Field experiments were performed during the 2018/2019 and 2019/2020 growing seasons from November to June at the experimental farm of the Academy of Agricultural Science (JAAS), Nanjing, Jiangsu, China (35.31°N 113.87 °E). The seeds of two cultivars of spring wheat i.e. Zheng Mai9 (ZM9, salt-sensitive) and Yang Mai25 (YM25, salt-tolerant), were brought from the Seed Center of Nanjing, Jiangsu, China (JAAS).

Both cultivars selected in the current experiment were widely cultivated in the delta of the Yangtz river of China, based on their excellent traits, e.g., ZM9 with more resistance to wheat diseases such as Fusarium head blight and powdery mildew, YM25 with stronger resistance to lodging, and both showing higher yield and better grain quality. The area has a humid subtropical climate, and the mean temperature and humidity of both growing seasons are shown in Supplementary Table S1. The soil's main properties were as follows: 24.9% clay; 55.6% silt; 19.5% sand. The soil chemical properties from 0 to 30 cm of the experimental site were represented in Supplementary Table S2. The plots' size used in this study was 13 m² (3.25 × 4 m), with a distance between each ridge of 45 cm. The experiments were performed using a complete randomized block design (CRBD, factorial), with three biological replications and eight treatments. The experiment field differs in salinity levels. The plots with 4.6 dSm⁻¹ were served as a control, and plots with 14.50 dSm⁻¹ were used as salinity stress. This salinity was designed as high salt stress for wheat plants as it was comparable to saline-alkaline lands in the coastal areas of Jiangsu Province, China (Ju et al., 2021). Salt soil plots were established by mixing normal soil with salt (1:100). In order to confine the lateral diffusion of salt with water movement among plots, a plastic membrane was mulched on the ridges of each plot.

For the priming treatments, two levels of JA (0 and 60 μM) were used. Prior to seed priming, diluted sodium hypochlorite was used as a sanitizer solution of wheat grains for 15 min and then washed several times to remove any debrides of the disinfectant. Then, the grains were divided into two sets, a group soaked in distilled water representing the control, and the second group was soaked in JA with a concentration of 60 μM at 15 °C in darkness for 24 h (Sheteiwy et al., 2021a). The time and the selected dose of JA were preliminarily tested and optimized. The seeds were left to dry, and the sowing was performed in the second week of November during the seasons (2018 and 2019).

The seeds were sown in rows at depth of 4 cm and 20 cm space between rows. The recommended amount of essential fertilizers such as N (388 kg h⁻¹), P (87.75 kg h⁻¹), and K (120 kg h⁻¹) was applied according to the soil fertility status. The entire P and K fertilizers were provided before planting as basal doses; however, the doses of N were completely applied at the stem elongation and node formation stages (after 50 days from sowing). The water requirement was supplied to soils based on the wheat growth stages, in which 15, 25, 30, and 25% were supplied to the soil at the initial stage, vegetative growth, reproductive stage, grain filling stage, respectively of the total water requirements. In comparison, the soil was naturally dried during maturation (5%). Irrigation water was supplied and pumped to the plots from a pond through pipes. The irrigation system has pipes buried in the soil at a specific depth and water is pumped to enter these pipes for irrigation. This system was equipped with a water meter to control the water flow to the plots at the different growth stages. Other agricultural practices for wheat production were similar to those applied by farmers' practices in the Jiangsu region of China.

In order to determine the physiological, morphological, and yield, 20 wheat plants with the same vigor were selected, tagged in each plot, and used for the different analyses. The morphological and physiological parameters were recorded after 60 days after sowing. Plant heights were measured from the base to the tip of the plant shoot. The fresh weight (FW) and dry weight (DW; dried in an oven at 80 °C for 24 h) of treated plants were recorded. After the maturity of plants and spikes, the spike/m², grains/ spike, and weight of 100 grains were estimated and it was determined during only the first growing season (2018/2019).

