Drought priming and plant growth‐promoting rhizobacteria enhance Solanum lycopersicum metabolites under drought stress

Abstract

BACKGROUND

Drought stress is an important factor that consistently decreases plant productivity. Bio‐stimulants such as plant growth‐promoting rhizobacteria (PGPR) and arbuscular mycorrhizal fungi (AMF) have been shown to modulate stress‐related pathways. The aim of this study was to enhance the metabolic memory of tomato under different drought stress conditions using these bio‐stimulants. Two weeks after planting, air‐dried sieved soil mixed with 150 g of Glomus deserticola and 1 mL of 10¹⁰ CFU mL⁻¹ Enterobacter ludwigii was applied to the root zone. Drought treatments included control, drought stress, and drought priming.

RESULTS

Drought priming increased the accumulation of proline, phenolics, and anthocyanins compared with drought stress (DS). Compared with control plants under drought priming, combining PGPR with drought priming enhanced ATPase by 81%, proline by 137%, soluble sugar by 296%, and phenylalanine ammonia‐lyase by 12%. Phenylalanine ammonia‐lyase is actively involved in the biosynthesis of secondary metabolites like flavonoids, which increased by 60%, and anthocyanins, which increased by 75%. An increase in biomass, a 100% survival rate, and low malondialdehyde content were observed. Combining AMF and drought priming significantly increased proline (108%), soluble sugar (240%), flavonoids, antioxidant enzymes, and phenylalanine ammonia‐lyase, resulting in a survival rate of 66.67%. Plant growth‐promoting rhizobacteria and AMF also enhanced the accumulation of NO, which is necessary for antioxidant enzyme accumulation.

CONCLUSION

Integrating PGPR with drought priming constitutes an effective strategy for enhancing immediate stress resilience and strengthening metabolic responses. Further research using epigenetic approaches is recommended to uncover drought stress memory under drought priming, with a focus on PGPR. © 2026 Society of Chemical Industry.

INTRODUCTION

Droughts are a recurring environmental challenge that plants encounter throughout their life cycles. ¹ Drought conditions lead to a decrease in key photosynthetic pigments, particularly chlorophyll a and b. This reduction culminates in lower total chlorophyll levels and a diminished presence of chlorophyll‐binding proteins. ² Prolonged drought further impairs the activity of ribulose bisphosphate, making it difficult for plants to absorb carbon dioxide (CO₂). It also reduces stem conductivity and decreases stomatal conductance. ³ These metabolic challenges contribute to a decline in photosynthesis, adversely affecting the production of ribulose bisphosphate. ⁴ Yield losses in the field due to drought typically range from 30% to 90%, varying among different crop species. ⁵ In Nigeria, major tomato‐producing states such as Benue, Kano, and Plateau recorded a 25% reduction in yield attributable to variability in rainfall. ⁶ Addressing the influence of drought stress is crucial for safeguarding tomato production and global food security.

Plants respond differently to repeated stress than to initial exposure, a phenomenon known as stress memory. ¹ This adaptive response is triggered by stress priming – including drought, heat, and flood priming – which enables plants to modify subsequent physiological and molecular responses. Drought priming, in particular, equips plants with the ability to retain information from previous stress events and adjust their responses to future stress conditions. This retention involves the storage of stress‐related signaling information and the capacity to retrieve it when re‐exposed to similar stress, which enhances tolerance. ⁷ Stress memory can be categorized into three distinct forms: somatic stress memory, which involves mitotically heritable changes that persist only within the lifespan of an individual plant; intergenerational stress memory, which is transferred to the immediate offspring; and transgenerational stress memory, which involves meiotically heritable changes observable in subsequent stress‐free generations. ⁸

Somatic stress memory is short‐term stress memory induced by stress priming, enabling plants that have undergone a stress event to utilize stored information for days, weeks, or even months, thus enhancing their response when faced with future stressors. ⁹ This mechanism is particularly well characterized in the processes known as cold hardening and drought priming. ⁹ Recent research on drought priming has demonstrated its role in maintaining reactive oxygen species (ROS) homeostasis, enhancing photosynthetic rates, and ultimately leading to increased grain yields when plants are subsequently exposed to abiotic stresses – including low temperatures, drought, and heat.10, 11

Arbuscular mycorrhizal fungi and plant growth‐promoting rhizobacteria (PGPR) are agents that colonize the rhizosphere, enhancing plant growth through various mechanisms. ¹² They enhance plant growth and improve drought stress tolerance in various ways. ¹³ They increase nutrient uptake, which supports overall plant health. They also promote root development, resulting in improved water absorption due to an expanded root surface area. These bacteria also play a pivotal role in controlling hormones like cytokinin, abscisic acid (ABA), indole‐3‐acetic acid (IAA), and gibberellic acid, and produce aminocyclopropane‐1‐carboxylate (ACC) deaminase, which lessens ethylene concentrations associated with plant stress. ¹⁴ Plant growth‐promoting rhizobacteria can also modify gene expression by altering the root framework and activate defense techniques via antioxidant generation.15, 16 They can boost root hair production, promote lateral root growth, and increase total root size, facilitating improved water uptake and nutrition through the synthesis of IAA.17, 18 Plant growth‐promoting rhizobacteria can modify the homeostasis of hazardous ions. They minimize the buildup of Na⁺ and Cl⁻ in leaves by supporting ion removal from root cells and activating the production of ion transporters, such as high‐affinity K⁺ transporters. These transporters facilitate accumulation in shoots while precluding these ions from leaves, ¹⁹ thereby regulating Na⁺ transportation in plants and forestalling intolerable Na⁺ concentrations in roots. ²⁰

