Biological Control Efficacy of Indigenous Antagonistic Bacteria Isolated from the Rhizosphere of Cabbage Grown in Biofumigated Soil against Pythium aphanidermatum Damping-Off of Cucumber

Abstract

Soil biofumigation is a widely used farming practice for soil-borne disease management in vegetable crops. Soil biofumigation is the practice of the incorporation of fresh plant materials from the members of the Brassicaceae family into the soil which release antimicrobial volatile organic compounds (VOCs), namely glucosinolates and their hydrolysis products. However, the VOCs may affect non-target beneficial soil biota including microbial biocontrol agents (MBAs) due to their broad-spectrum toxicity. In this study, native antagonistic rhizobacteria were isolated from cabbage plants grown in biofumigated fields and their potential in the management of Pythium aphanidermatum cucumber damping-off was evaluated with and also without biofumigation using cabbage leaf residue. A total of 20 morphologically distinct, culturable bacterial strains were recovered from the rhizosphere soil of cabbage cultivated in a biofumigated field, with the largest fraction of the isolates belonging to the genus Bacillus. The other bacterial genera isolated were Pseudomonas, Serratia, Stenotrophomonas, Microbacterium and Priestia. Of the 20 bacterial isolates, Pseudomonas aeruginosa B1-SQU, Pseudomonas indica B2-SQU, Serratia marcescens B3-SQU and Pseudomonas brenneri B4-SQU exhibited inhibitory activity against P. aphanidermatum in a co-culture assay. The deformation and shrinkage of P. aphanidermatum hyphae due to diffusible antimicrobial compounds from these bacterial strains were witnessed through scanning electron microscopy. A biofilm was formed by these bacterial strains in vitro and they exhibited compatibility with each other; however, they had no significant plant growth promotion effect on cucumber. These bacterial strains significantly reduced damping-off in cucumber under greenhouse conditions when applied to soil singly, but failed to show a significant effect when combined with biofumigation treatment, though the bacterial population in the soil was maintained at higher levels. Soil biofumigation alone was also not effective in suppressing the damping-off of cucumber. Gas chromatography–mass spectrometry analysis revealed that 1-butanol, n-butyl butyrate and butyl acetate were the predominant VOCs in the soil amended with cabbage leaf residue. The results suggest that soil biofumigation with cabbage leaf residue has no significant effect on P. aphanidermatum under high inoculum levels; furthermore, it affects the efficiency of the bacterial antagonists in controlling damping-off in cucumber.

1. Introduction

Cucumber (Cucumis sativus L., Cucurbitaceae), cultivated widely throughout the world, accounts for 21,72,193 ha. In Oman, it is cultivated on an area of 182 ha with a production of 93,114 tons [1]. Damping-off disease, caused by Pythium aphanidermatum, an oomycete plant pathogen, is a major constraint to the production of cucumbers in greenhouses and open fields in Oman [2]. P. aphanidermatum infects the hypocotyl of the germinating seedlings and causes shrinkage, disintegration of the stem, wilting and mortality of the seedlings [3]. The pathogen persists in the infested soil for a long period of time by producing oospores, and hence the management of the disease has been very challenging. An increase in the irrigation water salinity is also known to exacerbate the problem of damping-off in cucumber [2]. Soil drenching with some synthetic chemical fungicides has been demonstrated as an effective strategy to protect plants from soil-borne pathogens including oomycetes [4,5]. However, many of these chemical fungicides are reported to cause environmental pollution and induce the development of resistance in pathogens by inducing mutations [6,7]. It has been demonstrated that a single amino acid substitution at codon 143 of the Plasmopara viticola cytochrome b gene resulted in resistance in the pathogen to quinone outside inhibitor (QoI) fungicide [8]. The systemic fungicide metalaxyl-M has been shown to be highly effective against phytopathogenic oomycetes including Pythium spp. However, several studies have documented metalaxyl resistance in many species of Pythium including P. dissotocum [9], P. irregulare [10], P. aphanidermatum, P. sulcatum, P. graminicola, P. arrhenomanes, P. vanterpolii and P. ultimum [11,12,13,14], as well as other soil-borne pathogens belonging to the class of Peronosporomycetes [15,16]. Al-Balushi et al. [17] reported the occurrence of a hymexazol-resistant strain of P. aphanidermatum in Oman. Increasing public concerns regarding the undesirable effects of synthetic chemical pesticides on the environment, including soil health, and the promotion of the development of fungicide resistance in pathogens [18] lead to the desire to use environmentally benign methods for controlling this disease. Soil solarization and biofumigation [3,19,20], biological control using antagonistic microorganisms [21,22,23,24,25] and the field application of post-harvest mushroom compost and Bacillus aryabhattai [26] have been suggested as effective and eco-friendly strategies for controlling this disease.

Soil biofumigation is defined as the “suppression of soil pests and soil-borne pathogens by volatile poisonous isothiocyanates (ITCs), released in the soil after incorporation of glucosinolate (GSL)-containing plant tissues through hydrolysis” [27]. Soil biofumigation can be achieved via the soil incorporation of fresh plant materials from members of the Brassicaceae family including cabbage, cauliflower, broccoli, kale and various mustards, which contain high concentrations of GSLs in their tissues [28,29,30]. The presence of more than 132 different GSLs (thioglucoside compounds) that vary in their chemical structures (aliphatic, aromatic or indole) have been reported in Brassica spp. [31]. The concentration of GSLs present in plant tissues and the type of products released after the hydrolysis of GSLs vary greatly among the members of Brassicaceae [32]. The GSLs of plants are hydrolyzed by the enzyme myrosinase (EC 3.2.1.147). In the intact and undamaged plant tissues, GSLs and the myrosinase enzyme are physically separated. In Brassicaceae plants, myrosinases are stored in vacuoles or in specific myrosin cells [33]. Upon tissue damage, myrosinase and GSLs come into contact and the enzyme hydrolyzes the thioglucoside bonds of glucosinolates and produces an unstable compound, namely thiohydroximate-O-sulfonate, that degrades into a number of biologically active volatile compounds, namely ITCs, thiocyanates and nitriles [34]. The ITCs, which are volatile in nature, are known to have fungicidal, insecticidal and nematicidal activities [28,35]. The toxicity of ITCs is attributed to irreversible non-specific reactions with amines and sulfur-containing groups in proteins [28]. The steps involved in the biofumigation are (i) the incorporation of fresh plant biomass to the soil, (ii) irrigating the soil to its water holding capacity, (iii) using transparent plastic film to cover the soil surface and (iv) the removal of the plastic film 3–4 weeks later and the planting of the crops after 24 h. In addition to biofumigant activity, the incorporation of fresh plant biomass is known to enhance organic matter in the soil and thus improve soil health [36], and favor the build-up of beneficial antagonistic microorganisms and the release of other biocidal compounds apart from products of GSL [37,38].