The leaf physiological traits such as stomatal conductance (g_s), transpiration rate (T_r), net photosynthetic rate (P_n), and intercellular CO₂ (C_i) were determined using a portable photosynthesis system (LI-6400; Lincoln, NE, USA) after 2 h of acclimation in the growth cabinet at 27 °C, 70% relative humidity, the light intensity of 1000 μ mol m⁻² s⁻¹, and CO₂ of 380 μ mol mol⁻¹ (Sheteiwy et al., 2021a). The chlorophyll contents were measured spectrophotometrically using the method described previously by Sheteiwy et al. (2018). Briefly, the fresh leaf samples (0.2 g) were ground in 3 mL ethanol (95%, v/v). The homogenate was centrifuged at 5,000 xg for 10 min and the supernatant was extracted. The mixture was then determined by monitoring the absorbance at the wavelengths 665, 649, and 470 nm using a spectrophotometer (UV-2101/3101 PC; Shimadzu Corporation, Analytical Instruments Division, Kyoto, Japan). The chlorophyll fluorescence (CF) was measured by the chlorophyll fluorometer (IMAG-MAXI, Heinz Walz, Effeltrich, Germany) in the expanded leaves after dark adaptation for 15 min according to the methodology of Ali et al. (2015).

To deeply observe the changes in the cellular structures of the plant cell and major structural organelles in terms of chloroplast, cell wall, and starch grain and their functions in response to salt stress and JA priming, the leaf ultrastructural changes were investigated using a protocol reported by Yang et al. (2021) with slight modifications. Segments (6–8 per treatment) were randomly selected from wheat plants and overnight immersed in glutaraldehyde (2.5%, v/v) and washed four times with.1 M of phosphate-buffered saline (PBS), with 10-min intervals between each washing. Then, at 15–20 min intervals, the samples were dehydrated in a graded series of ethanol (50%, 60%, 70%, 80%, 90%, 95%, and 100%) and then washed with absolute acetone for 20 min. The samples were then assembled and fixed on copper grids to be scanned using the transmission electron microscope (JEOL TEM-1230 EX), as described by Yang et al. (2021).

The Na⁺ and K⁺ contents of the blade, sheath, stem, and roots of both wheat cultivars were recorded using the method described by Zhao et al. (2014). The tissues were washed with distilled water and then dried at 50 °C for 4 d. After that, the dried samples were ground into a fine powder by liquid nitrogen and digested in nitric acid (5 mL) overnight. Then, Na⁺ and K⁺ contents of different tissues were monitored using a flame photometer following the methodology of Zhao et al. (2014).

The water relation parameters such as water potential and osmotic potential were measured following the protocol reported by Ahmed et al. (2013). Briefly, the segments of the fully expanded leafy sample were macerated in a mortar. After extract filtration, the sap was centrifuged at 10,000 x g for 10 min at 4 °C, then the supernatant was used to estimate osmolality (c) using a vapor pressure osmometer (Wescor Inc., Logan, UT, USA). WUE was measured as the ratio between P_n and T_r as described by Jones (2013). Briefly, fresh eight of the leaf sample, and the dry weight (for 48 h in an oven at 75 °C) were recorded and used for the RWC determination. The leaf was then soaked for 24 h in distilled H₂O. The leaves were then dried using tissue paper prior to turgid weight calculation and the RWC was measured. The RWC was quantified in tissues using the equations of Barr and Weatherley (1962) as follows:

The hydrogen peroxide (H₂O₂) content was measured in the root and shoot according to Halliwell et al. (1987). Briefly, the root and shoot samples (0.5 g) were macerated in trichloroacetic acid (5.0 mL of 0.1%), and the supernatant was mixed with 1 mL of 1 mM KI, and the absorbance at the wavelengths 390 nm by using spectrophotometer was monitored. Proline was determined in the root and shoot samples according to Sheteiwy et al. (2021b). The leaf samples (100 mg) were homogenized in 3% sulfosalicylic acid (5 mL) and centrifuged at 5,000 xg for 10 min. The supernatant was mixed with acid-ninhydrin and acetic acid and boiled for 1 h at 100 °C followed by incubation in an ice bath. The absorbance of chromophore-containing toluene was monitored at 520 nm. Tocopherol in root and shoot samples was recorded at 520 nm following Backer et al. (1980). The flavonoid content in root and shoot samples was determined according to Zhishen et al. (1999). Total phenolic content in root and shoot were quantified and calculated using gallic acid (Sigma-Aldrich Chemie GmbH, Taufkirchen, Germany) as a standard (Shohag et al., 2012).

