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Zinc Oxide Nanoparticles: An Influential Element in Alleviating Salt Stress in Quinoa (Chenopodium quinoa L. Cv Atlas)

Aras Türkoğlu
Kamil Haliloğlu
Melek Ekinci
Metin Turan
Ertan Yildirim
Halil İbrahim Öztürk
Atom Atanasio Ladu Stansluos
Hayrunnisa Nadaroğlu
Magdalena Piekutowska
9 and
Gniewko Niedbała
Department of Field Crops, Faculty of Agriculture, Necmettin Erbakan University, Konya 42310, Türkiye
Department of Field Crops, Faculty of Agriculture, Ataturk University, Erzurum 25240, Türkiye
Department of Horticulture, Faculty of Agriculture, Atatürk University, Erzurum 25240, Türkiye
Department of Agricultural Trade and Management, Faculty of Economy and Administrative Sciences, Yeditepe University, Istanbul 34755, Türkiye
Health Services Vocational School, Binali Yıldırım University, Erzincan 24100, Türkiye
Department of Field Crops, Faculty of Agriculture, Upper Nile University, Malakal 71100, South Sudan
Department of Food Technology, Vocational College of Technical Sciences, Ataturk University, Erzurum 25240, Türkiye
Department of Nanoscience and Nano-Engineering, Institute of Science, Ataturk University, Erzurum 25240, Türkiye
Department of Botany and Nature Protection, Institute of Biology, Pomeranian University in Słupsk, 22b Arciszewskiego St., 76-200 Słupsk, Poland
Department of Biosystems Engineering, Faculty of Environmental and Mechanical Engineering, Poznań University of Life Sciences, Wojska Polskiego 50, 60-627 Poznań, Poland
Authors to whom correspondence should be addressed.
Agronomy 2024, 14(7), 1462;
Submission received: 30 May 2024 / Revised: 3 July 2024 / Accepted: 4 July 2024 / Published: 5 July 2024
(This article belongs to the Section Farming Sustainability)


Climate change has intensified abiotic stresses, notably salinity, detrimentally affecting crop yield. To counter these effects, nanomaterials have emerged as a promising tool to mitigate the adverse impacts on plant growth and development. Specifically, zinc oxide nanoparticles (ZnO-NPs) have demonstrated efficacy in facilitating a gradual release of zinc, thus enhancing its bioavailability to plants. With the goal of ensuring sustainable plant production, our aim was to examine how green-synthesized ZnO-NPs influence the seedling growth of quinoa (Chenopodium quinoa L. Cv Atlas) under conditions of salinity stress. To induce salt stress, solutions with three different NaCl concentrations (0, 100, and 200 mM) were prepared. Additionally, Zn and ZnO-NPs were administered at four different concentrations (0, 50, 100, and 200 ppm). In this study, plant height (cm), plant weight (g), plant diameter (mm), chlorophyll content (SPAD), K/Na value, Ca/Na value, antioxidant enzyme activities (SOD: EU g−1 leaf; CAT: EU g−1 leaf; POD: EU g−1 leaf), H2O2 (mmol kg−1), MDA (nmol g−1 DW), proline (µg g−1 FW), and sucrose (g L−1), content parameters were measured. XRD analysis confirmed the crystalline structure of ZnO nanoparticles with identified planes. Salinity stress significantly reduced plant metrics and altered ion ratios, while increasing oxidative stress indicators and osmolytes. Conversely, Zn and ZnO-NPs mitigated these effects, reducing oxidative damage and enhancing enzyme activities. This supports Zn’s role in limiting salinity uptake and improving physiological responses in quinoa seedlings, suggesting a promising strategy for enhancing crop resilience. Overall, this study underscores nanomaterials’ potential in sustainable agriculture and stress management.

1. Introduction

It has been documented that the adverse effects of environmental stressors on agricultural productivity are expected to escalate globally because of shifting environmental and climate conditions [1]. Approximately 10% of the world’s arable land is estimated to be free from stress factors, highlighting the widespread challenge of environmental constraints [2]. Among these, soil salinity emerges as a significant limiting factor, impacting agricultural productivity and posing a threat to global food security. It is estimated that over 800 million hectares of land are afflicted by salinity, constituting approximately 30% of cultivated areas and 50% of irrigated lands [3]. Plants exhibit various responses to salinity stress, encompassing cellular, genetic, physiological, and biochemical mechanisms [4]. The detrimental impact of salt stress on plant growth and development can vary depending on factors such as salt concentration, salt content, plant characteristics, duration of stress, and stage of plant development [5].
Among abiotic stresses, salt stress stands out as one of the most severe environmental factors affecting plants, triggering osmotic and ionic stresses that impede normal cell growth and division [6]. Salt stress additionally induces oxidative stress, resulting in substantial increases in the accumulation of reactive oxygen species (ROS) within the cell. These combined effects can cause metabolic dysfunction, including membrane damage, nutrient imbalance, enzymatic inhibition, and impaired photosynthesis, ultimately culminating in plant mortality [7,8]. Salinity detrimentally impacts osmotic and oxidative stress in plants. Reactive oxygen species (ROS) serve as crucial signaling molecules that contribute to the degradation of photosynthetic pigments in plants [9]. Osmotic adaptation plays a vital role in enhancing plant development parameters and is crucial for maintaining cell turgor, thereby positively impacting plant sustainability.
One of the effective strategies for mitigating the adverse effects of salinity is through the accumulation of proline in plant tissues [10]. Proline serves as both an amino acid and an osmo-protectant, playing a crucial role as a signaling molecule that accumulates in the plant cytosol. It contributes to the stabilization and protection of membranes, protein-containing enzymes, and various proteins. Proline regulates plant metabolism under stress conditions by modulating membrane proteins and ROS scavengers, while also maintaining cellular solute homeostasis [11]. Moreover, in coping with stress conditions in plants, compounds such as superoxide dismutase (SOD), peroxidase (POD), and catalase (CAT) possess the capability to scavenge ROS, thereby enhancing plant resistance [12]. Researchers are dedicating significant efforts to mitigate or eliminate the adverse effects of salinity on cultivated plants [13]. Various strategies, including different plant nutrition approaches, balanced irrigation practices, and the application of diverse organic and chemical compounds, have been explored to alleviate and mitigate the negative effects induced by salt stress [9].
In today’s world, ensuring a safe food supply has emerged as a paramount necessity for societies. The rapid expansion of the global population, escalating pollution of natural resources, economic vulnerabilities, and educational deficits exacerbate food insecurity challenges, making the provision of safe food increasingly challenging. Quinoa (Chenopodium quinoa Willd), a natural food boasting high nutritional value, has been recognized by the Food and Agriculture Organization of the United Nations (FAO) as a key crop for ensuring food security in the coming decades [14]. Quinoa has seen a surge in cultivation and widespread production in recent years, emerging as an alternative crop with numerous benefits. Renowned for its exceptional grain quality, quinoa serves as a safe grain alternative for individuals with celiac disease, as its seeds are gluten-free. Moreover, the predominant proteins found in quinoa seeds are albumins and globulins [15]. Additionally, quinoa seeds contain vital compounds such as polyphenols, carotenoids, dietary fiber, and oleic acid. Notably, quinoa stands out for its richness in essential amino acids like histidine and lysine, surpassing many traditional grains in nutritional value [16].
Quinoa boasts higher concentrations of essential amino acids, such as histidine and lysine, compared to numerous grains [17]; furthermore, in certain regions, the young green leaves of quinoa are also utilized as a vegetable for consumption [18]. It is rich in vitamins, proteins, minerals, and antioxidants [19]. Quinoa is often labeled as a “superfood” by some sources, attributed to its exceptional nutritional properties and beneficial impacts on human health. Additionally, quinoa plants possess the ability to accumulate salt ions in their tissues, aiding in the regulation of leaf water potential by sustaining transpiration through cell turgor maintenance [20]. Quinoa exhibits remarkable tolerance to various abiotic stresses, including high salinity environments. However, in conditions with moderate-to-high salt concentrations, salinity can impede germination and seedling development. This inhibition stems from both reduced water potential and elevated levels of salt ions (such as Na+ and Cl) in the environment. Minimizing the adverse impacts of salt stress, particularly during these critical periods, is of utmost importance [21].
Nano-fertilizers (NFs), also known as smart fertilizers, are gaining attention for their potential to boost crop yields and minimize environmental impact. They consist of nanoscale particles, typically under 100 nanometers, made from materials like metal oxides (e.g., iron, zinc, etc.) or organic compounds (e.g., chitosan, humic acid, etc.). NFs enhance nutrient delivery to plants, improving uptake efficiency. While currently limited in commercial availability and scale, NFs are applied in liquid or solid forms, seed coatings, or blended with traditional fertilizers. Research aims to optimize NFs for specific crops, improve soil health, and reduce environmental footprint, requiring ongoing safety and efficacy assessments through laboratory and field trials. Continued development seeks to enhance NF effectiveness, sustainability, and safety post-commercialization [22].
Microelements play a crucial and intricate role in the nutrition programs of cultivated plants [23]. Zinc is a highly significant microelement with essential functions in plants. It serves as a vital component in the structure of enzymes that fulfill numerous key roles within plant tissues. Additionally, zinc is recognized as a structural stabilizer for proteins, membranes, and DNA-binding proteins [24]. The primary challenge that hampers crop production in arid and semi-arid regions is salinity, and when coupled with zinc (Zn) deficiency, it further diminishes crop yield in these areas [25]. Moreover, in recent years, the widespread adoption of nano-Zn-chelated fertilizers has emerged as a popular method to fulfill the zinc requirements of plants [26]. These nanoparticles (NPs), increasingly utilized in agriculture in recent years, offer notable advantages by assisting plants in tolerating abiotic stresses like salinity and mitigating environmental pollution [27]. NPs are typically administered to the leaves of plants, resulting in quicker and more effective outcomes compared to soil-based fertilizer applications. The recognition of NPs’ beneficial effects on cultivated plants and their significant defensive role against abiotic stresses, which are prominent challenges in plant production, has sparked considerable interest in plant research [28,29,30]. According to the literature review, while there are studies on the role of zinc in quinoa plants, such as plant development, seed germination, and drought stress, there are no studies investigating the effects of zinc oxide nanoparticles under different stress conditions. Therefore, there are some gaps that the current study aims to fill in this area. There is not enough information on the evaluation of zinc nanoparticles in quinoa considering their morphological, physiological, and biochemical properties to determine their effects under salt stress. Considering these gaps, this study is expected to provide an important, original, and innovative contribution to the literature. Additionally, based on the factors of this study, the hypotheses are as follows: (a) zinc oxide nanoparticle (ZnO-NP) application will mitigate the negative impacts of salinity stress on quinoa seedling growth, physiology, and biochemistry, (b) quinoa seedlings treated with ZnO-NPs under salinity stress will exhibit enhanced antioxidant enzyme activities and reduced levels of oxidative stress markers compared to untreated seedlings, and (c) ZnO-NPs will improve ion homeostasis in quinoa seedlings under salinity stress, as evidenced by higher K/Na and Ca/Na ratios and the reduced accumulation of sodium ions. Therefore, in this study, to mitigate the risks associated with salinity toxicity, we investigated seedling growth and physiological and biochemical alterations in quinoa induced by ZnO-NPs under salinity stress.

