitosan (sourced from crustacean waste) and ferric salts account for roughly $3.00 per batch, while technician labor contributes an estimated $1.00 per cycle. Utilities include electricity for mixing (0.5 kW), ultrasonication (0.24 kW), and low‐temperature oven drying (0.5 kW)—add another $0.32 per hour. Capital amortization of core equipment (co‐precipitation reactor, LGMS, ultrasonicator, and oven) and routine maintenance contributes under $0.15 per hour when spread over a ten‐year lifespan at standard utilization rates. Altogether, the total operating cost is approximately $4.40 per per hour.

By contrast, conventional hydropriming methods and hormonal priming (e.g., gibberellic acid) at similar scales typically incur costs in the range of $10–15 and $12–18 per hour, respectively, once factors such as cold‐chain logistics, reagent stability, and waste disposal are incorporated [50]. Advanced nano‐delivery platforms such as carbon nanotubes or metal–organic frameworks often exceed $20–30 per hour due to multi‐step synthesis procedures, high‐purity precursor requirements, and extensive post‐treatment purification [51]. Furthermore, at a typical seeding density of 200 g/ha for lettuce, the per‐hectare material cost for a 1 ppm CS‐FeNP treatment equates to approximately $0.02 per hectare, rendering the nanopriming input cost negligible relative to standard fertilizer or pesticide applications.

From a process‐engineering standpoint, the magnetic recoverability of CS‐FeNPs substantially lowers downstream separation costs and minimizes particulate waste. The batch configuration affords simple scale‐up potential and seamless integration into existing seed treatment facilities or decentralized agricultural cooperatives. Because no hazardous organic solvents or high‐temperature furnaces are required, the technology imposes minimal regulatory burden and aligns with green chemistry and sustainable agriculture directives. Finally, a conservative payback analysis assuming a modest 5% yield improvement attributable to enhanced germination and early vigor indicates that the incremental revenue generated would offset the nanopriming operational costs within a single growing season.

In summary, the pilot‐scale process for CS‐FeNP nanopriming combines low‐energy, water‐based synthesis with effective reuse strategies to deliver a robust, eco‐friendly, and economically viable method for seed enhancement. By bridging colloidal engineering with agronomic performance and cost‐benefit analysis, this study demonstrates that CS‐FeNP nanopriming offers a scalable, low‐risk technology capable of improving early‐stage crop establishment, optimizing resource use, and supporting sustainable agricultural production on a practical timeline.

While the present study focused primarily on the germination and early seedling growth stages under controlled in vitro conditions, it is important to acknowledge that plant performance in open‐field environments may differ due to multiple interacting variables such as soil microbiota, nutrient heterogeneity, environmental stressors, and competition. As such, this study serves as a foundational exploration of a preliminary investigation into the efficacy of CS‐FeNP nanopriming. Future studies are warranted to validate these findings under greenhouse and field conditions, including assessments of seedling survival, biomass accumulation, yield performance, and long‐term environmental safety. Such translational steps are essential for bridging laboratory success with real‐world agronomic application and commercialization viability.

4. Conclusions

In this work, we have shown that CS‐FeNPs constitute a highly effective seed‐priming agent for lettuce (L. sativa), combining robust physicochemical performance with clear agronomic benefits and economic viability. Detailed TEM, FESEM, and DLS analyses confirmed that the chitosan shell endows FeNPs with “electrosteric” stabilization manifested as a uniform ∼102.8 nm dry‐state diameter, a reduced hydrodynamic size (∼112.7 vs. 234.2 nm for bare FeNPs), and a high positive zeta potential (+35.5 mV). These features collectively prevent aggregation, ensure prolonged suspension, and maximize NP‐seed surface interactions during priming.

Germination assays across five exposure durations (2, 6, 8, 16, and 24 h) revealed an optimal 8 h window that delivered a 96.7% germination rate double that of unprimed controls and produced the longest roots, shoots, and highest fresh biomass. This “sweet‐spot” balances sufficient NP uptake and iron release with avoidance of overexposure‐induced stress or premature radicle emergence. While the precise molecular underpinnings such as controlled ROS signaling, aquaporin activation, and reserve‐mobilizing enzyme induction remain to be fully elucidated, our results strongly implicate enhanced iron bioavailability and chitosan‐mediated bio‐stimulation as key drivers.

Crucially, a 10 L pilot‐scale process flow and simplified techno‐economic analysis demonstrated that CS‐FeNP nanopriming can be implemented at approximately $4.40 per hour substantially below conventional hydropriming or hormone‐based methods while leveraging magnetic recoverability for reagent reuse. This low‐cost, water‐based, moderate‐temperature approach aligns with green chemistry and circular‐economy principles, making it eminently scalable for decentralized seed‐treatment facilities. Taken together, our findings position CS‐FeNP nanopriming as a versatile, sustainable technology for accelerating seed germination and seedling vigor.

Although this study focuses on early‐stage germination and seedling development under controlled conditions, we acknowledge that the performance of nanoprimed seeds under field conditions may vary due to environmental factors. Hence, these findings should be viewed as preliminary, warranting further validation under greenhouse and field settings. Therefore, future studies should dissect the biochemical and genetic responses to CS‐FeNPs and validate long‐term yield benefits under field conditions, thereby advancing the integration of nanobiotechnology into mainstream agricultural practice.

Author Contributions

Nurfarwizah Adzuan Hafiz: writing – original draft, investigation, methodology. Anca Awal Sembada: writing – original draft, visualization, data curation. Mohamed Syazwan Osman: writing – review and editing, supervision, funding acquisition, conceptualization. Noor Fitrah Abu Bakar: writing – review and editing, supervision, conceptualization. Mohamad Sufian So'aib: writing – review and editing, supervision, conceptualization. I. Wuled Lenggoro: writing – review and editing, supervision.

Funding

Funding provided by the Ministry of Higher Education, Malaysia, through the Fundamental Research Grant Scheme (Grant No. FRGS/1/2022/STG05/UITM/02/10).

Ethics Statement

The authors have nothing to report.

Consent

Informed consent was not required as this study did not involve human participants.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Supporting File 1: cbdv70819‐sup‐0001‐SuppMat.docx

Data Availability Statement

Data will be made available on request.