Arbuscular mycorrhizal fungi (AMF) (Glomus deserticola), on the other hand, play a fundamental role in supporting plant health by creating a nutrient‐rich environment, primarily through the provision of phosphates. ²¹ In this symbiotic relationship, the plant supplies carbohydrates to the fungi, and the AMF develops a specialized mycelium that facilitates the transfer of carbon products generated during photosynthesis to the soil microbiome. ²² This mycelium, along with living hyphal exudates that are rich in carbohydrates, can promote bacterial growth in the soil due to their high turnover. ²³ Moreover, colonization by AMF leads to the accumulation of osmo‐protectants, such as soluble sugars and proteins, which play a crucial role in osmotic adjustment and maintaining cellular stability during water deficit conditions. ²⁴ This symbiotic relationship also enhances the architecture of the root system, facilitating improved soil exploration and resource acquisition. Notably, AMF mitigate oxidative damage associated with drought stress by reducing malondialdehyde (MDA) levels, a key marker of lipid peroxidation, through the activation of antioxidant defense mechanisms.25, 26 Together, these processes contribute to increased biomass production and enhance overall plant resilience in the face of drought stress. ²⁷

This study posits that co‐inoculation of PGPR and AMF will produce synergistic effects on plant drought tolerance by augmenting morphological, physiological, and biochemical adaptive responses beyond what is achieved by either microorganism alone. The individual contributions of PGPR and AMF in enhancing plant drought tolerance are well documented, but little is known about the combined effects of drought priming and bio‐stimulants on short‐term metabolic memory. Thus, the novelty of this study lies in examining the application of PGPR and AMF as bio‐stimulants that can enhance drought‐induced metabolic memory, ultimately improving plant resilience to subsequent drought events.

MATERIALS AND METHODS

Soil and microbial samples collection

Enterobacter ludwigii (a PGPR) was isolated from tomato root rhizosphere soil at the research farm, University of Ilorin, Ilorin, Kwara State, Nigeria. Soil tightly attached to tomato plants was collected in Ziplock bags and transported to the laboratory for isolation. Soil at a depth of 0–20 cm was collected at abandoned land, air dried, and sieved. The tomato cultivar ‘HORTI Tom 3’ was procured from the National Institute for Horticultural Research (NIHORT), Jericho, Ibadan, Nigeria. Exactly 3 kg of soil was measured into each labeled pot and placed in the screenhouse of the Federal University Oye‐Ekiti, Nigeria (7° 46' 36.98" N 5° 18' 54.94" E) with an average temperature ranging between 25 and 35 °C. Arbuscular mycorrhizal fungi (150 g) containing 15% spores per g were homogenously mixed with 2.85 kg sieved soil to make up to 3 kg soil before planting.

Isolation of PGPR

To isolate PGPR from these samples, serial dilution and subsequently the pour‐plate technique were employed using nutrient agar medium as the growth substrate. The prepared plates were incubated at a controlled temperature of 30 °C for 24 h to facilitate the development of bacterial colonies. After incubation, distinct colonies were selected and further subcultured on fresh nutrient agar to ensure the establishment of pure cultures free from contamination.

Identification and screening of PGPR

Enterobacter ludwigii (Isolate 2) was molecularly identified and screened for PGPR traits. According to David et al., ²⁸ the experimental isolate demonstrated the ability to solubilize phosphate, produce ACC deaminase, and generate siderophores. The nucleotide sequence has been deposited in the National Center for Biotechnology Information (NCBI) database under GenBank accession number PV276927 (https://submit.ncbi.nlm.nih.gov/).

Planting and PGPR application

The PGPR isolated from tomato was used on other varieties of tomatoes. Pots were grouped and labeled as control, AMF (soil mixed with Glomus deserticola before planting), and PGPR. Sterilized seeds were planted into the sieved soil, watered with 200 mL of water until field moisture capacity was reached, and thinned to three plants per pot during the first week of planting. Two weeks after planting, 1 mL of 10¹⁰ CFU mL⁻¹ Enterobacter ludwigii (Isolate 2) was applied directly to the root of each plant in the PGPR group using a sterile syringe. All pots were watered daily with 200 mL of water.

Drought stress applications

Pots were further divided into subgroups: (i) control (CNT), (ii) drought stress (DS) – consisting of plants subjected to 7 days of drought stress – and (iii) drought priming (DP) – consisting of plants subjected to 4 days of drought stress followed by a first recovery period of 3 days before the main 7 day drought stress period. Drought stress was introduced in the fourth week after planting by withdrawing water for 4 days (for primed pots) and 7 days (for both DS and primed pots), following a method described by Sintaha et al. ²⁹ Control pots were watered continuously with 200 mL of water every day. The following nine treatments were used: control + CNT (no stress or biostimulants), control + DS, control + DP, control + PGPR, PGPR + DS, PGPR + DP, control + AMF, AMF + DS, and AMF + DP. Each treatment was replicated three times, with 12 plants per treatment. All plants underwent a second recovery period and were watered until field moisture capacity was reached again for 3 days. After this second recovery stage, which was in the sixth week, the experiments were terminated, and fresh shoots, roots, and biomass were measured, and collected for biochemical analysis (Fig. 1).