2. Materials and Methods

2.1. Plant Materials, Growth Conditions, and Treatments

The described study was conducted as a pot experiment within the greenhouses of the Field Crops Department at Atatürk University Faculty of Agriculture. The research utilized Chenopodium quinoa L. Cv Atlas as the experimental plant material.
To induce salt stress, solutions with three different NaCl concentrations (0, 100, and 200 mM) were prepared [31]. Additionally, Zn and ZnO-NPs were administered at four different concentrations (0, 50, 100, and 200 ppm) [32]. Initially, quinoa seeds were sown in 36-compartment plastic vials containing a growing medium composed of a peat/perlite mixture (1:1, v:v). These seeds were nurtured in the greenhouses under salt-free conditions, using regular irrigation water, until they developed 3–4 leaves. Subsequently, seedlings were transplanted into perforated pots filled with soil/peat/sand (2:1:1, v:v:v) with a volume of 1.3 L (14.5 × 12.5 cm). The experiment was designed based on a factorial completely randomized design (CRD), with three replications and 4 plants per pot, totaling 288 plants across the NaCl, Zn, and ZnO-NPs treatments. Throughout the experiment, the greenhouse maintained an average temperature of 32 ± 1 °C and humidity of 55 ± 5%. Seedlings received initial watering with tap water after transplanting. Salt stress applications commenced at a dose of 25 mM to mitigate sudden stress in the root zone, gradually increasing to the final predetermined doses. Foliar application of Zn and ZnO-NPs solutions, prepared at concentrations, was conducted by spraying the seedling leaves. Control doses of Zn and ZnO-NPs were administered using distilled water. Each application involved 50 mL per seedling and was initiated one day after planting, repeated three times with 1-week intervals.

2.2. Synthesis of Zn Nanoparticles (ZnO-NPs)

For the synthesis of ZnO nanoparticles, Zn (NO3)2 (0.1 M) stock solutions were prepared by mixing them in 50 mL of distilled water. Plantain (Plantago major) extract was added to the solution and mixed at 500 rpm at 85 °C for 2 h to reduce it to ZnO using the green synthesis method. Then, after adding 1 mL NH3 to the reaction medium, it was transferred to Teflon-lined, leak-proof stainless steel autoclaves and kept at 180–200 °C for 8 h. They were then allowed to cool naturally to room temperature. After the reaction was completed, the resulting solid ZnO-NPs were washed with ethanol, filtered, and then dried under vacuum in an oven at 60 °C [33,34].

2.3. Morpho-Physiological and Biochemical Properties

After 4 weeks of planting, the study started, and various morphological, physiological, and biochemical analyses were conducted. These analyses included the measurements of plant height, stem diameter, and both fresh and dry plant weights. Additionally, chlorophyll readings were taken using a portable SPAD-502 chlorophyll-measuring device (SPAD-502; Konica Minolta Sensing, Inc., Sakai Osaka, Japan). This device measures the greenness of the middle leaves of the quinoa seedlings, providing insights into their chlorophyll content and photosynthetic efficiency [35]. After harvesting, the plants were carefully transported from the pots to the laboratory. Upon arrival, their fresh weight was promptly measured (g plant−1). Subsequently, the plants were dried at 75 °C to constant weight and weighed again to determine their dry weight (g plant−1), providing valuable data on plant biomass. For biochemical analyses, fresh samples were collected and stored at −80 °C to maintain their integrity until further processing. Dry shoot samples were then ground to a fine powder to facilitate the analysis of sodium (Na), potassium (K), and calcium (Ca) contents [36] within the quinoa seedlings.

2.4. K/Na and Ca/Na Ratios

Following the grinding of the dry samples (0.5 g), they were combined with a mixture of nitric acid and hydrogen peroxide (in a ratio of 2:3) as per the protocol outlined by Mertens [37]. These samples were then subjected to incineration in a microwave wet combustion unit capable of withstanding pressures up to 40 bar. After the samples had cooled, 10 mL of distilled water was added and thoroughly mixed. The resulting sediment was then filtered using filter paper, and the filtrate was diluted to 10 mL with distilled water. Subsequently, the extract was analyzed using Inductively Coupled Plasma Atomic Emission Spectrometry (ICP AES) to determine the sodium (Na), potassium (K), and calcium (Ca) contents within the samples, following the methodology described by Mertens [38].

