Introduction

Abiotic stresses, such as drought, heat, and salinity, reduce crop yield by more than 50%. Soil salinity is a worldwide problem due to global warming and climate change (Singh 2022, Wang, Han et al. 2022). Approximately 50% of the arable land is estimated to be affected by salinity by the year 2050 (Metwali et al. 2015; Shrivastava and Kumar 2015), which will likely impact the global food production. In Egypt, salinity affects 60%, 25%, and 20% of the cultivated lands in the Lower, Middle, and Upper Delta. Salinity has been reported to affect plant growth, yield, membrane integrity, chlorophylls, carotenoids and photosynthetic activity, osmotic adjustments, and water relations (Benjamin and Nielsen 2006; Akrami and Arzani 2018; Maswada et al. 2018). Saline soil drastically reduces crop yield by inhibiting seed germination, seedling growth, flowering, and fruit set (Sairam and Tyagi 2004). Physiological processes like respiration, photosynthesis, nitrogen fixation, and other metabolic processes are affected by salinity, resulting in stunted growth and decreased productivity (Acosta-Motos et al. 2017; Pal et al. 2021). Moreover, it disrupts the cellular osmotic balance and increases oxidative stress by generating reactive oxygen species (ROS), which can damage cellular components, such as proteins, lipids, and DNA, undermining vital cellular functions (Fahramand et al. 2014; El-Sheshtawy et al. 2022).

Plants have developed a range of sophisticated regulatory strategies, both enzymatic and non-enzymatic, to protect themselves from ROS (Fang and Xiong 2015). Endophytic plant growth-promoting bacteria (PGPB) offer the potential to reduce the impact of salt stress in plants through the accumulation of osmolytes and antioxidant compounds, in addition to their direct and indirect stimulatory effect on plant growth and productivity. These compounds are associated with osmotic adjustment and stabilize cell components and work as free radical scavengers (Kumar et al. 2020). The use of PGPB aligns well with the modern demands of agricultural, economic, social, and environmental sustainability (Chaparro et al. 2012; Ha-Tran et al. 2021). Several PGPB proved the ability to alleviate the effect of salinity in cumin and wheat crops (Ramadoss et al. 2013; Moradi and Piri 2018). Other species of PGPBs can be used to control rice phytopathogens responsible for grain loss (Ngalimat et al. 2021). Burkholderia seminalis enhances the growth of Arabidopsis sp, pak choi, Chinese amaranth, lettuce, and other vegetables (Hwang et al. 2021). Enterobacter cloacae enhanced the growth and productivity of wheat and soybean (Ramesh et al. 2014). Both Bacillus amyloliquefaciens and E. cloacae are considered as potential PGPBs as well as bio-control agents (Chen et al. 2007a, b; Mohamed et al. 2020; Ahmed et al. 2022).

Soybean (Glycine max L. (Merr.) is a subtropical member of the Leguminosae (Fabaceae) family, an erect bushy annual crop with considerable morphological diversity. Soybean is a crucial strategic crop because of its variety of uses as food, feed, and raw material for industry. (Khojely et al. 2018). Besides being one of the most economically significant oilseeds and biodiesel crops today, soybean also provides a significant amount of protein and oil for both human and animal sustenance (Zhang et al. 2022). The ability of Curtobacterium sp. SAK1 to produce phytohormones, antioxidants, and aminocyclopropane-1-carboxylic acid deaminase alleviates the salinity stress on soybean (Khan et al. 2019). Similarly, both Pseudomonas pseudoalcaligenes (SRM-16) and Bacillus subtilis (SRM-3) could mitigate the effect of salt on hydroponically grown soybean plants (Yasmin, Naeem et al. 2020).

Considering that salinity threatens agricultural production in general and soybean in particular which is considered a glycophyte, the aim of this study was to find compatible PGPBs with the ability to support the growth of this plant under salt stress. Additionally, the study aimed to investigate the response of antioxidant systems, the accumulation of osmolytes, and membrane integrity under this condition to understand the mechanism of response to the selected bacteria.

Materials and Methods

Proline, trichloroacetic acid, and citric acid were all supplied by Sigma chemical company. Other chemicals were supplied by different local companies (mostly,Al Gomhoria company) and they all were of analytical grade.