Figure 1.

Experimental design and drought treatments. ADS: air‐dried soil; Tom 3: tomato seed variety; DS1: 4 days drought stress; R1: 3 day recovery; DS2: 7 day drought stress; R2: 3 day recovery; CNT: control. Three bio‐stimulant treatments were applied: control (CNT), plant growth‐promoting rhizobacteria (PGPR), and arbuscular mycorrhizal fungi (AMF). Three drought treatments were imposed – control, drought priming (DP), and drought stress (DS). In primed plants, a 4‐day drought stress was applied before the main 7 day drought stress, whereas DS plants were subjected only to 7 days of drought stress. All plants were allowed a 3 day recovery period after each stress phase.

Biochemical assays

The plant shoots were analyzed to determine the activity of phenylalanine ammonia lyase and superoxide dismutase, as well as the levels of flavonoids, phenolic compounds, anthocyanins, nitric oxide, and MDA. Analysis of the root system focused on the activity of peroxidase (POD), which plays a critical role in plant metabolism.

To extract chlorophyll and carotenoid pigments from the leaves, a solution containing 75% (v/v) ethanol was utilized. Following extraction, the absorbance of the resultant extracts was determined at specific wavelengths of 470 nm (for carotenoids), 663 nm, and 645 nm (chlorophyll a and b), using a spectrophotometer (BioBase BK‐UV1600 Spectrophotomer, S/N: UPA 19D0004) to quantify the pigment concentrations.

Free proline content was quantified following Bates et al. ³⁰ Briefly, 0.5 g frozen leaf tissue was homogenized in 3 mg mL⁻¹ sulfosalicylic acid, filtered, and reacted with acid‐ninhydrin and glacial acetic acid. After incubation at 100 °C for 1 h, the chromophore was extracted with toluene, and the absorbance was measured at 520 nm. Results were expressed as μmol g⁻¹ fresh weight. The soluble sugar content was assessed using the 3,5‐dinitrosalicylic acid (DNSA) method, following a procedure modified from Jain et al. ³¹ The DNSA reagent was prepared by dissolving 1 g of DNSA and 30 g of sodium–potassium tartaric acid in 80 mL of 0.5 mol L⁻¹ NaOH at 45 °C. Once dissolved, the solution was cooled to room temperature and diluted to 100 mL with distilled water. For measurement, 2 mL of DNSA reagent was added to a test tube containing 1 mL of the plant extract (1 mg mL⁻¹) and heated at 95 °C for 5 min. After cooling, 7 mL of distilled water was added to the mixture, and the absorbance was measured at 540 nm using an ultraviolet‐visible (UV‐visible) spectrophotometer. The reducing sugar content was determined from the calibration curve of standard d‐glucose (200–1000 mg L⁻¹), and results were expressed as mg of d‐glucose equivalent (GE) per g of dry extract weight.

Phenylalanine ammonia lyase (PAL) activity was measured using the method described by Rahe et al. ³² For this photometric assay, 200 μL of enzyme extract was combined with 800 μL of a 15 mmol L⁻¹ l‐phenylalanine solution in borate buffer (25 mmol L⁻¹, pH 8.8) and incubated at 37 °C for 1 h. The reaction was stopped by adding 50 μL of concentrated HCl (5 mol L⁻¹). A control sample, prepared simultaneously with d‐phenylalanine, served as a blank. The absorbance of the samples was measured at 290 nm against the blank, and PAL activity was expressed as nanomoles of cinnamic acid formed per hour per unit of protein content.

Adenosine triphosphatase (ATPase) activity was measured as μmol of inorganic phosphate (P_i) released per mg of protein, following the protocol established by Rule et al. ³³ The color development reagent was prepared as follows:

• Solution A: 10 mg mL⁻¹ ammonium molybdate in 1 mol L⁻¹ HCl

• Solution B: 60 mg mL⁻¹ ascorbic acid in distilled water

• Solution C: 20 mg mL⁻¹ sodium citrate/2% (v/v) acetic acid

• Solution D: Equal parts of Solutions A and B mixed immediately before use, resulting in a yellow or slightly greenish‐yellow solution.

For the reaction, the following constituents were prepared in a test tube: 100 μL of 500 mM Tris–HCl (pH 8.0), 20 μL of 100 mM MgCl₂, 100 μL of 1 mol L⁻¹ KCl, 5 μL of 100 mmol L⁻¹ dithiothreitol, and 500 μL of the sample. This mixture was incubated at 37 °C for 5 min. The reaction was initiated by adding 60 μL of 10 mmol L⁻¹ adenosine triphosphate (ATP). To stop the reaction, 50 μL of the mixture was added to 50 μL of 10 mg mL⁻¹ sodium dodecyl sulfate. Color development was achieved by adding 100 μL of Solution D and 125 μL of Solution C. The mixture was incubated at 37 °C for 20 min, and the absorbance was measured at 850 nm. The amount of phosphate produced was quantified using a phosphate standard.