2.5. Antioxidant Enzyme Activities (CAT, POD, and SOD)

A 0.5 g quantity of frozen quinoa leaves was pulverized into powder using liquid nitrogen and then homogenized with 1.5 mL of extraction solution containing 100 mM K-PO4 buffer (pH 7.0), 1% (w/v) polyvinylpolypyrrolidone (PVPP), and 1 mM EDTA·Na2. The homogenates were centrifuged at 15,000 rpm for 15 min at 4 °C. The resulting supernatants were utilized for assessing the activities of SOD, POX, and CAT enzymes, as described by Turan et al. [10]. The CAT activity was determined following the method outlined by Liu et al. [39]. This method relies on monitoring the absorbance change over one minute at 240 µM, while hydrogen peroxide (H2O2) is being reduced by catalase (CAT). The reaction initiates upon the addition of the enzyme extract and is conducted at 25 °C for 3 min, with readings taken at 1 min intervals. CAT activity is quantified by the decrease in absorbance of H2O2 (with an extinction coefficient ε = 39.4 mM−1 cm−1) at 240 µM, and the results are expressed as micromoles of H2O2 decomposed per minute per gram of leaf fresh weight. Additionally, peroxidase (POD) activity was determined at 436 µM by measuring the conversion of guaiacol to tetra guaiacol (with an extinction coefficient ε = 26.6 mM−1 cm−1), following the protocol described by Liu et al. [39]. The reaction mixture consisted of 100 mM K-phosphate buffer (pH 7.0), 10 mM H2O2, 20.1 mM guaiacol, and the enzyme extract. The absorbance increase upon the addition of H2O2 was monitored at 436 µM for a duration of 5 min. This method relies on the capability of superoxide dismutase (SOD) to impede the photochemical reduction of nitro tetrazolium blue chloride (NBT) at 560 µM, as described by Liu et al. [39]. The reaction mixture comprised 50 mM K-phosphate buffer (pH 7.8), 13 mM methionine, 75 μM NBT, 0.1 μM EDTA, 4 μM riboflavin, and an appropriate quantity of the enzyme extract. Riboflavin was added to initiate the reaction, and the microplate was then incubated for 10 min under a fluorescent lamp. A complete reaction mixture that remained unirradiated was used as the blank. SOD activity was quantified by defining one unit as the amount of enzyme necessary to induce a 50% inhibition of NBT reduction, as observed at 560 µM.

2.6. H2O2, MDA, Sucrose, and Proline

The quantification of malondialdehyde (MDA) and hydrogen peroxide (H2O2) in quinoa leaf tissues was conducted following the protocol outlined by Ohkawa et al. [40]. Frozen quinoa leaf samples from each treatment were ground into a fine powder using liquid nitrogen. Approximately 0.3 g of fresh leaf samples was then homogenized in a mortar with 6 mL of 5% trichloroacetic acid (TCA). After centrifugation at 4100 rpm at +4 °C for 20 min, the collected supernatant was utilized for H2O2 and MDA analyses. For MDA measurement, 0.5 mL of the supernatant was mixed with 1 mL of 0.5% thiobarbituric acid (TBA) + 20% TCA solution, and 0.5 mL of 0.1 M Tris buffer (pH 7.6) was added. The reaction mixture was then incubated in a water bath at 95 °C for 60 min. After cooling the tubes in an ice bath to halt the reactions, the absorbance at 532 µM and 600 µM wavelengths was recorded using a spectrophotometer. The MDA concentration was calculated as mmol kg−1 FW using the formula [(absorbance 532 µM − absorbance 600 µM)/1.55 × 105] with an extinction coefficient of 155 mM−1 cm−1 [41].
For H2O2 measurement, 1 mL of 10 mM phosphate buffer (pH = 7) and 1 mL of 1 M KI were added to 0.5 mL of supernatant. The absorbance of the mixture was measured at 390 µM using a spectrophotometer. The H2O2 concentration in the leaf tissues was determined as mmol kg−1 FW with reference to a standard graph constructed using 0–50 μmol H2O2 concentrations [42].
Additionally, samples freeze-dried at −80 °C were ground to pass through a 1 mm mesh sieve. Leaf samples weighing 0.1 g from each treatment were mixed with 10 mL of 80% (v/v) ethyl alcohol and incubated in a 60 °C water bath for 1 h. Ethyl alcohol in the suspension was evaporated in a water bath at 60 °C. The resulting precipitate was dissolved in 1 mL of distilled water and centrifuged at 10,000 rpm at 4 °C for 10 min. Subsequently, 0.1 mL of the centrifuged sample was mixed with an equal amount of 30% potassium hydroxide (KOH) and incubated in boiling water for 10 min. After cooling to room temperature, 300 μL of distilled water was added. Sucrose standards were prepared, and 2500 μL of anthrone solution was added to both the samples and standards. The mixture was left to stand at room temperature for 1 h, and the amount of sucrose in the samples was determined in a spectrophotometer at a wavelength of 620 µM using a 96-well microplate [43].
Proline extraction and determination of proline content were conducted according to the method developed by Bates et al. [44]. The reaction relies on the formation of a pink-colored compound resulting from the color reaction of proline with ninhydrin. Leaf samples frozen at −80 °C were ground using liquid nitrogen. A 0.5 g quantity of leaf tissue was placed in test tubes and homogenized by adding 3% (w/v) aqueous sulfosalicylic acid. The mixture was then centrifuged at 10,000 rpm for 10 min. Subsequently, 2 mL of the extract was transferred to a test tube, and the reaction was initiated by adding 2 mL of freshly prepared 2.5% acid–ninhydrin solution and glacial acetic acid. The test tubes were then incubated at 100 °C for 60 min to allow the reaction to proceed. Upon completion, the reaction was terminated in an ice bath, and 5 mL of toluene was added to the mixture to separate the chromophores. The proline content of the toluene phase was measured at 520 µM using the spectrophotometric method in a 96-well microplate. Proline concentration was calculated using the formula [(μg proline in extract/115.5)/g sample], and the results were expressed as mmol kg−1 FW.

2.7. Statistical Analysis

All the data obtained from this research were subjected to the analysis variance based on the experimental design using RStudio statistical analysis software program, Version: 2023.12.1+402 “doebioresearch” package, and the differences among the means were compared according to Fisher’s Least Significant Difference (Duncan Multiple’s Range Test) (0.05).

3. Results

3.1. Characterization of ZnO-NPs

In Figure 1A, the TEM image of the synthesized ZnO nanoparticles is displayed, showcasing a magnification of approximately one million. Observations revealed that the ZnO NP structures exhibited a spherical morphology, with sizes ranging between 15 and 35 nm. Notably, Figure 1A illustrates that the surfaces of ZnO NPs appear rough. These findings align closely with the existing literature [33,34,35,36].
Utilizing Cu Kα radiation with a wavelength of 1.5406 Angstrom, the X-ray diffraction data were meticulously recorded over a 2θ range of 20–80°. The XRD analysis graph, which reveals the crystallized structure of ZnO nanoparticles, identified the 100, 002, 101, 102, 110, and 103 planes. The positions at 29.45°, 35.42°, 39.02°, 47.92°, 55.64°, and 63.57° correspond to the respective plane distances of the atoms in the ZnO nanoparticle structure. The average grain size of the samples was determined using the Scherrer equation, based on the diffraction intensity of the (101) peak. The X-ray diffraction analysis unequivocally affirmed the synthesis of ZnO materials, showcasing a wurtzite phase. All discernible diffraction peaks matched the reported JCPDS data precisely, with no characteristic peaks observed other than those of ZnO. The mean grain size (D) of the particles was deduced from XRD line-broadening measurements using the Scherrer Equation (1):
D = 0.89λ/(βCosθ)
Here, λ represents the wavelength (Cu Kα), β signifies the full width at half-maximum (FWHM) of the ZnO (101) line, and θ denotes the diffraction angle. The evident line broadening of the diffraction peaks indicates that the synthesized materials are within the nanometer scale. Additionally, the calculated lattice parameters closely aligned with the reported values.

