Screening and Characterization of Rhizobacteria as PGPR for Enhancing Growth and Yield of Aceh Patchouli

2.1 Study area

Soil samples were collected using purposive sampling from the rhizosphere area of healthy plants at several smallholder patchouli plantations located in Aceh Province, as shown in Figure 1, which includes the following locations: Alue Abed, Calang (4°33'19.2"N95°45'43.4"E), Krueng Itam (4°01’02.9"N96°23’46.3"E), Purwosari (4°04’29.4"N 96°16’30.6"E), Lambada (5°30'45.6"N 95°35'40.5"E), Paya Tampu (4°46’23.5"N 96°21’58.8"E), Air Tenang (4°18’24.7"N 98°04’06.7"E, Tanah Terban (4°18’45.0"N 98°03’52.3"E), Bandar Mahligai Sekerak (4°16’28.4"N 98°01’23.0"E), Pulo Gelime (4°08’25.9"N 97°07’29.4"E).

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Figure 1. Soil sampling map from the patchouli rhizosphere in several districts of Aceh Province

2.2 Isolation of rhizobacteria

In this study, the isolation method that was used for rhizobacteria was serial dilution. Initially, soil samples from several regions (Figure 1) were collected in 1 kg quantities from five different locations. Soil samples from each site were air-dried at room temperature (28-30℃) for 48 hours and then pooled [18]. After drying, the soil was sieved using a 9-mesh sieve shaker. One gram of the sieved soil was placed in a test tube, and 9 ml of distilled water was added. The mixture was vortexed at 100 rpm to create a uniform solution, which was labelled as the first dilution (10⁻¹). A 1 mL sample of the 10⁻¹ dilution was transferred to a new test tube with 9 ml of dilution solution, creating the second dilution (10⁻²). This process was repeated until the final dilution of 10⁻⁹. Nutrient Agar (NA) was prepared in Petri dishes, allowed to solidify, and used to isolate rhizobacteria. A 1 mL sample from dilutions 10⁻⁴ to 10⁻⁹ was plated in duplicate on NA agar plates and incubated at 28℃ for 48 hours. After incubation, bacterial identification was carried out, and pure rhizobacterial cultures were obtained.

2.3 Physiological identification

2.3.1 IAA production test

The Glickmann and Dessaux approach was used to assess the rhizobacterial isolates' capacity to generate IAA. The isolates were cultured on Sucrose Peptone Agar (SPA) medium for a whole day [19]. Each media received 0.5 g/l of the amino acid tryptophan to promote auxin production, which was then centrifuged for 10 minutes at 10,000 rpm. After being removed from the bacterial pellet, the supernatant (centrifugation liquid) was filtered through 0.2 μm Millipore filter paper, and its IAA level was measured. Salkowski's reagent was used to determine the amount of IAA present in the bacterial culture filtrate. A 2 ml test tube or an Eppendorf tube was filled with 1 ml of the bacterial culture filtrate and 1 ml of Salkowski's reagent. The absorbance was then measured using a spectrophotometer calibrated at a wavelength of 550 nm after 30 minutes of incubation in the dark at 26℃. To determine the amount of IAA in the bacterial culture filtrate, a standard curve based on the absorbance values of pure IAA solutions with concentrations of 0, 6.25, 12.5, 25, 50, 75, 100, 150, and 200 µg/ml was employed [20].

2.3.2 Phosphate solubilizing ability test

Phosphate source in an agar-based medium to examine the rhizobacteria's capacity to solubilize phosphate. In 1000 milliliters of distilled water, 15 grams of agar, 10 grams of glucose, 0.2 grams of KCl, 0.2 grams of NaCl, 5 grams of Ca₃(PO₄)₂, 0.5 grams of yeast extract, 2.5 grams of MnSO₄, 2.5 grams of FeSO₄, and 0.1 grams of MgSO₄ were dissolved to create the medium, which was then autoclaved to ensure sterilization. Using a cork borer, three wells were created in the test medium before it was transferred into Petri dishes. 0.2 ml of the rhizobacterial suspension under test was added to each well, and the wells were then incubated for seven days at 28℃. According to study [21], the development of a clean zone surrounding the well holding the bacterial suspension demonstrated the rhizobacteria's capacity to solubilize phosphate. The Phosphate Solubilizing Index (PSI) was computed using the clear zone that developed around each isolate. Following seven days of incubation, measurements were made of the colony's and the clear zone's diameters [22]. The following formula was used to determine the PSI:

$P S I=\frac{A+B}{B}$               (1)

where, A=Diameter clear zone (cm), B=Colony diameter (cm), PSI=Phosphate Solubilizing Index.