Total phenolic content was determined using the method outlined by Singleton and Rossi. ³⁴ Folin–Ciocâlteu's phenol reagent was used as an oxidizing agent. Briefly, a mixture was prepared using 0.1 mL of the sample and 0.9 mL of water, to which 0.2 mL of Folin–Ciocâlteu's phenol reagent was added before vortexing. After the mixture was allowed to stand for 5 min, 1.0 mL of a 7% (w/v) Na₂CO₃ solution was introduced, and the total volume was diluted to 2.5 mL. This solution was then incubated at room temperature for 90 min. The absorbance of the sample was measured at a wavelength of 750 nm against a negative control containing 1 mL of water instead of the sample. Gallic acid at a concentration of 0.1 mg mL⁻¹ was used as the standard to establish the gallic acid equivalent (GAE) of the sample following the preparation of a calibration curve. Distilled water served as the blank for the measurements.

Flavonoid content in leaf samples was quantified following established procedures. Distilled water (0.4 mL) was added to 0.1 mL of the extract or standard, followed by the addition of 0.1 mL of 50 mg mL⁻¹ sodium nitrite. After 5 min, 0.1 mL of 10 mg mL⁻¹ aluminum chloride and 0.2 mL of 1 mol L⁻¹ sodium hydroxide were added, and the volume was adjusted to 2.5 mL with distilled water. The absorbance was measured at 510 nm against a blank. The total flavonoid content was expressed as mg of quercetin equivalents per g of the plant extract. Total anthocyanin content was evaluated spectrophotometrically using the pH differential method, based on protocols established by Şakar et al. ³⁵ A pH 1.0 buffer solution was prepared by mixing 125 mL of 0.2 mol L⁻¹ KCl with 385 mL of 0.2 mol L⁻¹ HCl and 490 mL of distilled water. The pH was adjusted to 1.0 with 0.2 mol L⁻¹ HCl. A pH 4.5 buffer solution was prepared by mixing 440 mL of 1.0 mol L⁻¹ sodium acetate with 200 mL of 1.0 mol L⁻¹ HCl and 360 mL of distilled water, with the pH adjusted to 4.5 using 1.0 mol L⁻¹ HCl. A 0.5 mL aliquot of the extract was diluted to 12.5 mL in both the pH 1.0 and 4.5 buffers and allowed to equilibrate in the dark for 2 h. The absorbance of the samples at 512 nm (A ₅₁₂) and 700 nm (A ₇₀₀) was measured using a UV‐visible spectrophotometer. The anthocyanin content was expressed as 10⁻⁵ kg of cyanidin 3‐glucoside kg⁻¹ of sample, using a molar absorbance coefficient (ε) of 26 900 L mol⁻¹ cm⁻¹. ³⁶

Catalase (CAT) activity was measured using the method described by Aebi, ³⁷ monitoring the decomposition of H₂O₂ at 240 nm. Activity was calculated using the extinction coefficient (43.6 L mol⁻¹ cm⁻¹) and expressed as U mg⁻¹ protein. Superoxide dismutase (SOD) levels in the leaves were analyzed using the method established by McCord and Frederich. ³⁸ Tris–HCl buffer (2.5 mL of 75 mmol L⁻¹, pH 8.2), 30 mmol L⁻¹ ethylenediaminetetraacetic acid (EDTA), and 300 μL of 2 mmol L⁻¹ pyrogallol were added to 200 μL of the plant extract. The increase in absorbance was recorded at 420 nm for 3 min using a spectrophotometer (BK‐UV1600 Spectrophotomer, S/N: UPA 19D0004). Nitric oxide (NO) levels in the extracts were evaluated using the protocol described by Marcocci et al. ³⁹ An aliquot of the sample (100 μL) was added to each labeled Eppendorf tube, followed by 0.1 mL of 10 mg mL⁻¹ sulfanilamide in 50 mg mL⁻¹ phosphoric acid. This mixture was incubated at room temperature for 10 min before adding 0.1 mL of 0.1% N‐1‐naphthylethylenediamine dihydrochloride (NED) and incubating for another 10 min at 60 °C. The absorbance of the chromophore formed was measured at 546 nm. A standard curve was prepared using sodium nitrite (100 μmol L⁻¹), and the amount of NO in the sample was extrapolated from the curve. Finally, the concentration of MDA, an indicator of lipid peroxidation that signifies oxidative damage to cellular membranes, was determined using the thiobarbituric acid (TBA) method, following protocols established by Ohkawa et al. ⁴⁰ Phosphate buffer (0.5 mL of 0.1 mol L⁻¹, pH 8.0) and 0.5 mL of 24% (v/v) trichloroacetic acid (TCA) were added to 0.5 mL of the sample. This mixture was incubated at room temperature for 10 min and then centrifuged at 671 x g  for 20 min. The supernatant (1 mL) was combined with 0.25 mL of 3.3 mg mL⁻¹ TBA in 20% (v/v) acetic acid, and the resulting mixture was boiled at 95 °C for 1 h. The resultant pink product was cooled, and the absorbance was read at 532 nm (extinction coefficient of MDA, ε532 = 1.53 × 10⁵ L mol⁻¹ cm⁻¹).

Data analysis

All data were analyzed using analysis of variance (ANOVA) followed by Duncan's multiple range tests to separate means at a significance level of P ≤ 0.05. Bar charts and graphs were compiled using GraphPad Prism version 8.0. One of the images was created using BioRender (https://BioRender.com).