3.2. Plant Growth

Analysis of variance (Table 1) showed that there were significant differences in the three treatments (salinity, zinc, and concentration) pertaining to the plant height, plant weight, plant diameter, and chlorophyll content.
The mean comparison results for salinity indicated that the highest means for plant height, plant weight, plant diameter, and chlorophyll content were 10.10 ± 1.50 cm, 1.34 ± 0.23 g, 1.73 ± 0.24 mm, and 38.13 ± 2.59 SPAD, respectively, observed at 0 µM salt concentration. Conversely, the lowest means of 7.94 ± 1.19 cm, 0.71 ± 0.17 g, 1.58 ± 0.32 mm, and 30.67 ± 1.86 SPAD, respectively, were observed at 200 mM NaCl (Table 2). Based on zinc application, Zn-NPs treatment gave the highest mean plant height, plant weight, plant diameter, and chlorophyll content of 9.35 ± 1.29 cm, 1.04 ± 0.35 g, 1.82 ± 0.15 mm, and 35.51 ± 3.59 SPAD, respectively, whereas the lowest means were observed in the treatment of Zn of 8.54 ± 1.82 cm, 0.79 ± 0.33 g, 1.45 ± 0.27 mm, and 33.06 ± 3.65 SPAD, respectively (Table 2). In terms of the Zn concentrations, the highest values were observed in the treatment with 200 µM concentration for all the characteristics, i.e., plant height (9.22 ± 1.73 cm), plant weight (0.99 ± 0.38 g), plant diameter (1.74 ± 0.28 mm), and chlorophyll content (35.00 ± 4.73 SPAD) (Table 2).
Analysis of variance indicated that there were significant two-way interactions between salinity and zinc (S × Z) interaction in terms of plant height; salinity and concentration (S × C) in terms of plant height, plant weight, and chlorophyll content; and zinc and concentration (Z × C) in terms of plant weight (Table 1). Each salinity stress has a different response to the zinc application, which makes the salinity × zinc interaction significant. In terms of the S × Z interaction, the highest plant height (10.42 ± 1.20 cm) was obtained from 0 mM × Zn-NPs treatment; in terms of the S × C interaction, the highest plant height (11.17 ± 0.93 cm), plant weight (1.49 ± 0.14 g), and chlorophyll content (41.08 ± 1.37 SPAD) were observed in 0 mM × 200 ppm treatment; thus, in terms of the Z × C interaction, the highest plant weight (1.04 ± 0.35 g) was determined from the Zn × Zn-NPs interaction (Table 2). Under the three-way interaction of salinity, zinc, and concentration (S × Z × C); the effect of the three-way interaction was insignificant in all the characteristics except its effect on the plant weight, which was significantly affected. The highest plant height was measured to be 11.17 ± 1.04 cm in the control and 200 µM Zn and 200 µM Zn-NPs without NaCl treatment, while the lowest was measured to be 6.50 ± 0.87 cm at 0 µM Zn under 200 mM NaCl treatment (Table 2).
The application of Zn and ZnO-NPs generally has a positive effect on plant height, weight, diameter, and SPAD parameters in the absence of salt stress. The most significant improvements in these traits were observed at 200 ppm for both Zn and ZnO-NP applications, with ZnO-NPs yielding better results than Zn. Although salt stress negatively impacted these morphological traits, the 200 ppm concentration of Zn and ZnO-NPs stood out in mitigating the stress effects.

3.3. K/Na and Ca/Na Ratios

Analysis of variance showed that there were significant differences in the three main treatments (salinity, zinc, and concentration) on K/Na and Ca/Na (Table 3). The results of mean value comparison of salinity showed that the highest value of K/Na (50.45 ± 4.36) and Ca/Na (79.53 ± 25.39) was determined at 0 mM salt whereas the lowest was obtained at 200 mM salinity treatment (Table 4). Based on zinc application, Zn-NPs treatment gave the highest K/Na (37.47 ± 13.40) and Ca/Na (62.59 ± 32.71), whereas the lowest mean was observed in the treatment of Zn with 35.30 ± 10.50 and 53.20 ± 13.51, respectively (Table 4). When we look at the concentrations, the highest values were observed from 200 concentration treatment with the values of 38.42 ± 11.73 and 60.62 ± 27.03 for both K/Na and Ca/Na, respectively (Table 4).
Analysis of variance indicated that, while there is a significant difference in the two-way interactions between salinity and zinc (S × Z) and salinity and concentration (S × C) interactions in terms of K/Na and Ca/Na, the interaction of zinc and concentration (Z × C) showed nonsignificant differences (Table 3). Based on the mean values of the interactions, the highest values of K/Na (52.34 ± 4.10) and Ca/Na (103.99 ± 5.80) were determined from 0 mM × Zn-NPs interaction, whereas the lowest values were obtained from 200 mM × Zn-NPs. The salinity × zinc interaction showed a significant difference in both K/Na and Ca/Na. For the S × Z interaction, the highest values of K/Na (39.49 ± 13.40) and Ca/Na (64.95 ± 34.80) were obtained for Znt × Zn-NPs (Table 4).
The three-way interaction of different salinity levels, zinc, and concentration (S × Z × C) was highly significant in all the characteristics mentioned in Table 4. The highest K/Na and Ca/Na were measured at 54.96 ± 1.45 and 107.66 ± 6.03 from 0 mM × Zn-NPs × 50.0 ppm and 0 mM × Zn-NPs × 200 µM interactions, respectively (Table 4).
In general, the results indicate that stress conditions impair ion uptake in plants, leading to a decrease in the K/Na and Ca/Na ratios. However, both Zn applications (Zn and ZnO-NPs) proportionally increased these ratios. Based on the average values, the 200 ppm ZnO-NPs application has the most positive effect on these parameters and is recommended for use in quinoa.

3.4. Antioxidant Enzyme Activities

Analysis of variance showed that there were significant differences in the three main treatments (salinity, zinc, and concentration) and both the two-way and three-way interactions on the plant SOD, CAT, and POD (Table 5). The results of mean comparison of salinity showed that the highest mean of the SOD (472.14 ± 93.46), CAT (7378.96 ± 1127.04), and POD (242.87 ± 19.26) was determined for 200 mM salt, whereas the lowest was determined in 0 mM NaCl treatment (Table 6). Based on zinc application, Zn treatment gave the highest SOD, CAT, and POD with the means of 329.12 ± 148.77, 6267.71 ± 1371.00, and 210.59 ± 63.00, respectively, whereas the lowest mean was observed in the treatment of Zn-NPs (Table 6). In terms of the concentrations, the highest values of 344.90 ± 154.33, 6146.81 ± 1310.19, and 212.51 ± 59.67 were observed for 0 µM concentration treatments, respectively (Table 6).
The means of the salinity × zinc interaction for SOD, CAT, and POD varied between 189.30 ± 9.09 and 526.94 ± 22.9, 792.21 ± 389.57 and 7888.43 ± 550.24, and 128.18 ± 17.28 and 270.97 ± 13.04 and the lowest means for SOD, CAT, and POD were obtained for 0 mM × Zn treatment; in terms of the S × C interaction; the highest means for SOD (541.35 ± 17.54), CAT (7709.82 ± 412.53), and POD (250.59 ± 22.54) were observed for the 200 mM × 0 µM interaction, 0 mM × 0 µM interaction, and 0 mM × 100 ppm treatment; thus, in terms of the (Z × C) interaction, the lowest means for SOD (343.09 ± 152.23), CAT (6035.46 ± 1520.45), and POD (212.14 ± 56.15) were determined for 0 µM Zn (Table 6).
On other hand, for the three-way interaction of salinity, zinc, and concentration (S × Z × C), the three-way interaction had an insignificant effect on SOD, CAT, and POD (Table 5).
The highest means for SOD (556.58 ± 6.08) and CAT (8557.88 ± 84.03) were measured for the 200 mM × Zn × 100 µM treatment, and the highest mean for POD (283.29 ± 7.40) was obtined for the 100 mM × Zn × 50 µM treatment, while the lowest mean was measured for the 100 mM × Zn-NPs × 0 µM (175.89 ± 7.11), 0 mM × Zn-NPs × 100 µM (4461.25 ± 319.26), and 0 mM × Zn × 100 µM (110.17 ± 8.55) treatment (Table 6).
The study results show that salt stress increases enzyme levels, indicating an activation of the defense mechanism in plants. Applying Zn and ZnO-NPs to quinoa plants positively affected enzyme activity. Based on average values, 50 ppm Zn and 50 and 100 ppm ZnO-NPs were more effective. However, it is important to evaluate the most appropriate design proposal based on morphological characteristics. Therefore, applying 100 and 200 ppm ZnO-NPs might be more suitable for enhancing both enzyme activity and morphological development.