2.3.3 Nitrogen fixation ability test

A medium including MgSO₄·7H₂O (25 g), FeSO₄·7H₂O (0.01 g), NaMoO₄·2H₂O (0.01 g), MnSO₄·5H₂O (0.01 g), CaCl₂ (0.1 g), K₂HPO₄ in NaCl buffer (5 mL), and Agar (17–20 g) was used to examine the nitrogen fixing ability of rhizobacteria isolates. Pure isolates that were 48 hours old were cultivated on the medium, incubated for four days at room temperature (28℃), and the growth of the bacteria was monitored. The growth of the isolate in nitrogen-free media, as demonstrated by the shift in the liquid medium's turbidity, served as a signal for isolates that could fix nitrogen [20].

2.3.4 Hydrogen Cyanide (HCN) production test

A technique proposed by Bakker and Schipper [23] was used to qualitatively examine the formation of hydrogen cyanide (HCN) by rhizobacteria. 4.4 g of glycine, 2 g of picric acid, 8 g of sodium carbonate, 15 g of agar, 30 g of TSB, 1000 mL of distilled water, and pieces of sterile filter paper were used to prepare the testing medium. After dissolving glycine, TSB, and agar in 1000 milliliters of sterile distilled water and autoclaving the solution, it was transferred into Petri dishes. Two grams of picric acid and eight grams of sodium carbonate were dissolved in 200 milliliters of sterile distilled water to prepare the HCN detection solution. Sterile filter paper pieces were soaked in this solution before being placed in the Petri dish lid, and bacteria were then streaked onto the glycine medium. The filter papers were then placed in the center of the Petri dish lid. The plates were incubated at 28℃ for seven days. During incubation, HCN production by rhizobacteria was qualitatively observed. Isolates that did not cause any color change in the filter paper indicated no HCN production. HCN-producing isolates induced the following color changes in the filter paper: from yellow to light brown (+HCN low), light brown to dark brown (++HCN moderate), dark brown to brick red (+++HCN high), and dark red to black (++++HCN very high).

2.3.5 Siderophore production test

The production of siderophores by rhizobacterial isolates was tested by culturing the bacteria in Chrome Azurol S (CAS) medium, a standard method for detecting and screening siderophore producers. The test was carried out for 24 hours at room temperature. The medium was prepared with the following composition per liter: 20 g sucrose, 2 g L-asparagine, 1 g K₂HPO₄, and 0.5 g MgSO₄. After centrifuging rhizobacterial suspensions for 30 minutes at 11,000 rpm, the supernatant was filtered through a 0.2 µm nitrocellulose membrane. A control sample of supernatant without FeCl₃ was compared to 3 mL of the supernatant after 1 mL of 0.01 M FeCl₃ was added to evaluate siderophore formation. A spectrophotometer set at 410 nm was used to measure the synthesis of siderophores [24].

2.3.6 ACC-Deaminase production test

Following the protocol of Glick [25], ACC-Deaminase activity was assessed using Dworkin-Foster (DF) minimal salt medium supplied with the substrate 1-aminocyclopropane-1-carboxylate (ACC) or ammonium sulfate as the nitrogen source. Four grams of KH₂PO₄, six grams of Na₂HPO₄, 0.2 grams of MgSO₄·7H₂O, one milligram of FeSO₄·7H₂O, ten μg H₃BO₃, ten μg MnSO₄, seventy μg ZnSO₄, fifty μg CuSO₄, ten μg MoO₃, two grams of glucose, two grams of gluconic acid, two grams of citric acid, and twelve grams of agar (for solid media) were all dissolved in 1000 milliliters of distilled water. To the DF medium, 2 g of ammonium sulfate was added. All parts were then autoclaved for 30 minutes at 121℃ and 0.1 MPa to disinfect them. This was followed by inoculation with 1 mL of bacterial suspension (1 × 10⁹ cells/mL) onto both DF and DF + ammonium sulfate media. Growth of the isolates was observed after 48 hours, following the method described by the study [26]. Isolates that grew on DF + ammonium sulfate showed the presence of ACC-Deaminase activity. The DF medium without ammonium sulfate was used as a control to determine when the isolates were nitrogen-fixing bacteria capable of obtaining nitrogen by fixing N₂ from the air.