RESULTS

Phenotypic and chlorophyll content – response to drought stress

Plant growth‐promoting rhizobacteria increased root weight significantly under DS conditions (Fig. 1). As drought stress advanced, both root and shoot weights declined under primed drought conditions compared with untreated and unstressed plants (CNT). Root weight increased in plants treated with DP + AMF compared with DS + AMF. Notably, plants treated with PGPR showed increased root and shoot weights under the DS condition. This improvement was reflected in survival rates, which reached 100% for plants under control and PGPR conditions during both DS and primed drought scenarios, whereas survival rates were only 20% and 66.67% for AMF in DS and primed states, respectively. Arbuscular mycorrhizal fungi showed lower fresh biomass and a 20% survival rate under DS conditions (Fig. 2). Under drought priming, no differences were observed in the roots with or without bio‐stimulant treatments. Chlorophyll a in the control plants ranged from 21.60 to 21.81 regardless of the bio‐stimulants treatments; chlorophyll b ranged from 13.00 to 14.46, and carotene ranged from 1.12 to 1.44. Drought priming increased the chlorophyll b content (DS + CNT vs DP + CNT) and decreased the carotenoid content (DS + CNT vs DP + CNT). The chlorophyll b content of PGPR‐treated plants remained unchanged for each drought treatment (control, DS, DP). For AMF‐treated plants, DS and DP increased chlorophyll b compared with the control (Table 1).

Figure 2.

Morphological traits of tomato under drought stress. Arbuscular mycorrhizal fungi (AMF), plant growth‐promoting rhizobacteria (PGPR), and control (CNT) represent the bio‐stimulant treatments, and control, drought stress (DS), and drought priming (DP) are the drought treatments. Fresh shoot, root, and total biomass weights were recorded. Error bars represent standard errors. Mean values followed by the same letter are not significantly different at P ≤ 0.05 (Duncan's Multiple Range test). Plant growth‐promoting rhizobacteria positively boosted root, shoot, and biomass weights, leading to 100% survival under the DS and DP conditions.

Table 1.

Chlorophyll (mg g⁻¹ FW) content of the tomato plant under drought stress

	Treatment	Control	DS	DP
Chlorophyll a	CNT	21.60aA	21.85aA	21.65aA
	PGPR	21.67aA	21.60aA	21.68aA
	AMF	21.81aA	21.74aA	21.70aA
Chlorophyll b	CNT	14.46aA	12.99bB	14.50aA
	PGPR	14.63aA	14.44aA	14.55aA
	AMF	13.29bA	14.67aA	14.39aA
Carotenoids	CNT	1.21abB	1.50aA	1.21aB
	PGPR	1.18bA	1.21bA	1.20aA
	AMF	1.44aA	1.18bB	1.23aB

Note: Mean values sharing the same lower case letter in a column indicate no significant difference in bio‐stimulant effect, whereas values sharing the same upper case letter across a row indicate no significant difference in drought or priming effect at P ≤ 0.05 (Duncan's Multiple Range test). Fresh leaves of the plant were collected for chlorophyll analysis. FW, Fresh weight; DS, Drought stress; DP, Drought priming; CNT, Control soil without PGPR or AMF. Plant growth‐promoting rhizobacteria (PGPR) and arbuscular mycorrhizal fungi (AMF) showed no effect on chlorophyll a and b under the drought stress conditions.

Osmoregulatory role of PGPR and AMF in drought stress

Osmolytes such as proline (276% increase) and soluble sugars (114% increase) were accumulated as a response to drought under DS conditions, with PGPR‐treated plants showing the highest osmolyte content. A greater accumulation of proline (136% increase) and soluble sugar (296% increase) was observed under drought priming (Table 2 and Fig. 3). Arbuscular mycorrhizal fungi combined with drought priming also increased proline content and soluble sugar by 108% and 240%, respectively, compared with the control plants. The same trend was observed under DS conditions (Table 2).

Table 2.

Osmolyte content of tomato under drought stress

	Treatment	Control	DS	DP
Proline (mg mL−1)	CNT	0.022bB	0.021cB	0.034bA
	PGPR	0.036aB	0.079aA	0.080aA
	AMF	0.023bB	0.065abA	0.071aA
Soluble sugar (mg mL−1)	CNT	0.410aB	0.510bA	0.250cC
	PGPR	0.260cB	1.090aA	0.990aA
	AMF	0.440aB	0.930abA	0.850bA

Note: Mean values sharing the same lower case letter within a column indicate no significant difference in bio‐stimulant effect, whereas values sharing the same upper case letter across a row indicate no significant difference in drought or priming effects at P ≤ 0.05 (Duncan's Multiple Range test). DS, Drought stress; DP, Drought priming; CNT, Control soil without PGPR or AMF. Under DP and drought (DS) conditions, plant growth‐promoting rhizobacteria (PGPR) increased proline and soluble sugar content.

Figure 3.

Proposed synergistic effect of drought priming and plant growth‐promoting rhizobacteria (PGPR). Plants increase metabolite accumulation in response to drought stress, contributing to drought tolerance (DT). At the recovery (RC) stage, drought stress (DS) plants exhibit metabolic homeostasis and acclimation. In contrast, drought‐primed (DP) plants maintain elevated metabolic activity under recurrent drought stress.

Phenylalanine ammonia lyase and ATPase activity under drought stress

Drought priming was shown to reduce PAL activity in all treated plants compared with the CNT group (without stress and bio‐stimulants); however, activity in PGPR‐treated plants was the highest. Under DS conditions, PGPR intensified PAL action by 12%, but this was followed by a decrease in primed plants (Table 3). Drought stress increased the ATPase enzymes under both conditions; the ATPase activity in plants under drought stress ranged from 15.09 to 26.02 μmol P_i per mg protein. Under DS and primed drought conditions, PGPR increased ATPase activity by 100% and 81%, respectively, compared with the CNT (Table 3). Plants treated with AMF recorded reduced ATPase compared with CNT in both DS and primed conditions.