3.5. Hydrogen Peroxide (H2O2), Melondialdehyde (MDA), Proline, and Sucrose Contents

Analysis of variance showed that there were significant differences in the three main treatments (salinity, zinc, and concentration) on the H2O2, MDA, proline, and sucrose contents (Table 7). The results of mean comparison of salinity showed that the highest mean for H2O2 (12.79 ± 1.77), MDA (8.34 ± 1.01), proline (257.77 ± 45.14), and sucrose (37.79 ± 5.54) was determined for 200 mM salt, whereas the lowest mean for H2O2 (9.34 ± 1.02), MDA (6.16 ± 1.15), proline (164.45 ± 26.63), and sucrose (32.33 ± 6.11) was observed in 0 mM NaCl (Table 8). Based on zinc application, Zn treatment gave the highest means for H2O2 (11.03 ± 2.17), MDA (8.05 ± 1.08), proline (216.31 ± 61.97), and sucrose (36.73 ± 6.89), whereas the lowest means of 10.46 ± 1.64, 6.44 ± 1.73, 190.75 ± 54.00, and 34.04 ± 6.67 for Zn-NPs (Table 8). In terms of the concentrations, the highest means of 11.47 ± 1.98, 8.03 ± 1.34, 215.24 ± 59.45, and 39.46 ± 5.83 were observed for 0 µM concentration treatments, respectively (Table 8). Analysis of variance showed that there were significant differences in the interactions both two-way and three-way interactions for H2O2, MDA, proline, and sucrose (Table 7).
The means of the salinity × zinc interaction for H2O2, MDA, proline, and sucrose contents varied between 5.37 ± 0.36 and 8.60 ± 0.52, 9.21 ± 1.23 and 13.94 ± 0.48, 147.77 ± 24.90 and 263.32 ± 51.93, and 27.88 ± 1.86 and 38.67 ± 6.79, and the highest means for MDA, H2O2, proline, and sucrose were obtained for 0 mM × Zn-NPs, 0 mM × Zn, 100 mM × Zn-NPs, and 0 mM × Zn-NPs interactions; in terms of the (S × C) interaction, the highest means for MDA (9.22 ± 0.58), H2O2 (14.05 ± 0.19), proline (270.38 ± 25.85), and sucrose (41.74 ± 2.73) were observed for the 200 mM × 50 ppm and 200 mM × 0 ppm interactions, respectively; thus, in terms of the (Z × C) interaction; the highest means for H2O2 (11.53 ± 2.03), MDA (8.36 ± 0.87), proline (249.10 ± 76.13), and sucrose (43.04 ± 2.43) were determined for the Zn-NPs × 0 ppm, Zn × 50 ppm, Zn × 50 ppm, and Zn × 0 ppm interactions, respectively (Table 8).
On other hand, for the three-way interaction of salinity, zinc, and concentration (S × Z × C), the three-way interaction had an insignificant effect on MDA, H2O2, proline, and sucrose (Table 7). The highest means for H2O2 (14.52 ± 0.13), MDA (9.55 ± 0.10), proline (315.03 ± 20.08), and sucrose (48.58 ± 1.68) were measured for the 200 mM × Zn × 100 ppm interaction, 0 mM × Zn × 0.0 ppm interaction, 0 mM × Zn × 200.0 ppm interaction, 200 mM × Zn-NPs × 50 ppm interaction, 200 mM × Zn × 50 ppm, and 100 mM × Zn × 50 ppm, while the lowest means of 7.41 ± 0.32 (100 mM × Zn × 100 ppm), 4.51 ± 0.08 (100 mM × Zn-NPs × 50 ppm), 132.17 ± 7.66 (100 mM × Zn-NPs × 50 ppm), and 24.67 ± 1.96 (100 mM × Zn × 200 ppm) were measured, respectively (Table 8).
Salinity is a stress indicator for plants, and one of its detrimental effects is the accumulation of reactive oxygen species such as H2O2 and MDA, along with oxidative stress. Other substances that accumulate due to stress are proline and soluble sugars. Our study findings show that H2O2, MDA, proline, and sucrose accumulate in quinoa plants due to stress, which is consistent with previous studies. Our study found that the accumulation of these substances decreased after the application of Zn and ZnO-NPs. The average values indicate that applying 50 ppm Zn and 200 ppm ZnO-NPs effectively reduces the accumulation of these substances. Overall, it can be stated that ZnO-NP applications offer the most effective reduction and can be used in practice.

4. Discussion

Salinity stress poses a significant challenge, jeopardizing crop production worldwide and exerting detrimental effects on plant development [1]. In plants subjected to salt stress, nutrient uptake is hindered, leading to oxidative stress and substantial damage. This damage is primarily caused by the accumulation of toxic concentrations of sodium and chloride within the cells [45]. An early indication of salt stress in seedlings is the reduction in leaf area, suggesting that initial plant growth is adversely affected due to alterations in plant water balance [46]. Salt stress has been documented to induce a notable decrease in the morphological development of plants, including reductions in plant height and stem diameter [47]. Moreover, the salt content exerts a negative impact on plant growth by impeding assimilation processes within the plants and causing a significant reduction in plant weight [48].
In the study, the detrimental impact of salt stress on quinoa seedling growth was evident, manifested by the suppression of plant height and other developmental traits. A negative correlation between salt concentration and quinoa seedling growth characteristics was observed, with a significant decrease in plant height as salt concentration increased. Numerous studies have demonstrated that saline conditions disrupt plant ion balance and osmoregulation, consequently adversely affecting plant height, stem diameter, and overall biomass [41,42,43]. In a study, researchers examined the effects of zinc sulfate application on some physiological and morphological properties of quinoa (Chenopodium quinoa Willd) under drought stress. At the end of the research, they suggested that zinc sulfate reduced the damage caused by drought stress by improving the osmolytic level in plants and recommended the use of 6 μM zinc sulfate in nutrient solutions to alleviate the effect of drought stress [49]. In another study by Jorfi et al. [50], researchers investigated the root system development of quinoa affected by phosphorus and zinc sulfate application in alkaline soil. In the study, triple superphosphate fertilizer (0, 6, 12, 18 mg/kg soil) and zinc sulfate foliar fertilizer (0, 4, and 8 g/L) were applied to three quinoa varieties. According to the triple interaction, the applications that had the best effect on root properties were 18 mg/kg of triple superphosphate and 4 g/L of zinc sulfate. It was concluded that better performance is achieved when optimal amounts of macro/micro elements are used in alkaline soils by selecting varieties with larger root systems and higher nutrient uptake potential. In a similar study, the effects of foliar iron and zinc applications on seed germination and biochemical properties of quinoa under drought stress were determined. It was concluded that both iron and zinc applications positively affected seed germination under stress conditions [51].
In this study, both Zn and ZnO-NP applications exhibited a positive impact on plant height under salt stress conditions. When compared to Zn applications alone, ZnO-NPs demonstrated greater efficacy in enhancing plant height. This heightened effectiveness of ZnO-NPs can be attributed to their smaller size, which facilitates enhanced uptake, transportation, metabolism, and accumulation within plant cells [52]. In a similar study, the application of ZnO-NPs to tomato plants was found to enhance plant growth by mitigating the adverse effects of salt stress when compared to the control group [53]. Scientific studies indicate that the application of zinc enhances plant resistance to salt stress, and ZnO-NPs can aid plants in coping with the adverse effects of abiotic stress induced by severe soil salinization [54]. The study found a general decrease in the weight of quinoa plants under salt stress. After Zn and ZnO-NP treatments, there was an improvement and increase in plant weight, along with the alleviation of salt-induced stress. In the study conducted by Adil et al. [53], it was observed that the NaCl concentration in barley plants decreased as the concentration increased. The result showed that, following the application of ZnO-NPs, a notable enhancement in the dry biomass of barley plants under salinity stress was achieved. Another study demonstrated that the foliar application of ZnO-NPs resulted in a significant increase in plant growth and biomass [55].
Our findings indicated that SPAD values decreased with increasing salinity stress; however, the use of Zn-NPs mitigated the negative effects of salt. In this study, it was found that both Zn and ZnO-NP treatments led to a similar improvement in SPAD values. However, based on the findings, it can be concluded that ZnO-NPs were more effective. Salt stress induces structural damage to chloroplasts in plants, leading to a significant decrease in photosynthetic efficiency. One of the observed changes is the structural deterioration of leaf stomata. This damage to stomatal structures adversely affects CO2 uptake, consequently reducing the rate and efficiency of photosynthesis. Additionally, salt stress disrupts the number of chloroplasts and the photosynthetic pigment–protein mechanism responsible for photosynthesis. Furthermore, salt stress has a detrimental effect on photosynthetic pigments and plant activity, resulting in a negative impact on SPAD values. In a study, it was noted that the decline in SPAD values caused by salt stress was reversed by ZnO-NP treatment [56]. Singh et al. [57] reported that the application of ZnO-NPs to rice seedlings under salt stress led to an increase in chlorophyll a, chlorophyll b, and total chlorophyll content, resulting in improved photosynthesis. Similarly, in a study on rice plants, Zn application was applied to mitigate the negative effects of salt stress, and it was found to have a positive effect on SPAD values measured from leaves [58]. Additionally, ZnO-NP applications under saline conditions in species such as pepper [59], radish [60], and wheat [61] were reported to positively impact SPAD values, consistent with other studies. The findings from these studies are in line with the data obtained in the present study.
The most significant effects of salt stress on plant cells are both osmotic and ionic, which have a detrimental impact on cell metabolism. Under saline conditions, specific metabolic effects occur in plants, primarily attributed to the accumulation of toxic ions such as Na+ and Cl, or the reduction in essential elements like K+ and Ca2+. Additionally, plant lipid metabolism is adversely affected due to disruptions in ion uptake or balance [46]. Therefore, maintaining optimal ion uptake and balance is crucial for plant survival in saline environments [62].
Quinoa seedlings subjected to salt stress conditions exhibited lower K/Na and Ca/Na ratios compared to those not exposed to stress. Studies have indicated that this phenomenon is attributed to the increased accumulation of Na+ in the root zone under saline conditions, subsequently leading to deficiencies of Ca2+ and K+ in the plants [63,64,65]. Potassium is the predominant cation in plant cells and is vital for essential physiological processes such as photosynthesis and various enzymatic metabolic functions [57,66]. Following the application of Zn and ZnO-NPs to quinoa plants exposed to saline conditions, an increase in potassium and calcium uptake was observed, resulting in improved K/Na and Ca/Na ratios. Researchers have also found that the application of ZnO-NPs on Zea mays and Gossypium spp. plants under salt stress led to a reduction in the Na+ ratio and an increase in the K+/Na+ ratio, consequently mitigating the negative effects of salinity stress and promoting positive plant growth [67,68]. In plants treated with Zn, ZnO-NPs, and salt, there was a general increase in the activity of antioxidant enzymes such as SOD, POX, and CAT compared to plants not exposed to stress. Similar studies have demonstrated that the foliar application of ZnO-NPs enhances the activity of antioxidant enzymes such as SOD, POD, and CAT in the presence or absence of salt in the environment [56,69].
The results of the study show that salt stress in quinoa seedlings causes an increase in H2O2 and MDA levels, which are indicators of oxidative stress. In this study, Zn and ZnO-NPs applied to quinoa plants under salt stress increased the efficacy of antioxidant enzymes. Looking at the relative efficacy, ZnO-NPs seem to be more beneficial. Excessive ROS accumulation disrupts both the protein structure and the integrity of cell membranes and increases membrane permeability [10]. Studies have reported that ZnO-NPs applied under salt stress reduce the MDA concentration, which helps protect cell membrane integrity and reduce the negativities caused by oxidative stress [70]. In the study conducted by Yasmin et al. [71], it was reported that safflower plants exposed to salt stress exhibited an increase in antioxidant enzymes such as SOD, CAT, POD, and APX, and ZnO-NP applications mitigated the effects of salt stress. Similarly, in another study, it was found that ZnO-NP applications led to an increase in antioxidant enzymes in okra plants under salt stress [72].
In our study, there was a decrease in proline and sucrose sugar contents in quinoa plants under salt stress conditions, and it can be assumed that many applications of Zn and ZnO-NPs on the leaves reduced this accumulation by decreasing the proline and sucrose contents. In the study conducted by Ahmad et al. [73], there was a significant decrease in H2O2 production after foliar Zn application to Brassica juncea seedlings under salinity stress.