Plant Material and Bacterial Strains

A pure and homogenous line of soybean seeds (Giza 111) was obtained from the Agriculture Research Center, Ministry of Agriculture, Egypt. Three bacterial strains were used in this experiment, two of them were previously isolated for their potential as plant growth promoters and they were molecularly identified as Bacillus MAP3 (accession number MG214652) and Enterobacter Delta PSK (accession number MT012829) (Agha, Abbas et al. 2021, M. Mowafy, S. Agha et al. 2022). Bradyrhizobium japonicum (accession number EMCC No.1112) was obtained from the Microbial Research Center "Cairo Mircen", Ain Shams University, Cairo, Egypt. Additionally, the Al-Aqdin inoculum (a commercial product containing Bradyrhizobium japonicum and other PGPB specific for soybean designed and recommended by the Agriculture Research Center, Ministry of Agriculture, Egypt) was used as a positive control in this experiment.

Experimental Design

The soybean seeds were sterilized with 0.01% HgCl2 solution for 3 min, washed with sterilized distilled water, and then divided into five groups to have the following treatments: control (water treatment), B. japonicum, B. japonicum + Bacillus MAP3, B. japonicum + Enterobacter Delta PSK, and Al-Aqdin inoculum. The bacterial cultures were obtained from fresh LB (Luria–Bertani) media incubated at 28 °C with 150 rpm for 2 days and the concentration was adjusted to 108 CFU ml−1. For mixed treatments, equal volumes were used from the cultures. The soybean seeds were pre-inoculated with the aforementioned bacterial treatment for 1 h at room temperature before sowing. The seeds soaked in sterile distilled water served as the control. Five seeds were cultivated per pot (18 cm depth and 25 cm diameter) with six Kg of soil/each. There were 15 pots for each treatment. The soil (clay: sand, 2:1 wt/wt) was mixed well before use and the clay soil was obtained from the field close to Mansoura University. The experiment was conducted in the greenhouse of the Botany Department, Faculty of Science, Mansoura University, Egypt. The temperature, relative humidity, and light intensity levels were 25–30 °C, 57–60%, and 100–200 μmol m−2 s−1, respectively. The soil physical and chemical properties were analyzed, as shown in Table 1. After 21 days of plant growth, each of the described five treatments was subdivided into three groups to start salt treatments: control (tap water), 50 mM NaCl, and 100 mM NaCl. Plant materials were collected for analysis at two points before salinity and after it by 19 days (at 21 and 40 days old ages). Ten plants were randomly uprooted to assess shoot and root lengths, and dry weights (g/plant) were measured after drying the fresh tissues at 70 °C for two days. The dry tissues were grounded into a fine powder using a homogenizer and stored in sealed glasses at room temperature for various analytical experiments. The water content (WC) of shoot and root was calculated from Eq. 1 (Al Hassan, Fuertes et al. 2015).

$$ {\text{WC = }}\left[ {\left( {{\text{FW {-} DW}}} \right){\text{/FW}}} \right]{\text{ x 100}} $$
(1)

where FW refers to fresh weight and DW refers to dry weight.

Table 1 Physical and Chemical properties of soil used for cultivation of Soybean in the greenhouse pot experiment
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Germination percentage (GP) (Vibhuti et al. 2015) and Germination rate (GR) (Vashisth and Nagarajan 2010) were determined according to Eqs. 2 and 3, respectively.

$$ {\text{GP = }}\left( {{\text{No}}{\text{. of germinated seeds/total cultivated seeds}}} \right){*100} $$
(2)
$$ {\text{GR = }}\left( {\text{a/1}} \right){ + }\left( {\text{b - a/2}} \right){ + }\left( {\text{c - b/3}} \right){ + }.....{ + }\left( {{\text{n - }}\left( {\text{n - 1}} \right){\text{/N}}} \right) $$
(3)

where a, b, c, …,n are numbers of germinated seeds after 1, 2, 3, …, N days from the start of imbibition.