Table 3.

Phenylalanine ammonia lyase and ATPase under drought stress

	Treatment	Control	DS	DP
Phenylalanine ammonia lyase (PAL) (μmol cinnamic acid per mg protein)	CNT	25.75aA	11.19aB	8.07bC
	PGPR	10.03bB	11.51aA	9.04aC
	AMF	9.04cA	5.12bC	8.44bB
Adenosine triphosphatase (ATPase) (μmol Pi per mg protein)	CNT	10.64cC	26.02bA	15.09bB
	PGPR	43.12aB	52.04aA	27.24aC
	AMF	25.37bA	10.10cC	11.81cB

Note: Mean values sharing the same lower case letter within a column indicate no significant difference in bio‐stimulant effects, whereas values sharing the same upper case letter across a row indicate no significant difference in drought or priming effect at P ≤ 0.05 (Duncan's Multiple Range test). Plant growth‐promoting rhizobacteria (PGPR) induced phenylalanine ammonia lyase (PAL) and adenosine triphosphatase (ATPase) in drought stress (DS) and drought priming (DP) conditions.

Drought priming and bio‐stimulants accumulate secondary metabolites

Drought stress induced phenolic compound and flavonoid accumulation compared with CNT plants; however, drought priming significantly increased these levels further. In the presence of PGPR, drought priming led to a greater accumulation of flavonoids (60% increase) and anthocyanins (75% increase), further enhancing stress memory compared with control plants (Table 4 and Fig. 3). Phenolic compounds in drought‐priming plants were significantly higher than in the treated plants (AMF and PGPR) (Table 4).

Table 4.

The secondary metabolites of tomato plants under drought stress

	Treatment	Control	DS	DP
Phenolics (mg GAE g⁻¹)	CNT	164.770bB	172.840bA	179.050aA
	PGPR	163.060bA	167.410aA	141.010bB
	AMF	250.020aA	118.030cC	128.740cB
Flavonoids (mg QUE g⁻¹)	CNT	96.220cC	133.220cA	121.160cB
	PGPR	156.510aB	193.100aA	194.140aA
	AMF	133.220aB	168.980bA	168.150bA
Anthocyanin (10−5 kg of cyanidin 3‐glucoside kg−1 of sample)	CNT	350.670bA	125.242cB	275.532cA
	PGPR	350.097bB	459.219bA	480.093aA
	AMF	647.082aA	530.190aB	325.628bC

Note: Mean values sharing the same lower case letter within a column indicate no significant difference in bio‐stimulant effect, whereas values sharing the same upper case letter across a row indicate no significant difference in drought or priming effect at P ≤ 0.05 (Duncan's Multiple Range test). Plants increased phenolic and flavonoid compounds in response to drought stress (DS). Flavonoids and anthocyanins were consistently higher in plants treated with DP + plant growth‐promoting rhizobacteria (PGPR).

Antioxidant enzymes in plants under drought stress

Figure 4 shows the level of antioxidant enzyme activity during drought stress. Superoxide dismutase accumulated in response to DS and primed drought conditions under PGPR and AMF treatments compared with CNT plants. Under DS conditions, AMF increased CAT activity by 100% compared with CNT plants.

Figure 4.

Antioxidant defensive system under drought stress. Arbuscular mycorrhizal fungi (AMF), plant growth‐promoting rhizobacteria (PGPR), and control (CNT) represent the bio‐stimulant treatments, and control, drought stress (DS), and drought priming (DP) are the drought treatments. Error bars represent standard errors. Mean values followed by the same letter are not significantly different at P ≤ 0.05 (Duncan's Multiple Range test). Arbuscular mycorrhizal fungi and PGPR increased superoxide dismutase (SOD) activity under DS and DP conditions. Arbuscular mycorrhizal fungi also increased catalase (CAT) activity under DS.

Biosynthesis of NO and MDA content in drought‐stressed plants

Plant growth‐promoting rhizobacteria and AMF significantly elevated the concentration of nitric oxide (NO) in both DS and DP plants under drought stress compared with the CNT group. In plants exposed to DS, the MDA content was higher in those treated with PGPR and AMF. However, when drought priming was combined with PGPR treatment, a reduction of over 30% in MDA content was observed (Fig. 5).

Figure 5.

Nitric oxide and malondialdehyde (MDA) content of tomato plants under drought stress. Arbuscular mycorrhizal fungi (AMF), plant growth‐promoting rhizobacteria (PGPR), and control (CNT) represent the bio‐stimulant treatments, and control, drought stress (DS), and drought priming (DP) are the drought treatments. Error bars represent standard errors. Mean values followed by the same letter are not significantly different at P ≤ 0.05 (Duncan's Multiple Range test). Plant growth‐promoting rhizobacteria and AMF accumulated NO equally under DS and DP. The MDA content was reduced in plants treated with PGPR under primed drought stress conditions.