5. Conclusions

Although the effect of zinc on quinoa plants has been examined in many studies, there is a gap in the research concerning the application of nanoparticles, a more effective and recent approach, on quinoa plants under stress conditions. Therefore, our study investigates the effects of ZnO nanoparticles (ZnO-NPs) on quinoa seeds under salinity stress and aims to understand their impact on seedling growth, physiological parameters, and antioxidant activity. Salinity stress adversely affected various quinoa plant metrics, but the application of Zn and ZnO-NPs mitigated these effects, with ZnO-NPs showing significant improvements. X-ray diffraction confirmed the synthesis of crystalline ZnO nanoparticles with a wurtzite phase structure, identifying specific crystallographic planes. Overall, ZnO-NPs positively influenced all measured parameters, including antioxidant enzyme activities and biochemical markers, suggesting their potential in enhancing plant resilience under stress conditions. Further research into the application of ZnO-NPs in nano-fertilizers or other innovative approaches could aid in improving crop performance and stress resistance.

Author Contributions

Conceptualization, A.T. and K.H.; methodology, M.E., M.T., E.Y. and G.N.; software, A.T., A.A.L.S. and G.N.; validation, A.T., M.P. and G.N.; formal analysis, A.T., K.H., M.E., E.Y., H.İ.Ö., A.A.L.S., H.N., M.P. and G.N.; investigation, A.T. and A.A.L.S.; resources, A.T., K.H., E.Y., H.N. and G.N.; data curation, A.T., K.H., E.Y. and M.P.; writing—original draft preparation, A.T., M.P. and G.N.; writing—review and editing, A.T., H.İ.Ö., M.P. and G.N.; visualization, A.T., M.T. and E.Y.; supervision, A.T. and G.N.; project administration, A.T.; funding acquisition, A.T., K.H. and M.T. All authors have read and agreed to the published version of the manuscript.


This research received no external funding.

Data Availability Statement

The original contributions presented in the study are included in the article, further inquiries can be directed to the corresponding authors.


We thank the anonymous reviewers and editors for their constructive comments on this manuscript.

Conflicts of Interest

The authors declare no conflicts of interest.