Brassicaceous plants which produce dimethyl disulphide (DMDS; C₂H₆S₂) and dimethyl sulphide [(CH₃)₂S] in the soil have also been reported to be biofumigants in controlling soil-borne fungal pathogens including Fusarium oxysporum and Verticilium dahliae [39]. Recently, Garain et al. [40] reported the effectiveness of biofumigation using Indian mustard green biomass in combination with soil solarization and the application of Trichoderma sp. against the Athelia rolfsii collar rot of Piper betle. Neubauer et al. [41] demonstrated the biofumigation capability of brassicaceous green manures. Amendments to Brassica juncea shoot tissue significantly reduced the number of microsclerotia of V. dahliae with efficiencies ranging from 69.3 to 81.3%. Mattner et al. [42] reported the suppression of Phytophthora cactorum and other soil-borne pathogens of strawberry by volatile compounds released from the macerated roots of Brassica rapa and B. napus. Wang et al. [43] reported that rapeseed meal biofumigation suppressed the incidence of the Phytophthora blight of pepper plants. Morales-Rodriguez et al. [33] demonstrated that biofumigation with BioFence (a commercial Brassica carinata sold as pellets) reduced the inoculum density of P. cinnamomi and offered protection to Quercus cerris from root infection. In addition to the Brassica species, members of the plant family Alliaceae such as onion, garlic and leek are used for biofumigation to control soil-borne pathogens. The volatile compounds thiosulfinates and disulfides released by these plants upon tissue damage have been reported to possess antimicrobial activities [44,45]. Soil temperature plays an important role in biofumigation. It determines the concentration and volatility of ITCs produced during the process of biofumigation [46] and the activity of myrosinase [47]. Morales-Rodriguez et al. [33] reported that the efficacy of BioFence in restricting the growth of P. cinnamomi was maximum at 15 °C and reduced at 25 °C.

In soil, GSLs and ITCs are short-lived and quickly diminish within a few days after treatment and hence the environmental risks associated with biofumigation are low [48]. However, the biofumigants may affect non-target beneficial soil biota such as microbial biological control agents (BCAs) or other pest antagonists due to their broad-spectrum toxicity [37,49]. For instance, Henderson et al. [50] reported that the incorporation of Brassica carinata seed meal in soil disrupted the efficacy of the entomopathogenic nematodes Steinernema feltiae and S. riobrave in controlling root-knot nematode (Meloidogyne chitwoodi) and Colorado potato beetle (Leptinotarsa decemlineata) in potato. The objectives of this work were to isolate native antagonistic rhizobacteria from biofumigated soil and to evaluate their efficacy in controlling the damping-off of cucumber either individually or in combination with biofumigation; specifically, the following were studied: (i) the isolation and characterization of bacteria from the rhizosphere of cabbage cultivated in biofumigated fields; (ii) the in vitro antagonistic activity of the rhizobacterial strains against Pythium aphanidermatum; (iii) morphological changes in the mycelium of P. aphanidermatum due to the antagonistic effect of bacteria; (iv) the plant growth promotion of selected antagonistic bacterial strains; (v) the ability of the antagonistic bacterial strains to form biofilm; (vi) the efficacy of the application of antagonistic bacteria with and without soil biofumigation on Pythium aphanidermatum-induced damping-off of cucumber under greenhouse conditions and (vii) the profiling of the VOCs of biofumigated soil.

2. Materials and Methods

2.1. Soil Sample Collection and Isolation of Bacteria

Rhizosphere soil samples were collected from cabbage grown in Nehad Agronomy Services LLC located in Suwaiq, Oman (23°50′58″ N 57°26′19″ E). This farm contains sandy loam soil, and biofumigation using cabbage leaf residue has been practiced for the last 6 years. For the collection of rhizospheric soil, roots from a few cabbage plants growing in the biofumigated soil were excised and samples were collected including the soil particles adhering to the roots. The soil samples were transferred to the laboratory within 24 h and used for the isolation of bacteria. The soil sample (1 g) was suspended in 99 mL of sterile distilled water (SDW) in a 250 mL conical flask and mixed thoroughly using a magnetic stirrer at room temperature (25 ± 2 °C) for 1 h. Subsequently, the soil suspension was serially diluted with SDW and 100 μL of each dilution (10⁻⁴, 10⁻⁵ and 10⁻⁶) was plated on nutrient agar (NA) medium (Oxoid Ltd., Basingstoke, UK) [51]. The plates were incubated at 30 °C for 24–48 h and the bacterial colonies developing on the plates were transferred to new plates.

2.2. Pythium aphanidermatum

A virulent culture of P. aphanidermatum strain Sala1 (GenBank accession number: ON113866) isolated from a damping-off infected cucumber seedling was obtained from the microbial culture collections of the Department of Plant Sciences, Sultan Qaboos University. The culture was maintained on potato dextrose agar (PDA; Oxoid Ltd., UK) medium at 4 °C.

2.3. Antagonistic Activity of Bacterial Isolates against P. aphanidermatum

Bacterial isolates were screened for antagonistic activity against P. aphanidermatum in dual culture. A mycelial plug of P. aphanidermatum (6 mm diameter) cut from a 3-day-old PDA culture of the pathogen was placed on one end of the PDA plate (9 cm diameter), approximately 1 cm away from the margin. Test bacterial isolate was streaked on the opposite side of the PDA plate approximately 1 cm away from the periphery. The Petri dish inoculated with a mycelial disc of P. aphanidermatum alone served as the control. Three replicated plates were used for each bacterial isolate. The plates were incubated for 3–5 days at room temperature (25 ± 2 °C) or until the mycelium of P. aphanidermatum covered the entire plate in the control. The inhibition zone (the distance between the bacterium and the mycelium of the pathogen) was recorded [51].