The antioxidant enzyme activities, such as APX, SOD, and Phenylalanine ammonia-lyase (PAL) were determined in the leaf and root tissues spectrophotometrically at 290 nm by the method of Sheteiwy et al. (2021a). Briefly, fresh leaf and root samples (0.5 g) were homogenized in liquid nitrogen and then macerated in K-phosphate buffer (8 mL, 50 mmol L⁻¹, pH 7.8) in ice-cold conditions. The extract was then centrifuged at 10,000 x g for 20 min at 4 °C where the supernatant was utilized for the aforementioned enzyme activity measurements.

APX activity was determined following the methodology of Nakano and Asada (1981). The reaction mixture was composed of 1.4 mL PBS (25 mmol L⁻¹ pH 7.0 + 2 mmol L⁻¹ ethylene diamine tetra acetic acid (EDTA), 300 mmol L⁻¹ H₂O₂, 7.5 mmol L⁻¹ ascorbic acid 100 μL, and 100 μL enzyme extract in a total volume of 1.7 mL. APX enzyme was measured by noticing the reduction percentage of the absorbance of ascorbic acid at 290 nm for 2 min. SOD activity was measured according to the method of Giannopolitis and Ries (1977). The reaction mixture contained 3 mL 50 mmol L⁻¹ phosphate buffer (pH 7.8), 13 mmol L⁻¹ methionine, 1.22 mmol L⁻¹ riboflavin, 78.2 μmol L⁻¹ EDTA, 56 μmol L⁻¹ of Nitro blue tetrazolium (NBT) and 100 μL extract. The mixture was illuminated and the reaction was used to measure the inhibition of the photochemical reduction of NBT at 560 nm. One unit of SOD was defined as the amount required to inhibit the photoreduction of NBT by 50%. SOD activity was expressed as mg⁻¹ protein as reported earlier by Zhu et al. (2015).

The concentrations of cytokinins (CKs) were determined according to the method of Rauf and Sadaqat (2007). The indole acetic acid (IAA) was measured following the methodology of Sheteiwy et al. (2021c). Abscisic acid (ABA) was quantified in shoot and root tissues following the methodology of Gao et al. (2020a). For JA determination, frozen root and shoot samples (200 mg) were ground into fine powder using liquid nitrogen. The extracts were then centrifuged at 12000 x g for 10 min at 4 °C, and then JA was quantified using liquid chromatography-mass spectrometry (Waters, Milford, Massachusetts, USA) as described by Tsukahara et al. (2015).

The transcript levels of Na⁺ uptake and transport genes such as SOS1-NHX2- HVP1 were analyzed in leaf blade-, leaf sheath, stem, and root tissues. The total RNA isolation was performed using an RNA isolation (Takara Bio Inc., Shiga, Japan). cDNA was synthesized using Primer Script RT reagent Kit (Takara Bio Inc.) from 1 μ g of total RNA in a 20 μ L reaction, and diluted 4-fold with water. SYBR premix EX Taq (Takara Bio Inc.) was used to perform the quantitative real-time (qRT-PCR) analysis. The frozen tissues (100 mg) were homogenized in liquid nitrogen and qRT-PCR was performed following the methodology of Gao et al. (2020b) using primers used previously by Darko et al. (2017). ACT1 was used as a control to determine the transcription level of the other studied genes. The PCR program was the same as followed by Salah et al. (2015).

Two-way analysis of variance (ANOVA) was applied to determine differences among treatments. The results were described as the means of three replicates ± SD. The data were analyzed using the statistical package (IBM-SPSS, 19, USA). Mean values were compared by applying Duncan's multiple range test at the.05 level of significance. Asterisks indicate significant differences: ^*P < 0.05, ^**P < 0.001, ^***P < 0.0001. The Hierarchical Cluster Analysis (HCA) and Principle Component Analysis (PCA) were performed through the SPSS program on log₁₀ of the obtained data.