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Figure 1. (A) SEM image of ZnO-NPs synthesized by hydrothermal method. (B) XRD pattern of ZnO nanoparticles.
Figure 1. (A) SEM image of ZnO-NPs synthesized by hydrothermal method. (B) XRD pattern of ZnO nanoparticles.
Agronomy 14 01462 g001
Table 1. Analysis of variance of effects of different salinity levels, Zn treatments, and Zn levels on plant height (cm), plant weight (g), plant diameter (mm), and chlorophyll content (SPAD) of quinoa seedling.
Table 1. Analysis of variance of effects of different salinity levels, Zn treatments, and Zn levels on plant height (cm), plant weight (g), plant diameter (mm), and chlorophyll content (SPAD) of quinoa seedling.
Sources of VariationPlant
Content (SPAD)
F value (salinity)24.00 ***190.31 ***3.96 *163.27 ***
F value (Zn)9.81 **66.94 ***51.32 ***52.82 ***
F value (concentration)2.15 ns3.40 *2.91 *2.72 ns
F value (S × Z)4.16 *0.55 ns0.39 ns0.01 ns
F value (S × C)3.22 **4.14 **0.34 ns10.09 ***
F value (Z × C)0.92 ns2.71 *0.60 ns0.23 ns
F value (S × Z × C)2.20 *0.57 ns0.32 ns0.25 ns
CV %12.2014.0813.414.18
*, **, ***, ns: significant at p ≤ 0.05, p ≤ 0.01, and p ≤ 0.001 and nonsignificant, respectively.
Table 2. Mean comparison of the effects of different salinity levels, Zn treatments, and Zn levels on plant height, plant weight, plant diameter, and chlorophyll content of quinoa seedling.
Table 2. Mean comparison of the effects of different salinity levels, Zn treatments, and Zn levels on plant height, plant weight, plant diameter, and chlorophyll content of quinoa seedling.
SalinityZn TreatmentsZinc Concentration
Plant Height (cm)Plant Weight (g)Plant Diameter (mm)Chlorophyll Content
0 mMZn0.
Mean9.79 a1.20 b1.5736.93 b
Mean10.42 a1.48 a1.9039.33 a
Mean10.10 a1.34 a1.73 a38.13 a
100 mMZn0.09.830.661.2331.83
Mean8.79 b0.58 d1.3732.81 d
Mean8.79 b0.84 c1.8035.29 c
Mean8.79 b0.71 b1.58 b34.05 b
200 mMZn0.06.500.411.3130.90
Mean7.04 c0.60 d1.4029.43 e
Mean8.83 b0.81 c1.7531.91 d
Mean7.94 c0.71 b1.58 b30.67 c
Means of Zn
Zn0.09.17 ab0.691.2932.27
50.07.78 c0.731.4332.94
100.08.50 bc0.841.4833.01
200.08.72 abc0.911.5834.00
Mean8.54 b0.79 b1.45 b33.06 b
Zn-NPs0.09.33 ab1.081.7835.01
50.09.11 ab0.991.7535.57
100.09.22 ab1.041.8335.47
200.09.72 a1.071.9036.00
Mean9.35 a1.04 a1.82 a35.51 a
Means of Zn concentration (ppm) b1.54 b33.64 b
50.08.440.86 b1.59 ab34.26 ab
100.08.860.94 ab1.66 ab34.24 ab a1.74 a35.00 a
Differences between means shown with the same lowercase letter in the same column for each application are insignificant.
Table 3. Analysis of variance of effects of different salinity levels, Zn treatments, and Zn levels on K/Na value and Ca/Na value of quinoa seedling.
Table 3. Analysis of variance of effects of different salinity levels, Zn treatments, and Zn levels on K/Na value and Ca/Na value of quinoa seedling.
Sources of VariationK/Na Ca/Na
F value (salinity)547.46 ***1548.31 **
F value (Zn)12.26 **179.32 **
F value (concentration)9.61 ***19.67 **
F value (S × Z)129.49 ***798.88 **
F value (S × C)2.26 ns10.57 **
F value (Z × C)0.67 ns2.49 ns
F value (S × Z × C)4.26 **14.95 **
CV %7.215.13
**,***, ns: significant at p ≤ 0.01 and p ≤ 0.001 and nonsignificant, respectively.
Table 4. Mean comparison of the effects of different salinity levels, Zn treatments, and Zn levels on K/Na and Ca/Na ratios of quinoa seedling.
Table 4. Mean comparison of the effects of different salinity levels, Zn treatments, and Zn levels on K/Na and Ca/Na ratios of quinoa seedling.
SalinityZn TreatmentsZinc Concentration (ppm)K/Na ValueCa/Na Value
0 mMZn0.046.98 cd56.15 de
50.050.23 a–d56.54 de
100.046.04 d51.98 e
200.050.93 abc55.58 de
Mean48.55 b55.06 d
Zn-NPs0.048.64 bcd98.67 b
50.054.96 a105.33 a
100.052.22 ab104.29 a
200.053.56 a107.66 a
Mean52.34 a103.99 a
Mean0.047.8177.41 b
50.052.5980.94 ab
100.049.1378.14 ab
200.052.2581.62 a
Mean50.45 a79.53 a
100 mMZn0.022.28 jk55.02 de
50.021.18 jkl54.96 de
100.029.08 i77.97 c
200.029.89 hi76.56 c
Mean25.60 e66.13 b
Zn-NPs0.036.47 fg53.02 e
50.039.80 ef59.50 d
100.038.59 efg59.19 d
200.041.41 e58.83 d
Mean39.07 c57.63 c
Mean0.029.3754.02 d
50.030.4957.23 d
100.033.8468.58 c
200.035.6567.70 c
Mean32.34 b61.88 b
200 mMZn0.030.89 hi39.84 f
50.034.62 gh40.33 f
100.030.32 hi36.79 f
200.031.20 hi36.73 f
Mean31.76 d38.42 e
Zn-NPs0.017.51 l21.12 i
50.018.58 kl24.45 hi
100.024.39 j30.62 g
200.023.52 j28.36 gh
Mean21.00 f26.14 f
Mean0.024.2030.48 e
50.026.6032.39 e
100.027.3533.71 e
200.027.3632.54 e
Mean26.38 c32.28 c
Means of Zn
Mean35.30 b53.20 b
Mean37.47 a62.59 a
Means of Zn concentration (ppm)0.033.79 c53.97 c
50.036.56 b56.85 b
100.036.77 ab60.14 a
200.038.42 a60.62 a
Differences between means shown with the same lowercase letter in the same column for each application are insignificant.
Table 5. Analysis of variance of effects of different salinity levels, Zn treatments, and Zn levels on antioxidant enzyme activities (SOD: EU g−1 leaf; CAT: EU g−1 leaf; POD: EU g−1 leaf) of quinoa seedling under salinity.
Table 5. Analysis of variance of effects of different salinity levels, Zn treatments, and Zn levels on antioxidant enzyme activities (SOD: EU g−1 leaf; CAT: EU g−1 leaf; POD: EU g−1 leaf) of quinoa seedling under salinity.
Sources of VariationSODCATPOD
F value (salinity)9730.50 ***200.56 ***319.76 ***
F value (Zn)853.05 ***39.66 ***59.33 ***
F value (concentration)402.66 ***5.22 **11.26 ***
F value (S × Z)714.33 ***25.64 ***269.76 ***
F value (S × C)145.52 ***5.16 ***9.87 ***
F value (Z × C)130.28 ***4.13 *6.36 **
F value (S × Z × C)195.06 ***10.75 ***10.25 ***
CV %2.387.386.80
*, **, ***: significant at p ≤ 0.05, p ≤ 0.01, and p ≤ 0.001, respectively.
Table 6. Mean comparison of the effects of different salinity levels, Zn treatments, and Zn levels on antioxidant enzyme activities of quinoa seedling under salinity.
Table 6. Mean comparison of the effects of different salinity levels, Zn treatments, and Zn levels on antioxidant enzyme activities of quinoa seedling under salinity.
SalinityZn TreatmentsZinc Concentration (ppm)SOD
(EU g−1 Leaf)
(EU g−1 Leaf)
(EU g−1 Leaf)
0 mMZn0.0182.45 hi4621.41 i128.98 fg
50.0191.29 gh4909.01 ghi124.62 fg
100.0182.91 hi4682.49 hi110.17 g
200.0200.54 g4955.91 ghi148.96 f
Mean189.30 e4792.21 e128.18 f
Zn-NPs0.0327.24 d5905.06 cde244.50 cde
50.0189.96 gh5032.16 f–i129.71 fg
100.0203.34 g4461.25 i134.65 fg
200.0195.71 gh5296.80 e- i142.32 f
Mean229.06 d5173.82 d162.79 d
Mean0.0254.84 d5263.24 def186.74 d