2.4. Morphological Changes in the Mycelium of P. aphanidermatum Due to Antagonistic Effect of Bacteria via Scanning Electron Microscopy

Mycelial discs of 0.5 cm diameter were taken from the periphery of P. aphanidermatum at the inhibition zone in dual culture plates of the effective bacterial strains. Samples were prepared for the scanning electron microscopy according to the method described by Goldstein et al. [52] and were observed with a JEOL scanning electron microscope (Model JSM-7800F; JEOL USA, Inc., Peabody, MA, USA).

2.5. Identification of Antagonistic Bacterial Isolates

2.5.1. 16S rRNA Gene Sequencing

The DNA from the bacterial isolates was extracted using a commercial DNA extraction kit (foodproof StarPrep Two kit; BIOTECON Diagnostics GmbH, Potsdam, Germany) in accordance with the manufacturer’s instructions. The bacterial 16S rRNA gene was amplified using the universal primers 27F, 1429R and 534R [53,54] in a thermal cycler (Veriti 96-well Thermal Cycler; Applied Biosystems, Singapore). The PCR mixture comprised 1 μL of DNA (100 ng μL⁻¹), 1 μL of each primer (20 pmol μL⁻¹), 22 μL of water and a PuReTaq Ready-To-Go PCR bead (GE Healthcare, Hatfield, UK) for 25 μL total reaction volume. The PCR reaction conditions were as follows: initial denaturation for 2 min at 95 °C, followed by 35 cycles of denaturation for 30 s at 95 °C, annealing for 30 s at 54 °C, DNA extension for 60 min at 72 °C and a final extension for 10 min at 72 °C [51]. The PCR products (5 μL aliquots) were analyzed via electrophoresis on a 1% agarose gel followed by ethidium bromide (10 mg mL⁻¹) staining and sequencing (Macrogen, Seoul, Republic of Korea). The bacterial isolates were identified by comparing the 16S rDNA sequences with those existing in the GenBank (http://www.ncbi.nlm.nih.gov) database using the BLASTN program of the NCBI.

2.5.2. Plant Growth Promotion of Bacterial Strains

The plant-growth-promoting ability of the antagonistic bacterial strains was tested using the paper towel method. Cucumber seeds (cv. Jabbar, F1; US Agriseeds, Woodland, CA, USA) were soaked in the bacterial suspension (1 × 10⁸ cfu/mL) for 3 h at room temperature (25 ± 2 °C). The treated seeds were then placed in between two moist germination papers, rolled and incubated at 25 ± 2 °C. Sterile distilled water was used to treat seeds in the control. After 15 days of incubation, the % germination, root length and shoot length of the seedlings were recorded. The formula of vigor index = (mean root length + mean shoot length) × percentage germination was used [55].

2.5.3. Biofilm Forming Ability of Bacterial Isolates

The ability of the rhizobacterial strains to form biofilm was tested as described by O’Toole [56] with some modifications. The bacterial strains were cultured in 100 mL of nutrient broth (NB) at 30 °C for 24 h on a shaker at 180 rpm. Ten mL of bacterial suspension was transferred to a sterile 15-mL tube, and the optical density (OD) was adjusted to 0.2 at 600 nm with NB. The bacterial culture was then centrifuged at 3000 rpm at 4 °C for 10 min and the pellet containing bacteria was suspended in 10 mL of fresh NB medium. The bacterial suspension was added to the wells of a 96-well polystyrene plate (200 μL/well) and incubated at 28 °C for 24 h. Uninoculated NB, processed in the same way, served as the control. Ten replications were maintained for each bacterium. The bacterial culture was then carefully removed from the wells and the wells were washed with sterile distilled water (SDW) to remove unattached bacterial cells. Crystal violet (200 μL; 0.1% w/v; in distilled water) was added to the wells. After 15 min of incubation at room temperature (25 ± 2 °C), the wells were washed 5 times with SDW and air-dried. Ethanol (200 μL; 95%) was added to each well and incubated at 25 °C for 30 min. The absorbance of the solution was measured at 600 nm using a microplate reader (Model: MULTISKAN GO; ThermoFisher Scientific, Vantaa, Finland).

2.5.4. Testing Compatibility between Bacterial Isolates

The in vitro cross-streak method was used for testing the rhizosphere bacterial strain compatibility. A rhizosphere bacterial isolate was streaked parallelly on an NA plate and the other test bacterial strains were streaked perpendicular to the first one; the plates were incubated at 30 °C for 3 days. The bacterial growths at the interjunction were observed. The merger of bacterial growths indicates compatibility [51].

2.5.5. Efficacy of Antagonistic Bacteria and Soil Biofumigation on Pythium aphanidermatum Damping-Off of Cucumber

Pot culture experiments were conducted in a glass house at the Agricultural Experiment Station, SQU, to study the effect of biofumigation and biocontrol agents on the incidence of damping-off of cucumber caused by P. aphanidermatum. Cucumber cultivar Jabbar, F1 (US Agriseeds, USA), was used for pot experiments. The experiment was laid out in a completely randomized design with 12 treatments including the untreated control. The treatments included (1) Pseudomonas aeruginosa B1-SQU; (2) Pseudomonas indica B2-SQU; (3) Serratia marcescens B3-SQU; (4) Pseudomonas brenneri B4-SQU; (5) P. aeruginosa B1-SQU + biofumigation; (6) P. indica B2-SQU + biofumigation; (7) S. marcescens B3-SQU + biofumigation; (8) P. brenneri B4-SQU + biofumigation; (9) biofumigation + P. aphanidermatum; (10) P. aphanidermatum alone (infected control); (11) biofumigation alone and (12) untreated control.

The antagonistic bacteria were cultured in nutrient broth at 30 °C on a shaker for 48 h. The OD of the bacterial suspension was measured using a spectrophotometer and 10 mL of the bacterial suspension (9 × 10⁸ CFU mL⁻¹) was added to each pot and mixed well.