Chlorophyll and Carotenoid Contents Measurement

The chlorophyll and carotenoid contents of the soybean leaves were determined by following the procedure described by Hiscox and Israelstam (1979). The amount of 0.1 g (FW) leaves obtained from the first leaf was extracted with 7 ml of dimethyl sulfoxide at room temperature for 24 h. The extracted solution was filtered and the absorbance was measured at 470, 644, and 662 nm by Jenway 7315 UV–VIS. Chlorophyll a and chlorophyll b amounts were calculated using the following equations (Arnon, 1949; Ibrahim et al., 2014).

$$ {\text{Chlorophyll a }} = { 12}.{7 } \times \, \left( {{\text{OD}}_{{{662}}} } \right) \, {-}{ 2}.{69 } \times \, \left( {{\text{OD}}_{{{644}}} } \right) $$
(4)
$$ {\text{Chlorophyll b }} = { 22}.{9 } \times \, \left( {{\text{OD}}_{{{644}}} } \right) \, {-}{ 2}.{69 } \times \, \left( {{\text{OD}}_{{{662}}} } \right) $$
(5)

The carotenoid content was calculated according to the following equation (Villanueva et al. 1985).

$$ {\text{Carotenoids}} = \, \left[ {{\text{OD}}_{{{47}0}} {-}{ 1}.{28 }\left( {{\text{Chl}}.{\text{a}}} \right) \, + { 5}.{67 }\left( {{\text{Chl}}.{\text{ b}}} \right)} \right] \, / \, \left( {{256 } \times 0.{9}0{6}} \right) $$
(6)

Membrane Features

Electrolyte leakage (EL) was determined in plant leaf tissue (one cm2 piece away from the midrib from the second leaf) which was placed in test tubes containing 10 ml distilled water and the electrical conductivity (EC1) was recorded using EC meter. The tubes were placed on a shaker for 2 h and recorded the EC2. Then the tube was autoclaved at 120˚C and after cooling, EC3 was recorded. Equation 7 was used to estimate electrolyte leakage on the measured values (Lutts et al. 1996).

$$ {\text{Electrolyte leakage }}\left( {\text{\% }} \right) = { }\frac{{{\text{EC}}2 - {\text{ EC}}1}}{{{\text{EC}}3}}x{ }100 $$
(7)

Membrane Lipid peroxidation was determined by estimating malondialdehyde (MDA) formation as described by Heath and Packer (1968). One g of the fresh leaf sample (the second leaf) was macerated in 5 ml 0.1% trichloroacetic acid (TCA) and then centrifuged at 10 000 xg for 15 min. Then, 2 ml of 20% TCA and 2 ml of 0.5% thiobarbituric acid were added to 1 ml of the supernatant and the mixture was incubated at 95 °C for 30 min. The absorbance was measured spectrophotometrically at 532 and 600 nm (Jenway 7315 UV–VIS, Burlington, VT, USA). The concentration of MDA was calculated using MDA molar extinction coefficient (ε = 155 mM−1 cm−1).

Measurement of H2O2

The method of Velikova et al. (2000) was used to measure H2O2. First, 0.1 g of 2nd plant leaf was homogenized in 0.1% trichloroacetic acid (TCA). After centrifugation, 0.5 ml of the supernatant was mixed with 0.5 ml of 100 mM potassium phosphate buffer (pH 7.0) and 2 ml KI (1 M). The reaction was left to develop in dark for one hour, and the absorbance was measured at 390 nm (Jenway 7315 UV–VIS, Burlington, VT, USA). The standard curve was used to calculate H2O2 concentrations.

Assessment of Osmotic Regulators

Plant dry tissue powder was obtained by heating the fresh tissue at 70 °C for 48 h till getting constant dry weight. Osmolytes were determined in plant-water extracts prepared by incubating 0.1 g of dry tissue powder in 25 ml of distilled water at 90 °C for 60 min, followed by centrifugation. The EC of the plant-water extracts was measured to express osmotic pressure directly (Mickky et al. 2019).

In addition, citric acid was estimated as described previously (Snell and Snell 1937). To 5 ml of the extract, 15 ml of the deproteinizing solution (3 g HgCl2 and 3 g ZnSO4 in 100 ml distilled water) was added. After centrifugation, 4 ml of 10 N HCl and 1 ml of 6.2% FeCl3 were added and the absorbance was measured at 445 nm. Standard citric acid dilutions (1.0 and 100 µg) were used to construct the standard curve.