2.3.7 Analysis of soil nutrient content

In this study, soil was analyzed at the Agricultural Instrument Standardization Application Center's Testing Service Laboratory in Aceh Province before and after the application of rhizobacteria. Mon Alue in Aceh Besar's Indrapuri District provided the soil utilized as the planting medium. K-available (ppm), K-exchangeable, Na-exchangeable, Ca-exchangeable, Mg-exchangeable, C-Organic (%), N-Total (%), C/N ratio, pH H₂O, and Cation Exchange Capacity (CEC) (meq/100g) were among the soil metrics [27].

The organic carbon and total nitrogen content of the soil are measured to perform the carbon-to-nitrogen (C/N) ratio analysis. 0.5 g of air-dried soil sample is reacted with 1 N potassium dichromate (K₂Cr₂O₇) and concentrated sulfuric acid (H₂SO₄), shaken, and left to stand before being titrated with 0.5 N ferrous sulfate (FeSO₄) using ferroin indicator until the color changes from blue-green to red-brown. This is the Walkley and Black [28] method for determining organic carbon. Meanwhile, total nitrogen analysis follows the Kjeldahl method (1883) where 0.5 g of soil sample is digested with concentrated H₂SO₄ and a catalyst, then distilled with 40% NaOH to convert ammonium (NH₄⁺) into ammonia gas (NH₃), which is then trapped in 4% boric acid solution and titrated with 0.1 N H₂SO₄ or HCl using bromocresol green and methyl red indicator [29]. The C/N ratio is determined by dividing the organic carbon content by the total nitrogen content, serving as an indicator of the organic matter decomposition rate in the soil.

Cation exchange capacity (CEC) was analyzed using the ammonium acetate method at pH 7.0 [30]. A 5 g air-dried soil sample was placed in a centrifuge tube and mixed with 25 mL of 1 N ammonium acetate (CH₃COONH₄) solution at pH 7.0. The mixture was shaken for 30 minutes to exchange the cations adsorbed on soil colloids with ammonium ions (NH₄⁺). The solution was then separated from the soil by filtration or centrifugation, and the filtrate was analyzed to determine the concentrations of exchangeable cations such as calcium (Ca²⁺), magnesium (Mg²⁺), potassium (K⁺), and sodium (Na⁺). Inductively Coupled Plasma-Optical Emission Spectrometry (ICP-OES) or Atomic Absorption Spectroscopy (AAS) was used to quantify these cations. The total amount of exchangeable cations in cmol(+) kg⁻¹ soil was used to calculate CEC [31].

2.4 Agronomic characterization

2.4.1 Preparation of growing media and plant material

The initial step included preparing the planting medium by mixing topsoil and compost at a 1:1 ratio. For two weeks, this soil mixture was sterilized using 20 g m^-3 of the soil sterilant Basamid G. A pH meter was used to measure the medium's pH after sterilization; 6-7 is the ideal pH range. Subsequently, planting materials were prepared in the form of stem cuttings with 5-6 leaves from each variety. The varieties used in this study were Tapak Tuan, Lhokseumawe, and Sidikalang. The stem cuttings are initially propagated for 21 to 30 days using the sungkup (humidity chamber) method. This method ensured uniform and healthy growth of the patchouli seedlings.

2.4.2 Preparation of rhizobacteria suspension

Fifteen rhizobacteria isolates from various regions that showed superior physiological performance in preliminary tests, were selected to investigate their effects on patchouli plant growth. Without coming into contact with the growing medium, these isolates were cultivated and suspended in 100 milliliters of sterile distilled water. Using a spectrophotometer, the population density of the suspension was calculated to achieve a concentration of 10⁹ CFU mL^-1, corresponding to an absorbance of OD600 = 0.192 [32]. Subsequently, the bacteria suspension was transferred to culture bottles for application.

2.4.3 Planting and application of rhizobacterial isolates

After four weeks of growth, the patchouli seedlings were ready for transplantation into polybags measuring 40 × 25 cm, containing a 2:1 mixture of soil and compost with a total volume of 10 kg. The soil used for planting was obtained from the patchouli rhizosphere region. Following transplantation, a suspension of indigenous rhizobacteria was applied to the seedlings. The inoculation was carried out once a week after transplantation, with 100 mL of the bacterial suspension applied around the root zone of each plant. Other maintenance activities included regular watering and weed, pest, and diseases control to ensure healthy growth.