To produce an inoculum of the pathogen, P. aphanidermatum was grown on sterilized barley grains (150 g barley grain + 100 mL distilled water) at room temperature (25 ± 2 °C) for 2 weeks. The barley grains fully colonized with P. aphanidermatum were mixed with the potting soil (Bulrush Horticulture Ltd., Magherafelt, Ireland, UK) 3 days after the application of antagonistic bacteria at the rate of 10 g/pot.

For the biofumigation treatment, fresh cabbage leaves were cut into small pieces, added to the soil (10 g wet biomass/pot filled with 500 g sterilized soil) [57] 3 days after the application of antagonistic bacteria, thoroughly mixed and irrigated to field capacity. The soil was covered with a transparent 0.2 mm thick plastic sheet for 21 days and then removed. Three days after completion of the biofumigation treatment, cucumber seeds (cv. Jabbar, F1; US Agriseeds, USA) were sown in pots at the rate of 10 seeds/pot.

The plants were maintained at 25/20 °C (day/night), and 80% relative humidity. Damping-off disease incidence (%) was recorded 15 days after sowing. Each treatment was replicated five times.

The antagonistic bacterial population in the soil (Treatments 1–8) was determined at three time intervals, viz., 0 (before biofumigation), 27 (after biofumigation and the day of planting) and 42 days (the day of termination of the experiment) using NA medium and the serial dilution technique. Since these antagonistic bacterial strains were not marker strains, the total bacterial population in the NA medium was counted. A few colonies were randomly picked and confirmed via molecular analysis. Each treatment was replicated three times.

2.6. Analysis of VOC Emission from Soil Amended with Cabbage Leaf Residue

Cabbage plants were grown in pots under greenhouse conditions and the leaves were harvested before the flowering stage and cut into small pieces. Sterilized soil was taken in a solid-phase micro extraction vial of 50 mL capacity, amended with cabbage leaf tissues at the 5% level (wt/wt) and sealed with a polypropylene hole cap and PTFE coated silicone septa. SDW was added into the tube to the final moisture content of 10% (approximately field capacity) and incubated at room temperature (25 ± 2 °C) for 72 h. VOCs released by the rhizosphere soil in the headspace were extracted and analyzed via GC-MS. The GC-MS system used was Shimadzu GC-2010 Plus (Shimadzu), fitted with an Rtx-5MS capillary column (30 m × 0.25 mm, 0.25 μm) attached to a GCMS-QP2010 ULTRA MS as described by Al-Rashdi et al. [25]. The identification of VOCs was conducted by comparing retention time and mass spectra with those of the NIST (National Institute of Standards and Technology) 2011 v.2.3 and Wiley 9th edition mass spectrum libraries.

2.7. Statistical Analysis

SAS v8 software (SAS Institute, Cary, NC, USA) was employed for the statistical analysis of the data generated in this study. Analysis of variance (ANOVA) was performed using the general linear model. Duncan’s multiple range test (DMRT; p < 0.05) was used to determine the differences between treatment means.

3. Results

3.1. Isolation and Characterization of Antagonistic Rhizobacteria

A total of 20 morphologically different bacterial isolates belonging to six genera (Bacillus, Pseudomonas, Serratia, Stenotrophomonas, Microbacterium and Priestia) were obtained from the rhizosphere of cabbage plants grown in biofumigated soil. Among them, Bacillus was the most dominant genus (60%), followed by Pseudomonas (15%), Stenotrophomonas (10%), Serratia (5%), Microbacterium (5%) and Priestia (5%). The GenBank accession numbers of the 16S rRNA gene sequences of the bacterial isolates obtained in this study are given in

Table 1. Molecular identification of bacterial isolates from the rhizosphere of cabbage plants grown in biofumigated soil via sequencing of 16S rRNA gene.

Bacterial Isolate	Sequence Length (bp)	Closest Related Microorganism in GenBank Database	Similarity %	GenBank Accession Number
B1-SQU	412	Pseudomonas aeruginosa	100.00%	ON738574
B2-SQU	566	Pseudomonas indica	100.00%	ON738576
B3-SQU	597	Serratia marcescens	100.00%	OP837487
B4-SQU	385	Pseudomonas brenneri	100.00%	ON738575
B5-SQU	387	Bacillus oceanisediminis	100.00%	ON710971
B6-SQU	507	Bacillus endophyticus	100.00%	ON710992
B7-SQU	569	Bacillus oceanisediminis	100.00%	ON710993
B8-SQU	532	Bacillus altitudinis	100.00%	ON710994
B10-SQU	924	Priestia megaterium	100.00%	ON713424
B11-SQU	973	Bacillus aryabhattai	99.90%	ON713423
B12-SQU	967	Bacillus endophyticus	100.00%	ON713425
B13-SQU	934	Microbacterium paraoxydans	100.00%	ON713453
B14-SQU	824	Bacillus korlensis	100.00%	ON713456
B15-SQU	869	Bacillus firmus	100.00%	ON713454
B16-SQU	956	Bacillus endophyticus	99.79%	ON713457
B17-SQU	871	Bacillus oceanisediminis	99.89%	ON713452
B18-SQU	818	Stenotrophomonas maltophilia	100.00%	ON713458
B19-SQU	915	Stenotrophomonas maltophilia	100.00%	ON713455
B20-SQU	988	Bacillus firmus	99.90%	ON713467
B21-SQU	684	Bacillus firmus	100.00%	ON713466

.

3.2. In Vitro Anti-Oomycete Activity of Rhizobacterial Isolates

The rhizobacterial isolates were tested for their inhibitory activity against P. aphanidermatum under laboratory conditions. Of the 20 bacterial isolates, four isolates, viz., Pseudomonas indica B2-SQU, Serratia marcescens B3-SQU, Pseudomonas aeruginosa B1-SQU and Pseudomonas brenneri B4-SQU, inhibited the growth of P. aphanidermatum and produced clear inhibition zones (Figure 1;

Table 2. In vitro mycelial growth inhibition of Pythium aphanidermatum by bacterial isolates from the rhizosphere of cabbage cultivated in biofumigated soil.