The proline content of dry leaves was estimated by the method of Bates (1973). About 0.5 g of each sample was added to sulfosalicylic acid with an equivalent amount of glacial acetic acid and ninhydrin. The samples were heated at 100 °C for 2 h. After cooling, 5 ml of toluene was added to extract the developed color. The absorbance of the toluene layer was measured at 520 nm. Standard proline dilutions were used to construct the standard curve.

Total soluble sugars (TSS) were extracted by submerging 0.2 g of the dry shoot powdered tissue overnight in 10 ml 80% ethanol at 25 °C with periodic shaking. After filtration, an aliquot of 0.1 ml was added to 3 ml of freshly prepared anthrone reagent and incubated in a boiling water bath for 10 min and the absorbance was measured at 625 nm after cooling. The amounts of TSS in plant extracts were estimated using the standard curve of glucose (Hansen and Møller 1975).

Determination of Antioxidant Capacity

The methanolic extract of the dried plant samples was prepared as described by Kosem et al. (2007). An equal volume of the freshly prepared methanolic solution of DPPH (0.1% w/v) and sample extracts were mixed and kept for 30 min at room temperature in dark. Similarly, equal proportions of methanol and the DPPH solution were mixed to take the absorbance of the control. The absorbance was measured at 517 nm by using a spectrophotometer (Jenway 7315 UV–VIS, Burlington, VT, USA). The DPPH radical scavenging activity was determined using Eq. 8 (Şengül et al. 2014).

$$ {\text{DPPH radical scavenging activity }}\% \, = \, \left[ {{1} - \, \left( {{\text{A}}/{\text{B}}} \right)} \right] * {1}00 $$
(8)

where A is the absorbance of the sample and DPPH mixture and B is the absorbance of the methanol and DPPH mixture.

The reducing power was determined in the methanolic extract by mixing one ml with 2.5 ml of 0.2 M potassium phosphate buffer pH 6.6 in addition to 2.5 ml potassium ferricyanide (10 g/L). The mixture was further incubated at 50 °C for 20 min. and 2.5 ml trichloroacetic acid (100 g/L) was added. After centrifugation at 3000 rpm for 10 min, 2.5 of the supernatant was mixed with 2.5 ml dist. Water and 0.5 ml FeCl3 (1 g/L) then the absorbance was measured at 700 nm (Yildirim et al. 2001).

Assays of Antioxidant Enzymes

The activity of antioxidant enzymes was determined in the extracts prepared by homogenization of 2 g of liquid nitrogen frozen leaf tissue in 20 ml of 100 mM potassium phosphate buffer pH 6.8. After centrifugation, the following enzymes were assayed as described before (Oktay et al. 1995; Abd-ElGawad et al. 2020).

Catalase Activity

(CAT; EC 1.11.1.6) was assayed in 500 µl assay mixture containing 50 mM potassium phosphate buffer pH 7 and 10 mM H2O2. The reaction started by adding 20 µl protein extract and the absorbance was monitored for 2 min at 240 nm using the kinetic mode of Jenway 7315 UV–VIS spectrophotometer. One unit of enzyme activity is the decomposition of 1 µM H2O2/min/ml.

Peroxidase Activity

(POX; EC 1.11.1.7). was assayed in a 500 µl reaction mixture containing 50 mM potassium phosphate buffer pH 7, 50 mM pyrogallol, and 0.03% H2O2. The increase in absorbance was monitored at 420 nm after starting the reaction with 20 µl protein extract.

Polyphenol Oxidase PPO

(EC 1.14.18.1.) assay was committed in 1 ml reaction mixture containing 50 mM potassium phosphate buffer pH 7 and 50 mM pyrogallol. The measurement started by adding 50 µl protein extract and absorbance was monitored at 420 nm.