DISCUSSION

Root development plays an important role in plant adaptation to drought stress. These features exhibit plasticity under water‐limited environments and could be critical for developing crops with efficient root systems for adaptation under drought. ⁴¹ Plant growth‐promoting rhizobacteria mediate the effect of drought stress through the development of large roots, thereby improving plant biomass. Various studies report auxin production by rhizobacteria‐primed crop plants. ⁴² In wheat plants Bacillus thuringiensis showed elongation of root hairs through auxin and ACC deaminase synthesis, aiding the absorption of soil nutrients and water. ⁴³ Plant growth‐promoting rhizobacteria can associate with plants promptly by augmenting the accessibility of nutrients – such as nitrogen, phosphorus, and iron – which could enhance nutrient uptake and growth. ⁴⁴ Furthermore PGPR promoted better root structural defense in comparison with AMF. Plant colonization by PGPR improves plant growth and productivity due to their ability to enhance root elongation and phytohormone production. ²⁶

Drought priming exerted a significant positive impact on root weight. A combination of drought priming and PGPR or AMF stabilized the root weight. This demonstrates the beneficial effect of drought priming on the survival of plants under drought stress, and its synergistic effect with the bio‐stimulants could enhance tolerance. It has been established that drought priming can change the shape and density of stomata, improve water‐use efficiency, and reduce whole‐plant transpiration of potato. ⁴⁵ Tahiri et al. ⁴⁶ highlighted the combined benefits of AMF with other bio‐stimulants, such as PGPR and compost, for enhancing drought tolerance in comparison with their individual effects. A combination of AMF with drought priming increased the survival percentage from 20% in DS plants to 66.67%, while the PGPR‐treated plant showed consistent 100% survival rate in the DS and DP plants.

Drought often reduces photosynthesis due to diminished CO₂ diffusion from the atmosphere to the carboxylation site. ¹¹ Plants treated with PGPR or AMF under DS conditions have increased chlorophyll b levels, which are crucial for capturing the light energy necessary for glucose production and metabolic processes. The levels of chlorophyll a, b, and carotenoids remained relatively unchanged under drought priming, regardless of the treatment. Abdallah et al. ⁴⁵ reported that drought priming improved the photosynthetic performance and yield of potato under drought stress.

Osmolytes are organic solutes that protect cellular structure and maintain cell turgor under drought stress. Plant growth‐promoting rhizobacteria and AMF increased the accumulation of proline and soluble sugars under both drought stress conditions, indicating that PGPR enhanced osmotic adjustment during drought. However, osmolyte accumulation was greater in PGPR‐treated plants subjected to drought priming. The accumulation of osmolytes helps maintain root turgor, facilitating water absorption. Similar findings have been reported in fonio, where increased proline accumulation improved tolerance to osmotic shock. ⁴⁷ Enhanced stomatal conductance, osmotic adjustment, and reduced oxidative stress have also been attributed to the accumulation of glycine, cysteine, and betaine in PGPR‐treated plants. ⁴⁸ In wheat, increased proline and betaine accumulation contributed to enhanced drought tolerance following drought priming. ⁴⁹ Plant growth‐promoting rhizobacteria facilitate proline accumulation, and this effect is further enhanced by drought priming, suggesting that both treatments contribute to strengthening plant memory of recurrent stress events. Similarly, AMF colonization markedly improved drought tolerance in Cinnamomum migao by increasing levels of SOD, CAT, POD, soluble sugars, soluble proteins, and proline. ⁵⁰

Phenylalanine ammonia‐lyase is a key enzyme in the phenylpropanoid pathway and plays an important role in plant reactions to drought stress. ⁵¹ Arbuscular mycorrhizal fungi exhibited the opposite effect, increasing PAL activity in primed plants relative to their DS counterparts. Oxidative phosphorylation and phenylalanine metabolism and biosynthesis were triggered by AMF in response to drought stress. ²⁴ Phenylalanine ammonia‐lyase is essential for the secretion of secondary metabolites like phenols, flavonoids, and lignin, which contribute to plant tolerance by protecting against oxidative damage. Previous research has found that, under drought stress, PGPR induced a 6.3‐fold upregulation of PAL and a 0.6‐fold reduction in flavone synthase expression in maize. ⁴⁸ Phenylalanine ammonia‐lyase links primary and secondary metabolism through the phenylpropanoid route, which branches into multiple additional pathways. The enzyme catalyzes the deamination of phenylalanine to produce cinnamic acid, a precursor for the biosynthesis of other phenylpropanoids, including flavonoids, phenols, and lignin. ⁵²

Adenosine triphosphatases adjust their activity to ensure effective hydrolysis and pumping under varying conditions. ⁵³ Drought stress triggered the accumulation of ATPases under DS and DP conditions. H⁺‐ATPases are involved in ABA‐mediated stomatal closure to reduce water loss under drought stress. ⁵⁴ Upon activation, plasma membrane H⁺‐ATPase acidifies the apoplast, followed by the activation of enzymes involved in cell‐wall loosening. ⁵⁵ In the presence of PGPR, ATPases increased fourfold under non‐drought conditions. Auxin activates plasma membrane H⁺‐ATPases, which are believed to be the initial step in hypocotyl elongation. ⁵⁶ According to Liu et al., ⁵⁷ PGPR is involved in triggering hormones regulating K⁺ homeostasis and stomatal movement. Although abscisic acid (ABA) plays a dominant role in stomatal closure during drought stress, ATPases, particularly the plasma membrane H⁺‐ATPase, play a crucial role in maintaining cellular homeostasis and enabling plant survival. Maintenance of the membrane potential to regulate K⁺ transmembrane transport is critical for plant tolerance to drought. ⁵⁸ Enterobacter ludwigii (a plant growth‐promoting rhizobacterium) in the rhizosphere maintained a high cytosolic K⁺/Na⁺ ratio, which facilitated the movement of stomata and cell membrane integrity. ⁵⁷