Bacterial Isolate	Inhibition Zone (mm)
Pseudomonas aeruginosa B1-SQU	9.8 b
Pseudomonas indica B2-SQU	14.0 a
Serratia marcescens B3-SQU	11.8 ab
Pseudomonas brenneri B4-SQU	6.2 c
Control	0 d

Data are the means of five replications. Means followed by different letters in the column are significantly different from each other at p < 0.05, according to DMRT.

). None of the other 16 bacterial isolates inhibited the mycelial growth of P. aphanidermatum.

3.3. Morphological Changes in P. aphanidermatum Due to Antagonistic Effect of Rhizobacteria

The scanning electron microscopic imaging of P. aphanidermatum at the inhibition zone of culture assay plates revealed a severe damage to hyphal structures due to the effect of antagonistic bacteria (Figure 2). Shriveling, twisting and distortion were commonly observed in P. aphanidermatum hyphae when co-cultivated with antagonistic bacteria. P. aphanidermatum hyphae in the control were healthy, branched and tubular with a smooth surface.

3.4. Effect of Bacterial Isolates on Growth of Cucumber

No significant (p < 0.05) differences in the seed germination percentage, shoot length or root length of seedlings were observed between the control and rhizobacteria-treated cucumber seedlings in a rolled paper towel assay (

Table 3. Efficacy of seed treatment with antagonistic bacterial isolates from the rhizosphere of cabbage on seed germination and seedling vigor of cucumber under laboratory conditions.

Treatment	% Germination	Root Length (cm)	Shoot Length (cm)	Vigor Index
Pseudomonas aeruginosa B1-SQU	85.2 a	12.3 a	7.7 a	1719 a
Pseudomonas indica B2-SQU	88.9 a	13.1 a	9.0 a	2002 a
Serratia marcescens B3-SQU	81.5 a	10.3 a	6.8 a	1416 a
Pseudomonas brenneri B4-SQU	57.4 a	8.3 a	5.6 a	828 a
Control	72.2 a	10.4 a	7.4 a	1328 a

Data are means of five replications. Means followed by the same letter in the column are not significantly different from each other at p < 0.05, according to DMRT.

).

3.5. Biofilm Formation by Bacterial Isolates

All four bacterial isolates were shown to form biofilm in the in vitro microtiter plate assay; among them, Pseudomonas indica B2-SQU formed the highest density of biofilm as indicated by a higher OD value (A₆₀₀ 1.2) compared to the other rhizobacterial isolates tested in this study, but they were not statistically different from one another (

Table 4. Biofilm forming ability of bacterial isolates obtained from the rhizosphere of cabbage grown in biofumigated soil.

Bacterial Strain	OD at 600 nm
Pseudomonas aeruginosa B1-SQU	0.97 a
Pseudomonas indica B2-SQU	1.21 a
Serratia marcescens B3-SQU	0.91 a
Pseudomonas brenneri B4-SQU	1.05 a
Control	0.42 b

Data are the means of 10 replications. Means followed by different letters in the column are significantly different from each other at p < 0.05, according to DMRT.

). All were higher than the control.

3.6. Compatibility between Bacterial Antagonists

The compatibility between the bacterial isolates from cabbage rhizosphere was evaluated using the in vitro cross-streak method. The results showed that the bacterial strains were compatible as no inhibition was observed in the interjunction. The compatibility of P. brenneri B4-SQU with other bacterial isolates is shown in Figure 3.

3.7. Damping-Off of Cucumber Biocontrol

A significant reduction in the incidence of damping-off over the Pythium-infected control was observed in pots treated with BCAs (

Table 5. Biocontrol efficacy of bacteria grown in biofumigated soil against Pythium aphanidermatum-induced damping-off of cucumber.

Treatment	Seedling Mortality (%)
P. aeruginosa B1-SQU + P. aphanidermatum	22 (21.6) de
P. indica B2-SQU + P. aphanidermatum	42 (39.6) cd
S. marcescens B3-SQU + P. aphanidermatum	52 (46.1) bcd
P. brenneri B4-SQU + P. aphanidermatum	40 (36.0) cd
P. aeruginosa B1-SQU + P. aphanidermatum + biofumigation	100 (90.0) a
P. indica B2-SQU + P. aphanidermatum + biofumigation	72 (59.3) abc
S. marcescens B3-SQU + P. aphanidermatum + biofumigation	96 (82.6) a
P. brenneri B4-SQU + P. aphanidermatum + biofumigation	86 (76.1) ab
Biofumigation + P. aphanidermatum	100 (90.0) a
P. aphanidermatum alone (infected control)	100 (90.0) a
Biofumigation alone (no P. aphanidermatum)	10 (16.3) de
Untreated control	0 e

Data are means of 5 replications. Values in the brackets are arcsine transformed values. Means followed by different letters in the column are significantly different from each other at p < 0.05, according to DMRT.

). The lowest seedling mortality was recorded in pots treated with P. aeruginosa B1-SQU (22%), followed by P. brenneri B4-SQU (40%), P. indica B2-SQU (42%) and S. marcescens B3-SQU (52%), as compared to 100% mortality in the infected control. However, the effectiveness of BCAs was significantly reduced when combined with the biofumigation treatment. P. aeruginosa B1-SQU completely lost its effectiveness in suppressing the damping-off of cucumber when combined with biofumigation and showed 100% seedling mortality, whereas P. indica B2-SQU, S. marcescens B3-SQU and P. brenneri B4-SQU recorded 72, 96 and 86% seedling mortality, respectively. Under greenhouse conditions, the damping-off of cucumber was not controlled by soil biofumigation with cabbage leaf residue.

3.8. Population of Antagonistic Rhizobacterial Strains in Soil

Antagonistic bacterial strain populations in the soil were assessed at 0, 27 and 42 days after treatment. The results revealed that the population of introduced antagonistic bacteria generally decreased with time after application; however, a significantly higher population of P. indica was observed in the biofumigated soil even at 42 days after application (

Table 6. Population of antagonistic bacterial isolates in the rhizosphere of cucumber after application.