2.5 Molecular identification and phylogenetic tree construction

One popular technique in molecular biology is the polymerase chain reaction (PCR). PCR is widely used in molecular analysis, especially for amplifying specific regions (amplicons) and identifying specific genes within a sample. It is based on the ability of DNA polymerase to synthesize a new DNA strand from a given template. In molecular biology, real-time PCR, also known as quantitative real-time PCR (qPCR), is becoming increasingly popular. qPCR can be used to detect the presence of a specific gene, compare expression levels across different samples, and quantify gene abundance. In this study, DNA extraction from rhizobacteria was performed using the DNeasy Blood and Tissue Kit (https://ptgenetika.com/Lab. Genetika, Banten). After being moved to an Eppendorf tube with 500 µL of buffer solution, the genomic DNA was crushed with a grinder. After centrifuging the sample for ten minutes at 13,000 rpm, the supernatant was carefully poured into a fresh Eppendorf tube. Subsequently, 450 µL of a 24:25:1 Phenol: Chloroform: Isoamyl alcohol (PCI) solution was added, and the mixture was shaken for five minutes. After centrifuging the mixture for ten minutes at 13,000 rpm, the supernatant was carefully transferred into a new Eppendorf tube. Finally, to precipitate the DNA, 500 µL of cold 100% ethanol was added [27].

2.6 Observation parameters

The study's characteristics included the rhizobacterial isolates' capacity to fix nitrogen, solubilize phosphate, generate IAA, and create the enzyme ACC deaminase. At 12 weeks following application (WAA), measurements were also made of the plant's height (cm), number of leaves, number of branches, root volume, wet weight (g), and dry weight (g).

At 30, 60, and 90 days after application (DAA), established techniques were used to measure the plant growth characteristics. While the number of leaves and branches was counted based on completely formed organs, the height of the plant was measured from the base of the stem to the growing point. At 90 DAT, the leaf area was examined using ImageJ software. Following harvesting and drying, fresh and dry biomass were weighed using a digital balance. A ruler was used to measure the length of the roots, and a graduated cylinder's water displacement method was used to calculate the root volume.

2.7 Data analysis

This study employed a randomized block design (RBD) with a factorial arrangement. The first factor was the type of indigenous rhizobacteria (R) sourced from a prior study. The identification results indicated that these indigenous rhizobacteria functioned as biofertilizers and biostimulants, as demonstrated by their IAA production and phosphate solubilization abilities. This factor included 15 levels: R0 =no rhizobacteria (control), and R1 to R14. The second factor was the variety (V), which had three levels: V1 =Lhokseumawe variety, V2 =Tapak Tuan variety, and V3 =Sidikalang variety. As a result, there were 45 treatment combinations, replicated three times, leading to 135 experimental units, each consisting of two plants, totalling 270 samples. Data were analyzed using Analysis of Variance (ANOVA), and when the F-test showed significant effects (α =5%), further analysis was performed using Duncan's New Multiple Range Test (DNMRT) at the 5% significance level.

An auxin hormone called IAA is important for plant growth and development because it promotes tissue differentiation, lateral and adventitious root production, and cell division and elongation. These characteristics help roots and stems grow longer, which improves the plant's capacity to take up nutrients and water from the soil [33, 34]. Numerous factors, including the presence of microorganisms like rhizobacteria, can affect the formation of IAA. Numerous bacteria that live in the rhizosphere of plants produce the phytohormone IAA. These bacteria use a variety of IAA biosynthesis pathways, and some strains have multiple pathways. Study [34] shows that rhizosphere isolates may generate IAA at different doses up to 3.609 mg/L. Study [35] claimed that the soil microbiota influenced plant growth and development and contributed to health. The rhizosphere and roots of Ceanothus velutinus yielded 24 PGPR-producing IAA, which were found to promote plant growth. This finding highlighted how crucial rhizobacteria bacteria are for promoting plant growth by producing vital growth hormones.