Phenolics, flavonoids, and anthocyanins play a pivotal role in how plants react to drought stress. These compounds act as antioxidants, protecting plants from oxidative damage. Plants accumulated phenolics and flavonoids as a response to drought stress. However, DP improved metabolic memory, resulting in greater accumulation of flavonoids and anthocyanins, particularly in PGPR‐treated plants. Phenolics, flavonoids, and anthocyanins are stress‐induced metabolites that help scavenge free radicals, reducing damage caused by ROS. These compounds also contribute to osmotic balance and the regulation of stress‐responsive pathways, improving plant resilience under drought conditions. ⁵⁹ These results indicate that flavonoids, anthocyanins, and phenolics play important roles in drought stress responses, leading to greater tolerance. Anthocyanins, a class of flavonoid, also regulate compounds related to stress, including melatonin, proline, ABA, and salicylic acid. ⁶⁰ Flavonoids also contribute to drought tolerance by stabilizing cellular membranes and regulating stress‐responsive pathways. ⁶¹

Antioxidant enzymes such as SOD, CAT, and POD are key components of the active oxygen removal system. ⁶² Plant growth‐promoting rhizobacteria and AMF accumulated CAT and SOD in DS, which might indicate the maintenance of cell homeostasis and acclimation to stress. Previous studies have shown that PGPR strains, including Enterobacter and Pseudomonas fluorescens, improved SOD, CAT, and ascorbate peroxidase (APX) antioxidant systems under salinity stress. ⁶³ Similarly, AMF treatment increased CAT activity by 42% and APX activity by 66% in tomato leaves compared with non‐AMF‐treated plants. ⁶⁴ Nitric oxide, which is involved in stress responses, was higher in PGPR and AMF than in the control under drought stress. Beyond its role in drought tolerance, NO contributes to stomatal regulation, activation of antioxidant enzymes, and reduction of oxidative damage. ⁶⁵ Although NO is not essential for proline accumulation, drought stress induces both proline and endogenous NO accumulation in rice leaves. ⁶⁶ The involvement of PGPR induced NO and activated pathways that led to morphological and physiological modifications ultimately improved drought tolerance. Malondialdehyde content decreased under primed drought conditions, indicating reduced membrane leakage. This effect may also be associated with proline‐mediated maintenance of cell turgor. Abdallah et al. ⁴⁵ reported that primed plants exhibited increased accumulation of total sugar and proline, resulting in improved water status maintenance. Reduced levels of oxidative stress indicators, including hydrogen peroxide (H₂O₂), MDA, and electrolyte leakage in primed plants further confirmed alleviation of oxidative stress in olive (Olea europaea ‘Chétoui’), consistent with the present findings. ⁴⁵ Similar results were reported by Gontia‐Mishra et al., ⁶⁷ who showed that the PGPR Enterobacter ludwigii increased vegetative growth, increased relative water content, reduced MDA levels, and increased accumulation of proline and other stress‐associated metabolites under drought stress. The responses of plants to drought stress under E. ludwigii treatment are consistently high, indicating that they store metabolites for improved tolerance to future stress events.

CONCLUSION

Drought priming was a critical regulator of stress memory, modulating physiological and biochemical responses such as osmolyte accumulation, antioxidant metabolism, and secondary metabolite biosynthesis. The enhanced phenols, anthocyanins and lowered MDA content indicated that primed plants can be better equipped to mitigate oxidative damage and maintain membrane integrity.

Plant growth‐promoting rhizobacteria and drought priming play important and in many cases complementary roles in enhancing plant resilience to drought stress. In this study, PGPR application markedly improved plant biomass, survival rate, osmolyte accumulation, antioxidant defense, and ATPase activity, while reducing lipid peroxidation, highlighting its role in maintaining cellular homeostasis and promoting adaptive growth under stress. In contrast, AMF alone exhibited comparatively limited effectiveness under DS drought conditions but exhibited substantial improvement when combined with drought priming.

Future research should elucidate the molecular and epigenetic mechanisms underlying drought stress memory, particularly gene regulatory networks influenced by drought priming and Enterobacter ludwigii. Integrative studies combining PGPR and AMF under field conditions are also needed to optimize synergistic interactions and enhance sustainable crop resilience to drought stress.

CONFLICT OF INTEREST

The authors declare no conflict of interest.

AUTHOR CONTRIBUTIONS

Oyinade A. David conceptualized and designed the experiment, analyzed data, and wrote the first draft. Idowu Olawuni analyzed the biochemical assays. Oluwatoyin A. Fabiyi isolated and characterized the PGPR and reviewed and edited the final draft. Deborah M. Chukwuma and Joshua S. Ameh edited and reviewed the final draft. Peace I. Kolawole‐Idowu, Boluwatife O. Adedayo, Quareebat A. Shontan, Jacob V. Onimisi, and Amarachi F. Onyema contributed to the field work and methodology.

DATA AVAILABILITY STATEMENT

The data that support the findings of this study are available on request from the corresponding author. The data are not publicly available due to privacy or ethical restrictions.