Treatment	Rhizosphere Population Density (1 × 105 cfu/g Soil)
	Days after Application
	27	42
Pseudomonas aeruginosa B1-SQU	138 b	49 ab
Pseudomonas indica B2-SQU	228 ab	30 b
Serratia marcescens B3-SQU	189 ab	57 ab
Pseudomonas brenneri B4-SQU	149 b	52 ab
Pseudomonas aeruginosa B1-SQU + biofumigation	97 b	12 b
Pseudomonas indica B2-SQU + biofumigation	483 a	132 a
Serratia marcescens B3-SQU + biofumigation	125 b	25 b
Pseudomonas brenneri B4-SQU + biofumigation	213 ab	47 b

The population of antagonistic bacteria in the rhizosphere of cucumber in the pots was determined 27 and 42 days after application with 6.5 × 10⁷ CFU/g soil. Data are means of 3 replications. Means within a column followed by different letters are significantly different at p < 0.05, according to DMRT.

).

3.9. VOC Emission by Biofumigated Soil

Twenty-five volatile compounds were identified in the headspace of biofumigated soil via GC-MS. 1-Butanol (18.2%; RT 1.946), acetic acid, butyl ester (butyl acetate) (13.29%; RT 3.096) and Butanoic acid, butyl ester (n-butyl butyrate) (16.09%; RT 6.531) were identified as the major components in the soil amended with cabbage leaf residue, whereas Silanediol dimethyl- (19.7%; RT 1.98) was the predominant compound in the control soil (Figure 4;

Table 7. Volatile organic compounds detected in the control soil.

S.No.	Compound	Retention Time (min)	Area%
1	l-Alanine ethylamide, (S)-	1.38	3.62
2	Acetone	1.533	4.61
3	Borane carbonyl	1.587	9.38
4	Boric acid	1.67	2.71
5	n-Hexane	1.72	4.59
6	Trichloromethane	1.791	4.35
7	Ethane, 1,2-dichloro-	1.91	1.92
8	Silanediol, dimethyl-	1.98	19.7
9	Heptane	2.111	1.07
10	Hexanal	2.935	1.84
11	Cyclotrisiloxane, hexamethyl-	3.109	4.63
12	2-Hexanone, 3,4-dimethyl-	3.683	1.27
13	Oxime-, methoxy-phenyl-_	4.256	1.73
14	Cyclotetrasiloxane, octamethyl-	6.488	5.36
15	Nonanal	9.611	4.85
16	Cyclopentasiloxane, decamethyl-	11.13	6.57
17	3-Butynoic acid	21.843	1.42

and

Table 8. Volatile organic compounds detected in the biofumigated soil.

S.No.	Compound	Retention Time (min)	Area%
1	Carbon dioxide	1.416	2.92
2	Methanethiol	1.494	7.12
3	Isopropyl Alcohol	1.547	6.11
4	Ethanol, 2-chloro-, acetate	1.596	1.66
5	Carbon disulfide	1.642	3.59
6	Ethyl Acetate	1.777	2.45
7	1-Butanol	1.946	18.25
8	Methyl thiolacetate	2.156	2.48
9	n-Propyl acetate	2.215	0.27
10	Butanoic acid, methyl ester	2.273	1.03
11	Disulfide, dimethyl	2.501	3.13
12	Methallyl cyanide	2.595	0.59
13	Butanoic acid	2.763	3.77
14	Butanoic acid, ethyl ester	2.942	4.95
15	Acetic acid, butyl ester	3.096	13.29
16	Isopropyl butyrate	3.453	2.84
17	Butanethioic acid, S-methyl ester	4.249	0.38
18	Butanoic acid, propyl ester	4.365	0.42
19	Propanoic acid, butyl ester	4.557	1.55
20	Propanoic acid, 2-methyl-, butyl ester	5.496	0.3
21	Dimethyl trisulfide	6.043	1.97
22	Butanoic acid, butyl ester	6.531	16.09
23	D-Limonene	7.49	1.43
24	Butanenitrile, 4-(methylthio)-	9.066	0.69
25	Pentanenitrile, 5-(methylthio)-	12.711	2.72

).

4. Discussion

Biological control is a preferred option globally for the management of soil-borne vegetable crop diseases including Pythium diseases because of its environmentally friendly, cost-effect and user-friendly features. Many commercial biopesticide products based on highly efficient strains of antagonistic microorganisms are available worldwide for field application on a larger scale [58]. However, the successful biocontrol of plant diseases is dependent on the fitness level of the microbial biocontrol agents in the soil. Hence, the search is centered on identifying indigenous antagonistic bacterial strains that are suitable to the local environmental conditions such as saline soil, high soil temperature, etc.

In this study, 20 morphologically different bacteria were isolated from the rhizosphere of cabbage plants grown in biofumigated soil. Bacillus was identified as being the most predominant genus, followed by Pseudomonas. The tolerance of Bacillus spp. and Pseudomonas spp. to abiotic stresses including high temperature, salinity and drought has been well-documented [59,60]. Bacillus spp. abundance in the biofumigated soil can be attributed to their ability to form endospores which are dormant resistant structures capable of surviving under extreme environmental conditions [61]. Out of 20 bacterial isolates screened, Pseudomonas indica B2-SQU, Serratia marcescens B3-SQU, Pseudomonas aeruginosa B1-SQU and Pseudomonas brenneri B4-SQU exhibited antagonistic activity against P. aphanidermatum. These bacterial isolates produced a clear zone of inhibition in the in vitro culture assay. The diffusion of antimicrobial metabolites released by the antagonistic bacteria into the agar medium explains the inhibition zone [62]. The differences in the level of inhibition of P. aphanidermatum between the bacterial isolates in the present study can be attributed to the quantity and toxic nature of the inhibitory compounds produced by these bacterial strains. Several strains of P. indica [63], S. marcescens [64], P. aeruginosa [23,24] and P. brenneri [65] have been previously reported as antagonistic microorganisms against phytopathogenic fungi and oomycetes and used as BCAs for the control of various soil-borne diseases of crops.