Several studies have demonstrated that the application of IAA can stimulate plant growth, leading to increases in height, root length, and other growth parameters [36]. Thus, IAA plays a vital role in promoting physical growth and improving plants' ability to absorb water and nutrients from the soil. For instance, study [37] found that Serratia marcescens strain MBC1 produced IAA, resulting in significant increases in root and stem length in soybean plants. Furthermore, IAA also influenced plant growth in hydroponic systems, as evidenced by its significant effect on celery plants.

IAA production was demonstrated by rhizobacteria isolated from patchouli plants, which improved plant growth and yield [38]. IAA, which is crucial for physiological activities such as lateral root development and stem elongation, was produced by several rhizobacteria isolates in this investigation at different doses. Higher total yields were the result of patchouli plants' vegetative development being promoted by the high concentration that rhizobacteria produced. Bacteria that produce IAA can also increase the productivity of Arabidopsis biomass production and plant propagation [35]. Furthermore, it has been demonstrated that IAA promotes gas exchange and stem diameter expansion, which both improve coffee plant growth [39]. In this study, patchouli plants' stem diameter and shoot and leaf counts were enhanced by the use of IAA-producing rhizobacteria isolates. This demonstrated that the isolates promoted the physical development and general production of the plants.

L-tryptophan is one of the organic substances that naturally occur in plant roots and are utilized by rhizobacteria for IAA production to resist biotic and abiotic stress [36]. Rhizobacteria that produce IAA encourage the host's root system to flourish, which may lessen the requirement for chemical pesticides on patchouli plants. Other studies have shown that rhizobacteria such as Pseudomonas fluorescens, Stenotrophomonas rhizophila, and Bacillus cereus enhance plant growth and resistance to pathogens [40, 41]. According to study [42], IAA-producing rhizobacteria isolates were shown to be effective in inducing plant resistance to viral infections and reducing disease intensity in tobacco plants. This mechanism also applies to patchouli plants, where rhizobacteria compete with pathogens and produce antimicrobial compounds to protect against disease attacks [35]. Enterobacter sp. capable of producing IAA significantly enhances maize growth by increasing root length, plant height, fresh weight, and dry weight [36]. Therefore, the application of IAA-producing rhizobacteria can be an environmentally friendly alternative to reduce dependence on chemical pesticides. The results indicated that all rhizobacterial isolates were capable of producing IAA. The rhizobacteria with the highest IAA production included Enterobacter sp., Delftia tsuruhatensis, and Paenibacillus polymyxa. In contrast, Pseudomonas aeruginosa produced less IAA compared to the other species.

Competition for iron consumption through siderophore synthesis is one of the ways rhizobacteria regulate plant diseases [20]. In the rhizosphere, different Plant Growth-Promoting Bacteria (PGPB) help plants absorb iron which speeds up the physiological and biochemical processes of plants in adverse situations. These bacteria can produce and release siderophores, which improve plant nutrition and growth, and they can also control the bioavailability of iron for plants [43-45]. Numerous kinds of bacteria, including Bacillus, Enterobacter, Klebsiella, Paenibacillus, Pseudomonas, and Streptomyces, have been documented to produce siderophores [46-50]. Pseudomonas aeruginosa and Enterobacter sp. were among the rhizobacteria isolates in this investigation that were able to produce siderophores.

Previous studies reported that siderophore-producing rhizobacteria had significant potential in enhancing plant resistance to pathogens and environmental stress. Study [51] found that rhizobacteria isolates from local rice plants (Kamba) could produce siderophores, effectively inhibiting pathogen growth by binding Fe³⁺ for pathogen development. Additionally, siderophore-producing rhizobacteria can help mitigate heavy metal stress in Panax ginseng, enhancing the plant's resilience to harmful environmental conditions [52]. Some rhizobacterial isolates, such as Glutamicibacter nicotianae and Brevibacillus invocatus, have demonstrated the ability to produce siderophores, significantly promoting maize growth [53]. These isolates contribute to improved nutrient absorption, enhancing leaf quality and plant growth rates. As a result, rhizobacteria that produce siderophores can act as potential biocontrol agents, supporting environmentally friendly and sustainable agricultural practices [51].