Results revealed that salt stress caused a significant (P ≤ 0.05) reduction in growth and numbers of tillers in wheat seedlings compared to the control seedlings (Table 1). The reduction of shoot length, FW, and DW in response to salinity reached 23, 40, and 44% in the ZM9 cultivar and 22, 45, and 44% in cultivar YM25, respectively, compared with the control. Priming with JA resulted in improved plant height, FW, and DW as well as tiller numbers in both wheat cultivars during both growing seasons, compared to unprimed plants (Table 1).

Comparing both cultivars, the highest DW and FW values were observed in the YM25 cultivar primed with JA in both seasons. On the other hand, JA-primed plants of YM25 in 2019 and ZM9 in 2020 attained taller plants than the corresponding non-primed plants. We noticed that both cultivars showed a similar pattern of growth upon the treatment with 60 μM JA (Table 1). Plant height, FW, DW, and the numbers of tillers were significantly (P ≤ 0.05) enhanced by JA priming under salt stress compared to the salt-stressed plants of both cultivars during the two growing seasons.

As presented in Table 1, and compared with the control, P_n and T_r were enhanced in the salt-stressed primed with JA compared to salt-stressed non-primed plants, which had reductions in both traits. The primed plants of the cultivar ZM9 showed the highest value of Tr in 2019, while the cultivar YM25 produced the highest value of Tr in 2020 growing season (Table 1). A significant decrease was observed in C_i, g_s, chlorophyll a (Chl a), Chl b, total chlorophyll, and Chlorophyll fluorescence (FC) in the salt-stressed plants compared to the control plants (Table 2). Interestingly, C_i, g_s, Chl a, Chl b, carotenoids, and CF [represented by maximal PSII photochemical efficiency (Fv/Fm)] were increased in the JA-primed control compared to the non-primed seedlings (Table 2). This could potentially mitigate the damaging effect of salinity stress in both wheat cultivars; yet, still lower than the control plants in both growing seasons.

In comparison to the control plants (no salinity stress), there were considerable increases in Na⁺ concentrations in different tissues of wheat (blade, sheath, stem, and roots) of the two wheat cultivars in both growing seasons in response to salt stress (Table 3). These cultivars showed the highest accumulation of Na⁺ content in the sheath but the lowest in stems. In salt-stressed plants, Na⁺ accumulation was higher in the blades, followed by sheath, roots, and finally, stem in both cultivars during the two experimental years. On average, the increments of Na⁺ content in blade, sheath, root, and stem, were 184.0, 268.0, 227, and 201% in ZM9 and 172, 172, 96, and 133% in YM25, respectively, compared with their corresponding controls.

Interestingly, JA priming significantly (P ≤ 0.05) restricted the uptake of Na⁺, which resulted in reductions of Na⁺ in the tissues of the tested wheat cultivars during the two growing seasons (Table 3). In contrast, K⁺ content was highly retarded by salinity, mainly in blades (Table 3). Thus, the reduction of K⁺ reached 49, 40, 39, and 45% in the blade, sheath, stem, and root tissues, respectively, in the ZM9 cultivar salt-stressed plants compared to control plants of the same genotype. In the YM25 cultivar, K⁺ was reduced by 50, 30, 41, and 47% in the blade, sheath, root, and stem tissues, respectively, compared to the non-stressed control plants. JA retarded the reduction of K⁺ in different organs partially for more affected organs (blade and sheath) and enhanced its content to be higher than the control plants, especially for root (Table 3).

Plants exposed to salinity stress exhibited significant (P ≤ 0.05) reductions in the water and osmotic potential of the studied wheat cultivars during the two growing seasons compared with that in non-salinized control plants (Figures 1A,B). On the other hand, JA priming significantly (P ≤ 0.05) increased the water and osmotic potential under saline or non-saline conditions relative to their control. Salinization of soil drastically decreased the RWC of both wheat cultivars comparably throughout the two studied years (Figure 1C). Although WUE was enhanced by salinity, priming in JA profoundly alleviated the damaging impact of salinity on RWC to be statistically highly significant relative to control. These results suggest that JA plays a vital role in alleviating the damage effects of salinity in wheat cultivars through regulating water status and the osmoregulation process.