Estimation of Secondary Metabolites

The methanolic extract used for the estimation of total phenols and flavonoids was prepared by incubating 0.1 g plant dry powder in 10 ml methanol (80%) for one week at room temperature. After centrifugation, the supernatant was kept at 4 °C for analysis. Total phenolic content was estimated using Folin–Ciocalteu reagent, as described earlier (Dhungana et al. 2015; Zeitoun et al. 2017). A volume of 50 µL extract was mixed with 1000 µl 2% Na2CO3. After 3 min, 50 µL of 1 N Folin-Ciocalteu reagent was added and the mixture was left at room temperature for 30 min. The absorbance was measured at 750 nm. The total polyphenol content was expressed in microgram gallic acid equivalents per gram dry weight of the sample (µg GAE/g). The total flavonoid content was measured by adding 30 µl of 5% NaNO2 to 300 µL of the methanolic extract then 60 µL of AlCl3 (10%) was added to the mixture followed by 200 µL of 1 M NaOH and the absorbance was measured at 500 nm. The total flavonoid content was measured and expressed in microgram quercetin equivalents (QE) per gram dry weight of the sample (µg QE/g) (De Souza et al. 2018).

Statistical Analysis

Measurements were gathered from 10 samples of the biological replicates (physical growth parameters) in a completely randomized method. Three samples were analyzed for the technical replicates (pigment content, membrane features along with ROS, osmolytes, antioxidant capacity, antioxidant enzymes, phenols, and flavonoids). The statistical analysis was conducted using COSTAT. With a 95% confidence level, Fisher's One- and Two-way analyses of variance (ANOVA) were used for the measurements made on samples from the first (before salinity) and second stages (after salinity application) respectively. The Levene test (normality), a parametric distribution was used. Using COSTAT software, Fisher's test was performed with a probability level of P < 0.05.

Results

Bio-priming with the used PGPBs significantly affected the germination parameters of soybean. Compared with the control, the highest values of GP and GR were recorded for B. japonicum + Delta PSK-treated seeds. Significant differences were observed in the growth parameters of soybean in response to the used bacterial treatments after 21 days (Table 2). Shoot length increased by 6.3%, 31.13%, 49.58%, and 37.2%, while the dry weight increased by 8%, 20%, 40%, and 20% in response to B. japonicum, B. japonicum + MAP3, B. japonicum + Delta PSK, and Al-Aqdin inoculum, respectively. Leaf area, root length, and dry weight significantly increased by 78.58%, 20.19%, and 93.54% after B. japonicum + Delta PSK treatment, which also induced a remarkable increase in water contents of both shoot and root. Compared with the control, the highest increases in the contents of Chl. a, Chl. b, Chl. a/Chl. b, and carotenoids were obtained in response to B. japonicum + Delta PSK (57.35%, 46.26%, 7.33%, and 49.19%, respectively) (Table 3).

Table 2 Effect of different bacterial treatments on germination percentage and rate, shoot and root lengths and dry weights, leaf area, and relative water contents of soybean after 21 days of growth
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Table 3 Effect of different bacterial treatments on Chl.a, Chl.b, Chl a/Chl b, and carotenoid contents of soybean after 21 days of growth
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The lowest value of electrolyte leakage was recorded in plants treated with B. japonicum + Delta PSK, which also caused lipid peroxidation and malondialdehyde (MDA) content to decrease as well as hydrogen peroxide (Table 4). Levels of citric acid and proline contents were at the lowest values; however, total soluble sugars were at the highest level in response to that combined treatment. The measured osmotic pressure was at the lowest value for plants bacterized with B. japonicum + Delta PSK. Compared with the control, the total antioxidant content of soybean shoots increased significantly after bacterial treatment. Plants bacterized with B. japonicum + Delta PSK showed the lowest values for total antioxidants, reducing power, and DPPH free radical scavenging activity (Table 5). The activities of antioxidant enzymes catalase, peroxidase, and polyphenol oxidase decreased significantly in response to bacterization and B. japonicum + Delta PSK combined treatment led to the lowest activity of these enzymes.