Both organic and inorganic forms of phosphate are found in soil naturally; these forms are insoluble or only weakly soluble, which restricts the amount of phosphate that soil organisms can use. In acidic soils, inorganic phosphate minerals are typically bonded as AlPO₄·2H₂O (variscite) and FePO₄·2H₂O (strengite); in alkaline soils, they are typically bound as Ca₃(PO₄)₂ (tricalcium phosphate). Phosphate-solubilizing microbes that generate organic acids with functional carboxyl (-COO⁻) and hydroxyl (-O⁻) groups can aid in the release of phosphate from these bonds. This can raise the amount of soluble phosphate in the soil by forming complexes with cations such Ca₃(PO₄)₂, FePO₄·2H₂O, or AlPO₄·2H₂O [54]. In this study, the rhizobacteria isolates capable of solubilizing phosphate included Pseudomonas aeruginosa.

Various types of bacteria capable of solubilizing phosphate in the soil have been identified, including Burkholderia, Azotobacter, Pseudomonas, Bacillus, Enterobacter, Citrobacter, and Pantoea. Numerous organic acids, including oxalate, succinate, tartrate, citrate, lactate, α-ketoglutarate, acetate, formate, propionate, glycolate, glutamate, glyoxylate, malate, and fumarate, are produced by the metabolism of phosphate-solubilizing bacteria. When it comes to phosphate solubilization, each of these bacteria creates a variety of organic acids in variable quantities and types [55]. It has been demonstrated that phosphate can be efficiently dissolved by phosphate-solubilizing rhizobacteria, including Bacillus species and Pseudomonas species [30]. Furthermore, by producing growth-promoting hormones and having antibiosis actions against pathogens, rhizobacteria improve plant growth and yield [56].

The process of transforming atmospheric nitrogen into a form that plants can absorb, known as nitrogen fixation, promotes growth and raises plant output. Despite being present in the atmosphere in large quantities (about 80%) as N₂, plants are unable to directly absorb nitrogen. This method has not been effective in maximizing nitrogen use efficiency, as a significant amount of nitrogen is lost into the atmosphere and water bodies, leading to negative environmental impacts. Therefore, nitrogen fixation in plants offers a practical solution to decrease the reliance on synthetic nitrogen fertilizers and enhance the sustainability of food production systems [57]. Previous studies reported that bacteria with good nitrogen fixation activity had been found in species such as Pseudomonas aeruginosa and Brucella intermedia.Ara.

PGPR such as Bacillus pumilus S1r1 can be a biological alternative for fixing nitrogen from the atmosphere and delaying remobilization in plants, potentially increasing harvest yields [58]. The nitrogenase enzyme system, consisting of two metalloproteins (iron and molybdenum-iron proteins), plays a significant role in converting dinitrogen (N₂) into ammonia (NH₃) through ATP [59]. Various studies have shown some examples of rhizobacteria capable of nitrogen fixation, including Azospirillum spp., Bradyrhizobium spp., Paenibacillus spp., and Paraburkholderia phymatum [60, 61]. This nitrogen fixation process enhances plant growth and supports sustainable agriculture through improved nitrogen efficiency in the soil.

Through the induction of defensive mechanisms against attacks, HCN contributes positively to plant responses to pathogens. Although it is produced in small amounts during the basic metabolism of plants, HCN can be released in larger quantities after tissue damage in cyanogenic species, becoming an important defense mechanism in plants [62]. Various studies have shown that many rhizobacteria, including strains of Pseudomonas, Bacillus, and Enterobacter, are capable of producing HCN. By producing extracellular lytic enzymes and secondary metabolites, PGPR can successfully manage Vicia faba root rot and wilt illnesses. Subsequent research revealed that 34 out of 104 rhizobacteria isolated from the patchouli rhizosphere produced HCN, which shielded plants from infections and other dangerous creatures [63-66]. Halimursyadah et al. [65] also identified nine HCN-producing bacteria isolates from cassava rhizosphere soil. In this study, the bacteria capable of producing HCN included Pseudomonas aeruginosa, Enterobacter sp., and Brucella intermedia.

ACC-deaminase-producing PGPR plays an essential role as 'stress modulators' by reducing ethylene levels in plants, helping to withstand various stress conditions [67]. In this study, ACC-deaminase-producing bacteria were found in species like Pseudomonas aeruginosa, Enterobacter sp., Delftia tsuruhatensis, and Klebsiella pneumoniae. ACC-deaminase (1-aminocyclopropane-1-carboxylate deaminase) in rhizobacteria can break down ACC, ethylene, especially when plants are under stress, is a precursor that can lead to negative effects. Elevated ethylene concentrations can cause chlorosis, inhibit growth, and may even result in plant death.