The H₂O₂ significantly (P ≤ 0.05) increased in response to salinity in the shoots and roots of both cultivars (Figures 2A,B). JA treatment lowered the content of H₂O₂ in roots of salt-stressed plants effectively to be comparable to the control plants. In contrast, the effect of JA on the reduction of shoots' H₂O₂ was comparable to saline plants which were generally higher than the control (Figure 2A). The specific activities of antioxidant enzymes viz., SOD (Figures 2C,D), and APX (Figures 2E,F) of both wheat cultivars increased by salinity stress in the different tissues in both wheat cultivars. JA profoundly exacerbated the activities of the enzymes in the root and shoot higher than their control.

In non-salinized and salinized plants, JA priming notably enhanced SOD and APX activities compared to the non-JA treated plants in root and shoot tissues of both cultivars (Figures 2E,F). The stimulation power of JA on SOD and APX activities was higher in the salinized plants than in non-salinized ones. PAL activity was also enhanced in response to salinity stress reaching ≥ 6-fold in shoots or roots of the two wheat cultivars throughout the two studied seasons. Further increment of PAL activity was denoted for saline and non-saline plants in plants receiving JA compared to their corresponding control (Figures 2G,H).

The contents of proline (an important cell osmolyte) and tocopherols (a non-enzymatic antioxidant) were progressively boosted by salinity stress in the shoots and roots of the two cultivars (Figures 3A–H). The study also included the effect of salinity stress and priming with JA on secondary metabolites of wheat plants. In this regard, flavonoids (Figures 3A,B) and total phenols (Figures 3C,D) accumulated in response to salinity stress in the shoots and roots of both wheat cultivars. JA application had a regulatory role on higher biosynthesis of both metabolites compared to their corresponding control; the highest content was denoted for the non-salinized plants primed with JA. Likely, the antioxidant molecule, tocopherol, was stimulated by soil salinization, and higher exacerbation of tocopherol biosynthesis was also denoted by JA application (Figures 3E,F). Further accumulation of proline could be attributed to the priming of wheat plants with JA. For example, the salinized plants primed with JA demonstrated the highest accumulation capacity of proline in tissues (Figures 3G,H).

The application of JA to wheat seeds caused significant positive changes in hormonal homeostasis (Figure 4). In this regard, the contents of CKs were dramatically (P ≤ 0.05) reduced (by 31.9–66.7%) in salt-stressed plants, especially in the shoots, for both cultivars during both growing seasons (Figures 4A,B). However, JA increased the content of CKs in shoots and roots of non-stressed plants compared to the control plants. The JA not only retrieved the contents of CKs in stressed plants higher than their corresponding stressed plants in those not treated with JA but also increased it to higher levels than in the non-stressed plants (Figures 4A,B). Moreover, IAA was highly attenuated in salt-stressed plants with approximately the same effect on both tissues as well as the two tested cultivars during the two seasons with a percent reduction of ~ 23% compared with non-salinized plants (Figures 4C,D).

JA curtailed the reduction of IAA contents in shoots and roots to be higher than in control plants as well as stimulated IAA contents of non-stressed plants, which is recommended for both cultivars, tissues, and studied growing seasons (Figures 4C,D). Except for the stressed shoots of YM25, endogenous JA contents in shoots and roots were not affected by salinity stress shoots and roots (Figures 4E,F). Furthermore, the exogenous application of JA significantly (P ≤ 0.05) boosted the JA endogenous content in stressed and non-stressed plants to higher levels than the control treatment (non-primed seeds). It is worth mentioning that the highest endogenous JA contents were noted in JA-primed plants exposed to the high salt concentration.