The inhibitory effect of the bacterial isolates was further confirmed via the examination of P. aphanidermatum hyphae at the inhibition zone using SEM. P. aphanidermatum hyphae when co-cultivated with the antagonistic bacterial strains showed morphological abnormalities including shriveling, twisting and distortion. Similar observations were reported by Halo et al. [66] when P. aphanidermatum was co-cultivated with Aspergillus terreus isolate 9F isolated from the roots of Tephrosia apollinea. Likewise, shrinkage and twisting were reported by Al-Daghari et al. [23] when P. aphanidermatum was co-cultivated with Pseudomonas aeruginosa AT3 and P. resinovorans B11. The shrinkage of pathogen hyphae might be due to the leakage of cell contents by the action of the bacterial metabolites on the cell membrane [67]. A study performed by Troppens et al. [68] revealed that 2, 4-diacetylphloroglucinol (DAPG), a secondary metabolite produced by Pseudomonas fluorescens, inhibited Neurospora crassa growth and conidial germination by inducing morphological changes in mitochondria, as well as caused the rapid loss of mitochondrial membrane potential.

All four tested bacterial isolates in this study were positive for biofilm formation in vitro. P. indica B2-SQU formed the maximum level of biofilm compared to the other rhizobacterial isolates. Several reports described the biofilm forming potential of bacterial and yeast biocontrol agents [25,69,70,71]. The biofilm is known to enhance the resistance of bacteria to various environmental stresses including alterations in pH, antimicrobial agents and ultraviolet (UV) light and help them to colonize the rhizosphere of plant roots [72]. The biofilm offers protection from pathogens to the host plants because of competition for essential nutrients on the plant surface [73]. Furthermore, a few yeasts in the biofilm have been reported to suppress fungal pathogens via the production of cell wall lytic enzymes [74]. Bais et al. [69] demonstrated that Bacillus subtilis 6501 showed antagonistic activity against Pseudomonas syringae pv. tomato which formed biofilm, whereas the non-antagonistic strain B. subtilis M1 did not form biofilm. Haggag and Timmusk [70] reported that biofilm-forming Paenibacillus polymyxa offered protection against Aspergillus niger-induced peanut crown rot disease. The results of the present study suggest that biofilm formation by these rhizospheric bacterial strains could be one of the mechanisms that contribute to damping-off suppression in cucumber.

In the cross-streak assay in the present study, the growth of all four bacteria merged at the intersections suggesting that these bacterial strains are compatible with each other. Mixtures of microbial biological control agents have been shown by other studies to offer better disease control due to different modes of action than the application of single isolates [51,75,76,77,78,79]. For example, Al-Hussini et al. [51] found that mixtures of Bacillus cereus and Exiguobacterium indicum were superior in controlling P. aphanidermatum damping-off compared to individual antagonists in tomato. Similarly, the combined application of Pseudomonas fluorescens strain F113 (2,4-diacetylphloroglucinol producer) and Stenotrophomonas maltophilia strain W81 (lytic enzyme producer) offered better protection from damping-off in sugar beet than with single bacterial inoculations of either strain [75]. The combined application of Rhizophagus intraradices and Bacillus pumilus INR7 effectively controlled Rhizoctonia solani root rot of common bean [76]. Actinobacteria mixtures containing Streptomyces africanus KAI-32 + S. coelicolor KAI-90 + S. griseus strains CAI-24, CAI-121 and CAI-127 or a mixture of S. griseus CAI-127 and S. africanus KAI-32 were found to be effective in the control of chickpea wilt incited by Fusarium oxysporum f. sp. ciceri [78]. In another study, the consortium of Trichoderma harzianum and Pseudomonas aeruginosa significantly reduced the Fusarium wilt severity in banana [79]. Liu et al. [77], while evaluating the potential of Bacillus altitudinis and B. velezensis for the biological control of damping-off of cucumber (Pythium ultimum), damping-off of pepper (Rhizoctonia solani), bacterial speck of tomato (Pseudomonas syringae pv. tomato) and bacterial spot of tomato (Xanthomonas axonopodis pv. vesicatoria), reported that the levels of disease suppression were higher with bacterial mixtures than with individual bacterial antagonists. The compatibility among the rhizosphere bacterial strains used in this study makes them ideal candidates to prepare antagonist mixtures for improved efficacy.

The results of the in vitro plant-growth promotion test in this study revealed that the treatment of cucumber seeds with the BCAs had no significant effect on seed germination or seedling vigor. The role of rhizosphere bacteria in plant growth promotion and disease control has been well-documented [80,81]. Several mechanisms including phosphate solubilization and the production of indole acetic acid are employed by rhizobacteria to promote plant growth [81], while the production of antibiotics, siderophores, hydrogen cyanide (HCN) and other antimicrobial compounds such as 2,4-diacetylphloroglucinol (DAPG) and phenazine derivatives are involved in the suppression of phytopathogens [82]. However, a few studies have reported the suppression of plant growth due to the bacterization of seeds with rhizobacteria, probably due to the secretion of phytotoxic metabolites by the bacteria. For example, Al-Hussini et al. [51] found that seed treatment with Klebsiella oxytoca strain D1/3 reduced the shoot length and root length of tomato seedlings and seedling vigor compared to the control. In this study, seed bacterization with the selected rhizobacterial strains had no detrimental effect on seed germination or the growth of cucumber seedlings, demonstrating the safeness of these bacterial isolates for use in agriculture.

The results of the pot culture experiments revealed that there was a significant reduction in the damping-off incidence in cucumber after soil application of all four bacterial strains as compared to the infected control. P. aeruginosa B1-SQU exhibited the highest efficacy with a 78% reduction in damping-off incidence as compared to the infected control. However, when combined with the biofumigation treatment, no significant (p ≤ 0.05) reduction in the damping-off disease incidence was observed. Soil biofumigation via the incorporation of cabbage leaf residue also had no significant effect on the damping-off of cucumber. Although soil biofumigation has been reported to suppress several genera of soil-borne pathogens including P. aphanidermatum [19,40], our studies showed no significant reduction in the incidence of damping-off of cucumber. Davis et al. [83] reported similar findings, and found that the incorporation of Brassica napus var napus in soil had no effect on Verticillium dahliae population. Similarly, Hartz et al. [84] reported that biofumigation with Brassica juncea had no significant suppressive effects on the population of Verticillium dahliae or Fusarium spp. in soil and on the disease incidence in tomato. The variations in the GSL content among Brassica spp. [85], the partial conversion of GSLs to ITCs and inefficient field management practices [84] have been attributed to the inconsistent performance of soil biofumigation on plant disease suppression. Several other factors including the timing of the incorporation of GSL-containing plant residues in soil, the timing of the release of GSLs coinciding with the susceptible stage of the pathogens, myrosinase enzyme activity, losses in antimicrobial GSLs from soil due to evaporation and leaching and the degradation of GSL by soil microbiota also determine the efficiency of biofumigation [28]. The ineffectiveness of biofumigation with cabbage leaf residues in controlling the damping-off of cucumber in this study can be attributed to the high inoculum level of P. aphanidermatum in the soil, the tolerance of the pathogenic strain to ITCs or both of them working together [86,87].