Previous studies have demonstrated that ACC-deaminase-producing bacteria, such as Streptomyces venezuelae, can enhance plant growth and salinity tolerance by reducing ethylene levels, reactive oxygen species, and Na+ ions [68]. Similarly, Pseudomonas putida and P. fluorescens have shown benefits for wheat plants under salt stress, leading to increased plant height, root length, biomass, and grain yield. Furthermore, ACC-deaminase enzymes in rhizobacteria have been found to improve germination rates and promote the growth of roots and shoots, helping plants tolerate salt stress [69].

According to Danish et al. [70], salinity tolerance in Camelina sativa was significantly increased by soil inoculation with ACC-deaminase-producing PGPR or the expression of the acdS gene in transgenic plants. According to a study by Heydarian et al. [71], ACC-deaminase-producing bacteria are effective in enhancing plant growth under various stress conditions, including salinity and drought. According to Maulidia [72], phosphate-solubilizing rhizobacteria help dissolve tricalcium phosphate, enhancing availability and supporting the biological efficiency of nitrogen fixation. Study [73] found that seed pre-germination treatments with rhizobacteria effectively increased vegetative growth and chili yield, particularly for species such as Azotobacter sp., Bacillus megaterium, B. coagulans, Flavobacterium sp., and Pseudomonas dimuta, which gave the most significant results.

According to Han et al. [74], the D. tsuruhatensis HR4 strain, which was isolated from the rice rhizosphere, successfully inhibited the growth of a number of plant diseases. This resulted from siderophore production, which lessened soil iron restriction. Furthermore, D. tsuruhatensis MTQ3, which was isolated from the tobacco rhizosphere, is a PGPR that is safe for the environment and can improve plant growth and antibacterial activity [75, 76]. Pseudomonas, Azotobacter, Azospirillum, Acetobacter, and Bacillus are other well-known PGPR. Pseudomonas aeruginosa is known to be one of the most powerful PGPR strains among these. According to Ghadamgahi et al. [77], Pseudomonas aeruginosa promotes plant growth by producing siderophores and solubilizing phosphate, which leads to increased mineral absorption and enhanced lateral root development in host plants.

In general, Plant Growth-Promoting Rhizobacteria (PGPR) are crucial for controlling the balance of plant hormones and forming the rhizosphere's microbial population, both of which contribute to the resilience and growth of plants. PGPR has the ability to increase the synthesis of phytohormones that are involved in root growth, stem elongation, and cell differentiation, including auxins (IAA), gibberellins, and cytokinins [78]. Application of rhizobacterial isolates to patchouli plants increased the number of branches by providing nutrients during the vegetative phase, stimulating cell division, differentiating new shoots, and increasing photosynthetic activity through phosphate solubilization [79].

In terms of PGPR interactions with rhizosphere microbial communities, root exudates produced by plants in response to PGPR colonization can also influence the dynamics of soil microbial communities, increasing the population of beneficial microorganisms, and creating a healthier rhizosphere environment [80]. Through this mechanism, PGPR not only acts as a biofertilizer that increases nutrient availability but also functions as a biostimulant and bioprotectant that strengthens the plant defense system and increases productivity. Additionally, PGPR helps create a more stable microbial community in the rhizosphere by promoting nutrient exchange and producing bioactive compounds that support the growth of symbiotic microorganisms. Some PGPR strains are known to enhance the activity of soil enzymes, such as dehydrogenases and phosphatases, which are involved in the mineralization process and improve nutrient availability for plants. In addition, PGPR can affect the soil microbiome by inducing plant systemic resistance (ISR), a defense mechanism that increases plant resistance to pathogen attack and environmental stress. Through competition for nutrients and space, as well as the generation of hydrolytic enzymes that can break down pathogen cell walls, PGPR may also reduce the number of harmful bacteria [81, 82]. Therefore, PGPR has the potential to be a solution for a more sustainable agricultural system since it not only directly promotes plant development through phytohormone synthesis but also stabilizes the soil ecology by improving microbial interactions in the rhizosphere.

Moreover, this study does not address the influence of greenhouse conditions compared to field conditions on the effectiveness of superior PGPR bacteria in enhancing onion crop yields, nor does it evaluate the long-term stability of different PGPR strains. Consequently, a more detailed and specific analysis is warranted to assess the enduring stability and efficacy of these promising bacterial strains.