ABA was differentially affected by salinity stress; the effect depended on the examined tissue and growing season (Figures 4G,H). The shoots of the two cultivars attained significantly (P ≤ 0.05) higher levels of ABA under salinity stress regardless of the tested season. However, in the roots, salinity stress reduced ABA contents in 2019 but increased it in 2020 for both cultivars. JA increased ABA contents of shoot tissues in non-stressed plants compared to the corresponding treatment. The levels of ABA in shoots of salinized plants treated with JA were lower than in stressed plants but still higher than in non-stressed plants. ABA contents were significantly (P ≤ 0.05) enhanced in the roots of the non-stressed plant primed with JA in the two growing seasons (Figure 4H). On the other hand, JA treatment restrained the reduction of ABA levels in roots in 2019. During the growing season of 2020, the increased levels of ABA were highly (P ≤ 0.05) attenuated by salinity stress in response to JA priming, yet the levels were higher than in control plants (Figures 4G,H).

Salinity stress differentially induced the expression levels of SOS1 based on the organ tested (Figure 5). Upon exposure to high salt, the transcript levels of SOS1 were significantly (P ≤ 0.05) increased by 2-, 1.5-, 3.1-, and 4.2-fold in roots, stem, leaf sheaths, and blades, respectively compared to the same tissues of non-stressed plants (Figures 5A,D,G,J). Salinity stress induced the expression level of NHX2 progressively by 5.8-, 3.3-, 3-, and 3-fold and HVP1 by 1.6-, 1.92, 1.84-, and 1.92-fold for roots, stems, and leaf sheaths, and blades, respectively compared to non-stressed plants. Priming of JA exhibited stimulatory effects on the expression of SOS1, NHX2, and HVP1 genes in wheat plants under saline and non-saline conditions (Figure 5).

Salinity dramatically affected the ultra-structural configurations of the cell, where the disintegration of plastid compactness in salinized wheat cultivars was the resultant (Figure 6). Due to salinity stress, starch grains and plastoglobuli particles were reduced alongside the loss of the integrity of grana. Interestingly, the cultivar ZM9 exhibited swollen chloroplasts with non-well-developed and sometimes, unrecognizable granum structures compared to its corresponding control or the other wheat cultivar. By combining JA priming and increased NaCl application on plants, the chloroplasts were oval shaped, more consistent with the regular configuration. In addition, several numbers of starch grains and higher plastoglobuli particles relative to deteriorated chloroplasts' of saline treatments were only found in the JA-treated plants (Figure 6).

The number of spike/m² and grains/spikes were dramatically (P ≤ 0.05) reduced by 25 and 21%, respectively, in response to salinity stress compared with their relative control (Figures 6A,B). The weight of 100 grains was slightly affected by salinity stress with an 8% reduction in the cultivar ZM9 (Figure 6C). On the other hand, the same stress in the YM25 cultivar did not affect the weight. Intriguingly, JA significantly (P ≤ 0.05) increased the yield of non-stressed plants. Salinized-wheat plants treated with JA highly mitigated the detrimental effects of salinity on wheat yield as compared with the unprimed stressed plants.

All mean values of the morphological and biochemical parameters were subjected to hierarchical clustering as a heat map of five 5 clusters: Group (A) included ABA and H₂O₂ (in shoot and root), and Na⁺ content (in the root, stem, blade, and sheath); Group (B) included IAA and CKs (in shoot and root), and K⁺ (root, shoot, blade, and sheath); Group (C) included JA, PAL and proline (in shoot and root), and SOD activity (in roots); Group (D) included the expression of genes HVP1, NHX2, and SOS1 (in the stem, root, and leaf sheath and blade) and SOD activity (in the shoot); while Group (E) included APX activity, tocopherol, flavonoids and total phenol (in shoot and root) (Figure 7).

PCA analysis presented in Figure 8 indicated that the variables of group A strongly connected with stressed plants without JA priming, especially ABA. The group–B variables were relatively strongly delineated with JA-primed non-stressed plants (Figure 8) especially Ci, IAA, and cytokinin. On the other hand, the group –C, and –D variables were relatively strongly delineated with JA-primed salinity stressed plants especially proline and SOD (shoot and root).