The survival of antagonistic bacteria in the soil after inoculation is crucial for effective biocontrol. Martin et al. [88] while studying soil population density relationships between Laetisaria arvalis and Pythium ultimum damping-off in table beet reported that the decrease in disease incidence was linearly related to an increasing population density of L. arvalis in the soil. Several other factors may influence the population densities of BCAs. For example, Berger et al. [89] while studying the effect of root exudates of Daphne (Daphne blagayana) plants on Bacillus subtilis reported that the poor biocontrol activity of B. subtilis was because the Daphne roots released inhibitory compounds. The results of the current study revealed that the population of antagonistic bacterial strains gradually decreased in all of the treatments with the increase in crop age; however, some of the bacterial populations remained relatively high even 42 days after application. The population of P. indica B2-SQU in biofumigated soil was higher compared to other bacterial strains, suggesting the tolerance of this isolate to ITCs and other toxic volatiles released during the biofumigation process.

Solid-phase microextraction-gas chromatography/mass spectrometry (SPME-GC/MS) analysis of the untreated control soil identified 17 compounds, of which Silanediol dimethyl- was the predominant compound. On the contrary, 25 compounds were detected in the soil amended with cabbage leaf residue; among them, 1-butanol, Butanoic acid, butyl ester (n-butyl butyrate) and Acetic acid, butyl ester (butyl acetate) were detected as being the major components. Several studies have reported that organic amendments can affect VOC emissions from soil [90]. VOCs are odorous, secondary metabolites with low molecular weight (<300 Da), low boiling point, high vapor pressure and a lipophilic moiety [91]. These characteristics enable VOCs to diffuse well through rhizosphere soil and air [91]. The type, quality and nutrient contents of the organic matter influence the composition of VOCs [92]. For example, methanol was reported to be the most abundant compound in Mediterranean soils [93], whereas acetone was found as a major compound after amendment with straw [94]. The soil microorganisms contribute to VOC emissions from soil through different processes including the degradation of sugars, amino acids, fatty acids, alcoholic fermentation and sulfur reduction [95]. The VOCs released by microorganisms belong to different chemical classes including alcohols, alkenes, benzenoids, ketones, sulfides and terpenes [96,97,98,99]. The amount and composition of VOCs depend on the microbial communities in the soil [100]. Abis et al. [101] reported that reduced levels of microbial diversity in soil induce larger VOC emissions from soils; however, the number of different VOCs emitted decreased.

Butanoic acid, butyl ester has been reported as one of the volatile compounds of biocontrol agent Trichoderma koningiopsis VM115 [102]. Li et al. [103] reported the production of butyl acetate by Ceratocystis fimbriata in the biological control of Penicillium digitatum and Monilinia fructicola. The inhibition of mycelial growth and spore formation in Colletotrichum gloeosporioides by butyl acetate produced in a co-culture system of Bacillus subtilis and Trichoderma sp. has been demonstrated [104]. The inhibition of laccase enzyme activity which is necessary for melanin synthesis and pathogenicity has been suggested as the mechanism of action of butyl acetate on C. gloeosporioides [104]. The production of 1-butanol as one of the main volatile organic compounds by Bacillus velezensis strains showing inhibitory activities against Botrytis cinerea, Monilinia spp. and Penicillium spp. has been documented [105]. Mu et al. [106] reported that 1-butanol and acetic acid butyl ester that exhibited strong antifungal activity against B. cinerea were produced by the Bacillus subtilis strain M29 isolated from vermicompost. However, these VOCs had no effect on P. aphanidermatum-induced damping-off in this study. Further, the ability of BCAs in controlling the damping-off of cucumber was drastically reduced upon biofumigation. It has been demonstrated that environmental factors greatly influence the potential of biocontrol agents against phytopathogens. For example, Ng et al. [107] found that Salinispora arenicola M413 produced more Rifamycin antibiotics when grown under low-NaCl (1% NaCl) growth conditions as compared to high-NaCl (3% NaCl) growth conditions. Similarly, the antagonism of Trichoderma harzianum towards Verticillium dahliae decreased with the increase in NaCl concentration in the growth medium [108]. In this study, the ineffectiveness of BCAs, though higher populations were maintained in the soil, to protect cucumber plants from P. aphanidermatum when combined with biofumigation treatment might be due to VOC-induced metabolic alterations in BCAs. Several BCAs including Pseudomonas resinovorans [23], P. aeruginosa [24] and Acinetobacter johnsonii [25] have been reported to control cucumber damping-off with varying levels of effectiveness. This is the first report, according to our knowledge, demonstrating the effectiveness of native antagonistic rhizobacteria from biofumigated soil in controlling Pythium aphanidermatum cucumber damping-off.

5. Conclusions

In this study, soil application of the native antagonist Pseudomonas aeruginosa B1-SQU effectively reduced the incidence of cucumber damping-off under greenhouse conditions. No significant effect of soil biofumigation with cabbage leaf residue on Pythium ahanidermatum damping-off of cucumber was observed. There was no synergism between biofumigation and the use of BCAs on damping-off. Pseudomonas indica B2-SQU showed moderate tolerance to biofumigation treatment and protected cucumber seedlings from P. ahanidermatum infection. This strain was capable of forming more biofilm and showed compatibility with P. aeruginosa B1-SQU. Hence, these two bacterial strains can be used as microbial mixtures for enhanced efficacy in controlling the damping-off of cucumber. Further investigation is needed to elucidate the underlying mechanisms of their antagonistic effects and the effect of GSLs and ITC on metabolic alterations in BCAs, including the production of antibiotics and lytic enzymes. The efficacy of these bacterial strains should be evaluated under field conditions and against other soil-borne diseases of crops in combination with biofumigation in different plant pathosystems.