Table 4 Effect of different bacterial treatments on electrolyte leakage, lipid peroxidation, hydrogen peroxide, osmotic pressure, citric acid, proline, and total soluble sugar contents of soybean after 21 days of growth
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Table 5 Effect of different bacterial treatments on the antioxidant capacity, antioxidant enzymes activity, and the secondary metabolites of soybean plant after 21 days of growth
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The growth parameters and pigment contents, assessed after 19 days of salinity treatment decreased considerably according to the stress level. Generally, soybean growth was reduced as far as NaCl conc increased, however, bacterization particularly with B. japonicum + Delta PSK, considerably alleviated this deleterious impact of both salinity levels (Table 6). The contents of both chlorophylls (a and b) and carotenoids considerably decreased according to salinity level. This effect was retarded particularly in response to the combined bacterial treatment (B. japonicum + Delta PSK) as shown in Table 7. However, the ratio of Chl. a/ Chl. b did not significantly change in response to different treatments. As indicated in Tables 6 and 7, salinity affected significantly most of the measured growth parameters except dry weights, and leaf area, however, different bacterial treatments considerably affected all the growth parameters. Salinity and bacterial treatments affected pigments but Chl. a/ Chl. b was not considerably affected. The combined effect of salinity and bacterial treatments was considerable on growth parameters, chlorophylls, and carotenoids.

Table 6 Effect of different bacterial treatments in combination with three salinity levels (0, 50, and 100 mM NaCl) on shoot and root lengths and dry weights, leaf area, relative water contents, and number of nodules of soybean after 40 days of growth
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Table 7 Effect of different bacterial treatments in combination with three salinity levels (0, 50, and 100 mM NaCl) on Chl. a, Chl. b, Chl. a/Chl. b, and carotenoid contents of soybean after 40 days of growth
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According to NaCl conc, electrolyte leakage, MDA, and hydrogen peroxide values increased, however, this effect was reduced considerably in response to different bacterial treatments (Table 8). The levels of citric acid, total soluble sugars, proline, and consequently osmotic pressure increased in response to salinity. Both NaCl, bacterial treatments, and their combinations considerably affected these parameters.

Table 8 Effect of different bacterial treatments in combination with three salinity levels (0, 50, and 100 mM NaCl) on electrolyte leakage, lipid peroxidation, hydrogen peroxide, osmotic pressure, citric acid, proline, and total soluble sugar contents of soybean after 40 days of growth
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The antioxidant capacity, reducing power, DPPH free radical scavenging activity, and total phenolic and flavonoid contents in soybean plants increased significantly, particularly in the presence of B. japonicum + Delta PSK at different salinity levels (Fig. 1). Bacterial treatments considerably affected all these parameters but salinity did not show a significant effect on phenols and flavonoids. The combination between salinity and bacterial treatments developed a considerable change in these parameters. The activity of antioxidant enzymes (catalase, peroxidase, and polyphenol oxidase) increased progressively, particularly in the presence of B. japonicum + Delta PSK (Figs. 1 and 2) along with salt treatments. Salinity, bacterial treatments, and their combinations significantly affected the activity of these enzymes.

Fig. 1
figure 1

Effect of different bacterial treatments in combination with three salinity levels (0, 50, and 100 mM NaCl) on a total antioxidant capacity, b reducing power, c DPPH (1,1-diphenyl-2-picrylhydrazyl), d total phenolic contents, and e total flavonoid contents of soybean after 40 days of growth. Fisher’s test at p < 0.05 reveals significant variations in means (± standard error), which are different letters (a–k) on the same bars. *, **, and *** imply significance levels of 0.01, 0.05, and 0.001, respectively

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Fig. 2
figure 2

Effect of different bacterial treatments in combination with three salinity levels (0, 50, and 100 mM NaCl) on the activity of a catalase (CAT), b peroxidase (POX), and c polyphenol oxidase (PPO) of soybean after 40 days of growth. Fisher’s test at p < 0.05 reveals significant variations in means (± standard error), which are different letters (a–k) on the same bars. *, **, and *** imply significance levels of 0.01, 0.05, and 0.001, respectively

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Discussion

Beneficial plant bacteria have been shown to have a dual role in improving plant responses to abiotic and biotic stressors in addition to promoting the growth and productivity of plants (Badawy et al. 2022). Their interaction with plant metabolism and hormonal balance directs the synthesis of secondary metabolites required to alleviate stress. These effects represent PGPBs as good solutions for salinity problems (Dawood, Sofy et al. 2022) and effective alternatives for chemical fertilizers (Abu-Shahba, Mansour et al. 2022). The data represented in the current study show the impact of such beneficial organisms and their role in mitigating salinity stress on soybean.