50.0190.62 f4970.58 fg127.17 f
100.0193.13 f4571.87 g122.41 f
200.0198.12 f5126.36 ef145.64 e
Mean209.18 c4983.01 c145.49 c
100 mMZn0.0301.09 ef6573.88 c276.80 ab
50.0295.15 f5712.01 d–g283.29 a
100.0292.48 f5816.25 c–f256.72 bc d
200.0195.75 gh6387.83 cd267.08 ab c
Mean271.12 c6122.49 c270.97 a
Zn-NPs0.0175.89 i4867.29 hi139.39 f
50.0197.69 g4816.63 hi147.22 f
100.0202.05 g4725.70 hi138.48 f
200.0189.79 gh4820.66 hi144.49 f
Mean191.35 e4807.57 e142.39 e
Mean0.0238.49 e5720.59 d208.09 bc
50.0246.42 de5264.32 def215.25 b
100.0247.26 de5270.98 def 197.60 cd
200.0192.77 f5604.24 de205.79 bc
Mean231.24 b5465.03 b206.68 b
200 mMZn0.0556.58 a8557.88 a 232.90 de
50.0524.01 b7724.36 b237.23 de
100.0527.41 b7840.75 ab236.21 de
200.0499.78 c7430.72 b224.12 e
Mean526.94 a7888.43 a232.62 c
Zn-NPs0.0526.13 b5922.03 cde252.52 bc d
50.0510.34 c5488.63 e–h263.94 ab c
100.0310.24 e8078.36 ab243.68 cd e
200.0322.62 d7988.93 ab252.37 bc d
Mean417.33 b6869.49 b253.13 b
Mean0.0541.35 a7239.95 b242.71 a
50.0517.17 b6606.50 c250.59 a
100.0418.83 c7959.55 a239.95 a
200.0411.20 c7709.82 ab238.24 a
Mean472.14 a7378.96 a242.87 a
Mean of
Zn0.0346.71 a6258.15 a212.89 a
50.0336.82 bc6584.39 bc215.05 a
100.0334.27 c6113.17 bc201.03 a
200.0298.69 d6115.12 ab213.39 a
Mean329.12 a6267.71 a210.59 a
Zn-NPs0.0343.09 a6035.46 d212.14 a
50.0299.33 d5564.79 e180.29 b
100.0238.54 e5755.10 cd172.27 b
200.0236.04 e5112.47 bc179.73 b
Mean279.25 b5616.96 b186.11 b
Means of Zn concentration (ppm)0.0344.90 a6146.81 a212.51 a
50.0318.07 b6074.59 b197.67 b
100.0286.41 c5934.13 a186.65 b
200.0267.37 d5613.80 a196.56 c
Differences between means shown with the same lowercase letter in the same column for each application are insignificant.
Table 7. Analysis of variance of effects of different salinity levels, Zn treatments, and Zn levels on H2O2 (mmol kg−1), MDA (nmol g−1 DW), proline (µg g−1 FW), and sucrose (g L−1) contents of quinoa seedling under salinity.
Table 7. Analysis of variance of effects of different salinity levels, Zn treatments, and Zn levels on H2O2 (mmol kg−1), MDA (nmol g−1 DW), proline (µg g−1 FW), and sucrose (g L−1) contents of quinoa seedling under salinity.
Sources of VariationH2O2MDAProlineSucrose
F value (Salinity)415.53 ***164.03 ***255.97 ***37.15 ***
F value (Zn)30.02 ***265.35 ***53.38 ***26.09 ***
F value (concentration)24.55 ***39.91 ***14.95 ***27.12 ***
F value (S × Z)82.48 ***41.08 ***68.11 ***36.69 ***
F value (S × C)8.45 ***11.01 ***9.77 ***9.85 ***
F value (Z × C)33.04 ***11.35 ***30.91 ***66.54 ***
F value (S × Z × C)17.92 ***26.41 ***30.19 ***17.70 ***
CV %5.774.047.296.33
***: significant at p ≤ 0.001.
Table 8. Mean comparison of the effects of different salinity levels, Zn treatments, and Zn levels on H2O2, MDA, sucrose, and proline contents of quinoa seedling under salinity.
Table 8. Mean comparison of the effects of different salinity levels, Zn treatments, and Zn levels on H2O2, MDA, sucrose, and proline contents of quinoa seedling under salinity.
SalinityZn TreatmentsZinc
Concentration (ppm)
(mmol kg−1)
(nmol g−1 DW)
(µg g−1 FW)
(g L−1)
0 mMZn0.010.01 e–h5.27 ef 151.66 ijk33.76 hij
50.08.53 i5.75 e 134.18 k32.64 ijk
100.09.24 hi5.17 ef 144.04 jk45.58 ab
200.010.09 efg5.27 ef 196.67 fg35.14 f–i
Mean9.21 de5.37 e156.64 d 36.78 ab
Zn-NPs0.09.34 gh 5.45 e206.49 f 27.24 lm
50.010.39 ef 6.81 d171.24 g–i 27.00 lm
100.09.70 fgh 7.34 d168.38 hij 29.86 ijk
200.07.41 j 8.19 c142.98 jk 27.42 lm
Mean9.47 e6.95 c 172.27 c27.88 c
Mean0.09.68 f5.36 f 179.07 d 30.50 f
50.09.46 f6.28 e 152.71 e29.82 f
100.09.47 f6.26 e156.21 e 37.72 bc
200.08.75 g 6.73 de169.83 de31.28 ef
Mean9.34 c6.16 c164.45 c32.33 c
100 mMZn0.010.10 efg5.09 ef 280.60 bcd 29.77 jkl
50.09.52 gh5.16 ef 266.01 cde 24.67 m
100.09.21 hi4.51 f 197.10 fg 43.01 bcd
200.09.86 e–h8.78 abc 172.13 ghi48.58 a
Mean9.67 d5.89 d228.96 b36.51 b
Zn-NPs0.011.26 d8.22 c132.17 k 40.64 cde
50.09.89 e–h8.79 abc179.27 fgh 37.78 e–h
100.010.36 ef8.84 abc139.75 k 34.81 ghi
200.010.67 de 8.53 bc139.87 k29.01 kl
Mean10.55 c8.60 a 147.77 d35.56 b
Mean0.010.68 d6.65 de 206.39 c 35.21 cd
50.09.71 f6.98 d 222.64 c31.23 ef
100.09.78 ef6.68 de168.42 de38.91 b
200.010.26 de 8.65 b156.00 e 38.80 b
Mean10.11 b7.24 b188.36 b36.03 b
200 mMZn0.014.12 ab6.90 d 315.03 a 38.61 efg
50.013.36 bc6.84 d 289.37 bc44.49 bc
100.013.75 ab9.55 a 186.76 fgh42.94 bcd
200.014.52 a 9.03 ab 262.14 de 28.63 klm
Mean13.94 a8.08 b263.32 a36.91 ab
Zn-NPs0.013.98 ab8.45 bc203.02 f33.45 ij
50.012.64 c8.69 bc251.40 e 34.72 ghi
100.09.99 e–h8.89 abc297.66 ab 40.53 cd
200.09.92 e–h 8.38 bc256.75 de 38.95 def
Mean11.63 b8.60 a 252.21 a38.67 a
Mean0.014.05 a7.67 c 259.02 ab 36.03 cd
50.013.00 b7.76 c 270.38 a 39.61 ab
100.011.87 c9.22 a242.21 b 41.74 a
200.012.22 c 8.71 b259.44 ab33.79 de
Mean12.79 a8.34 a257.77 a37.79 a
Means of
Zn0.011.41 a5.75 d 249.10 a 32.33 d
50.010.47 c5.92 d 229.85 b30.68 d
100.010.73 bc 6.41 c 175.96 d43.04 a
200.011.49 a7.70 b 210.31 c 40.89 b
Mean11.03 a6.44 b 216.31 a36.73 a
Zn-NPs0.011.53 a7.37 b180.56 d35.50 c
50.010.98 b8.09 a200.64 c 36.43 c
100.010.02 d 8.36 a201.93 c35.87 c
200.09.34 e8.37 a179.87 d 28.35 e
Mean10.46 b8.05 a190.75 b34.04 b
Means of Zn concentration (ppm)0.011.47 a6.56 d214.83 a 33.91 b
50.010.72 b7.01 c215.24 a33.55 b
100.010.38 c 7.39 b 188.95 b39.46 a
200.010.41 c8.03 a 195.09 b34.62 b
Differences between means shown with the same lowercase letter in the same column for each application are insignificant.
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Türkoğlu, A.; Haliloğlu, K.; Ekinci, M.; Turan, M.; Yildirim, E.; Öztürk, H.İ.; Stansluos, A.A.L.; Nadaroğlu, H.; Piekutowska, M.; Niedbała, G. Zinc Oxide Nanoparticles: An Influential Element in Alleviating Salt Stress in Quinoa (Chenopodium quinoa L. Cv Atlas). Agronomy 2024, 14, 1462.

AMA Style

Türkoğlu A, Haliloğlu K, Ekinci M, Turan M, Yildirim E, Öztürk Hİ, Stansluos AAL, Nadaroğlu H, Piekutowska M, Niedbała G. Zinc Oxide Nanoparticles: An Influential Element in Alleviating Salt Stress in Quinoa (Chenopodium quinoa L. Cv Atlas). Agronomy. 2024; 14(7):1462.

Chicago/Turabian Style

Türkoğlu, Aras, Kamil Haliloğlu, Melek Ekinci, Metin Turan, Ertan Yildirim, Halil İbrahim Öztürk, Atom Atanasio Ladu Stansluos, Hayrunnisa Nadaroğlu, Magdalena Piekutowska, and Gniewko Niedbała. 2024. "Zinc Oxide Nanoparticles: An Influential Element in Alleviating Salt Stress in Quinoa (Chenopodium quinoa L. Cv Atlas)" Agronomy 14, no. 7: 1462.

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