In this study, the physical growth parameters recorded after 21 and 40 days of growth (Tables 2 and 6, respectively) showed that soybean-bacterized plants, especially those that received B. japonicum + Enterobacter Delta PSK with and without stress, surpassed control plants in their growth. It is apparent that salinity stress directs plants toward metabolism to produce the materials needed to counter these conditions. The Enterobacter strain used in this study was selected because of its ability to promote plant growth (Agha, Abbas et al. 2021). Additionally, several reports identified Enterobacter as a potential PGPB (Panigrahi et al. 2020, Sheteiwy et al. 2021, Pérez‐Rodriguez, Pontin et al. 2022). B. japonicum is regarded as a compatible symbiont of soybean (Ramongolalaina et al. 2018). The ability of B. japonicum + Enterobacter Delta PSK to produce high amounts of indole-3-acetic acid and gibberellic acid might be responsible for stimulating growth. This ability remained even in the presence of salt (Agha, Abbas et al. 2021). The production of indole acetic acid (IAA) supports root growth and proliferation (Tables 2 and 6). Enterobacter has a high ability to colonize plant roots, especially the primary roots, and specifically in the elongation zone. It also could colonize and re-open stomatal cells in the host plant, indicating the ability to establish even under severe conditions (Synek et al. 2021). The ACC deaminase activity reported for Enterobacter (Singh, Pandey et al. 2022) might be among the mechanisms for its ability to alleviate salt stress by reducing ethylene levels.

Due to osmotic stress and ion toxicity, salinity dramatically reduces the ability of plant roots to absorb water (Alsaeedi et al. 2019). Furthermore, ROS are generated more rapidly (ALHaithloul, Khan et al. 2022). Under salt stress, the mechanism of photosynthesis and its efficacy are inevitably compromised and damaged (Agha, Abbas et al. 2021). In this study, bacterization particularly with B. japonicum + Enterobacter Delta PSK increases the chlorophylls contents even under salinity stress indicating the ability of this strain to alleviate the oxidative and ionic stress of NaCl (Tables 3 and 7). The same result was observed in tomato plants (Pérez‐Rodriguez, Pontin et al. 2022) in response to Enterobacter. Both Bacillus subtilis and Pseudomonas fluorescens have exhibited substantial potential for enhancing photosynthetic pigment biosynthesis under salt stress in pea plants (Sofy et al. 2021). In salt-stressed Vigna radiata plants, Enterobacter cloaca has been found to positively affect photosynthesis and the levels of chlorophyll, carotenoids, and all photosynthetic pigments (Bhise et al. 2017). Enhanced P, N, and K absorption might underlie PGPB's stimulation of pigment production in soybean plants (Zhao et al. 2018). The prominent increase in total soluble sugars in response to the combination treatment supports this speculation (Tables 4 and 8). Furthermore, the significant increase in carotenoids in response to bacterization supports the ability of Delta PSK to activate plant antioxidant defenses (Tables 3 and 7). The same response was recorded in Enterobacter-treated salinity-stressed tomato plants (Pérez‐Rodriguez, Pontin et al. 2022).

Salinity decreased the number of nodules (Table 6), but combination-treated plants showed higher nodule numbers, probably due to the ability of the used Enterobacter to produce siderophores (M. Mowafy, S. Agha et al. 2022) that make iron available, which is essential for nodule formation. The effect of PGPB on nodulation under salinity stress has been previously reported (Elsheikh and Wood 1990).

After 21 days of growth, salinity stress was not applied, indicating that the observed responses, including the profiles of citric acid, proline, total soluble sugar contents, total antioxidant, reducing power, DPPH free radical scavenging activity, the activities of CAT, POX, and PPO, total phenolic contents, and total flavonoid contents, were due to bacterization (Tables 4 and 5). The induction of phenols and proline in bacterized plants indicates the activation of the defense response. These results complement those recorded in grapevine plantlets in response to Burkholderia phytofirmans (Ait Barka et al. 2006). The detectable increase in flavins (Table 4) has also been reported in another study in Astragalus sinicus inoculated with Burkholderia cepacian under iron deficiency conditions (Zhou et al. 2018).

Due to NaCl stress, soybean produced higher levels of MDA and hydrogen peroxide and showed greater EL, which caused membrane damage and lipid peroxidation (Table 8). Membrane integrity was maintained in bacterized (B. japonicum + Enterobacter Delta PSK) plants and the detected amounts of MDA and EL were lower too. Damodaran et al. (2014) reported that membrane damage during NaCl stress altered membrane function in mustard cultivars. The same effect was reported in Astragalus sinicus plants inoculated with B. cepacian (Zhou et al. 2018).

Although NaCl stimulates antioxidant enzymes, inoculating the plant with B. japonicum + Delta PSK further boosts the antioxidant process and removes toxic ROS. The same effect was recorded, although in another experimental purpose, in which Enterobacter cloacae antagonized the effect of Ralstonia solanacearum on potato plant (Mohamed et al. 2020). In the Haber–Weiss reaction, CAT is a vital antioxidant enzyme that scavenges superoxide radicals, hydroxyl radicals, and hydrogen peroxide (Sofy, Mancy et al. 2022). Therefore, CAT, POX, and PPO are upregulated in soybean plants to protect membranes from free radical damage caused by NaCl in this study. The increased POX activity in the combination-treated plants might be attributed to improved lignin biosynthesis and other antioxidant compounds that reduce oxidative stress as reported previously in Vigna unguiculata under salt stress Chen et al. (2007a, b).

Salinity stress for 19 days significantly affected plant growth and metabolism across the different treatment groups. The physical growth parameters were reduced with salt concentration but bacterization improved them compared with the control. Salinity induces a remarkable reduction in plant growth, pigmentation, and metabolism (Parida et al. 2004; Shabala and Cuin 2008). Various strains of PGPB alleviate this devastating effect under salinity stress (Hamdia et al. 2004; Rojas-Tapias et al. 2012), as corroborated by our results. As shown previously, the increase in the number of root nodules in bacterized plants under salinity stress supports the role of PGBP in enhancing growth and nodulation (M. Mowafy, S. Agha et al. 2022). Compared with the control, the increase in chlorophyll content in Delta PSK-treated plants is the real reason for better growth, indicating the compatibility of the Enterobacter strain used in this study with soybean plants and its ability to relieve salinity stress.

The ability to produce indole-3-acetic acid, siderophores, and solubilized phosphate may have conferred upon Delta PSK the ability to support soybean growth under salinity stress (Agha, Abbas et al. 2021), resulting in antioxidant responses via regulation of osmolyte accumulation and the development of free radical scavenging activities. The same conclusion was drawn in several studies (Mishra, Mishra et al. 2021; Fouda and Sofy 2022). Simultaneously, the amounts of citric acid, total soluble sugars, and proline increased leading to an elevated internal osmotic pressure in the combination-treated plants compared with the control. This was accompanied by lower lipid peroxidation and EL, indicating the integrity of the cellular membrane under such stress. Similar results have been reported in other studies (Prittesh et al. 2020, Pérez‐Rodriguez, Pontin et al. 2022). Furthermore, the increase in TA capacity (Fig. 1) and antioxidant enzyme activity (Fig. 2) reflects the stimulatory effect of B. japonicum + Delta PSK on systematic resistance, which eventually decreases the levels of ethylene, a metabolite that accumulates due to salinity stress (Fan et al. 2020). Further, plants that received Delta PSK showed higher levels of total phenolic compounds (Fig. 1), a result reported previously for Enterobacter-treated tomato and maize plants (Ali et al. 2022, Pérez‐Rodriguez, Pontin et al. 2022).

Conclusion

The mutualistic interactions with proper PGPB can increase plant ability to withstand abiotic stress for their numerous direct and indirect mechanisms including phytohormones production, ACC deaminase activity, and nutrient sequestration which are beneficial to plant growth and productivity. This study demonstrated the ability of Enterobacter Delta PSK to serve as a soybean growth promoter under salinity stress leading to improved physiological and growth parameters. The impact of these isolates on soybean yield is currently being reported. An open-field experiment is ongoing to ensure the effectiveness of Enterobacter Delta PSK along with B. japonicum on soybean under salinity stress in the local environment.