Isolation of Bacillus spp. from soil
Six soil samples were collected from different localities in Sharkia province, Egypt according to the methods described by Johnson et al. (1959). The protocol for isolation of Bacillus spp. from soil, described by Wollum (1982), was used with some modifications. One gram of each soil sample was mixed with 10 ml sterile distilled water. Thereafter, supernatants were diluted by 10-fold serial dilution as 100 μl of the 10-2 up to 10-7 dilutions in saline solution. One milliliter of soil dilution was spread on a nutrient agar plate (4 plates for each dilution) and incubated at 35 °C for 24 h. The different-looking bacterial colonies developed on each plate were selected to sub-culturing and purifying, using nutrient agar medium. These colonies were characterized according to the key of Bergey’s Manual of Systematic Bacteriology (Holt et al., 1994) and depending on Gram’s stain reaction, morphological and biochemical tests. The confirmed Bacillus isolates were tested for their ability to produce biofilm.
Detection of biofilm-forming isolates
Congo red agar (CRA) method described by Bose et al. (2009) was used to detect biofilm production in CRA medium, which was prepared with brain heart infusion: sucrose: agar and Congo red stain (37:50:10 and 8 g/l, respectively). Congo red stain was separately prepared as concentrated aqueous solution and autoclaved at 121 °C for 15 min, then added to the autoclaved brain heart infusion agar with sucrose at 55 °C, CRA plates were inoculated with test organisms and incubated aerobically at 37 °C for 24 h. Black colonies with a dry crystalline consistency indicated biofilm production. The experiment was repeated 3 times.
Quantity estimation of biofilm formation
Thirty Bacillus cultures were isolated and screened for their biofilm formations. The overnight cultures were grown in 96-well micro titer plate, filled with brain heart infusion broth medium (BHI), then the plates were incubated for 48 h at 30 °C.
The biofilm formation by each Bacillus culture was measured and quantified, using Crystal Violet (CV) assay as described by Castelijn et al. (2012). After incubation, the wells were gently washed 3 times with phosphate-buffered saline (PBS) pH 7.1 (1 mM KH2PO4, 10 mM Na2HPO4, 3 mM KCl, 140 mM NaCl). The attached bacteria were fixed with 0.1% (W/V) crystal violet for 30 min. The wells were washed 3 times with PBS. Thereafter, 0.2 ml of ethanol (70% (V/V)) were added to each well and incubated for 30 min to dissolve bounded biofilm. The optical density (OD) of solubilized bounded biofilm in each well was measured at 595 nm, using an ELISA Microplate Reader. The negative control was determined in uninoculated BHI broth under the same conditions.
Biofilm production and ECM extraction
Single colony of tested isolates were inoculated into 3 ml of BHI medium and incubated overnight at 37 °C with shaking. The overnight cultures were 1000-fold diluted in 10 ml of the Brain heart infusion medium supplemented with 1% glucose and incubated at 37 °C for 24 h under static conditions. After the incubation, the cultures were centrifuged at 6000 rpm for 10 min at 25 °C, and the pellets were collected for extraction of biofilms. Pellets were weighted and suspended with 1.5 M NaCl solution at various concentrations 5, 10, and 20% (w/v). The suspensions were centrifuged at 1000 rpm for 10 min at 25 °C, and the supernatants were transferred to a new test tube as ECM which would be subjected to bioassay (Chiba et al., 2014).
Insect culture and experimental conditions
Insect cultural rearing and experiments were carried out under controlled environmental conditions in the Department of Pest Physiology, Plant Protection Research Institute, Sharkia branch, Egypt. C. maculatus adults, collected from naturally infested cowpea seeds, were reared on infested cowpea seeds. The cultures were maintained in a controlled chamber under a 12/12 h light: dark photoperiod at 28 ± 2 °C and 70 ± 5% relative humidity (RH). Newly emerged adult weevils were used for bioassay.
Bioassays of biofilm against cowpea weevil, C. maculatus
Totally. 90 seeds of cowpea were taken in a conical flask and separately mixed with each concentration of ECM previously prepared, and seeds treated with NaCl 1.5 M solution alone was used as a control. The treated seeds were air-dried and they were separated into 3 plots, each having 30 seeds, each placed in a separate plastic container; 5 pairs of newly emerged adults were introduced into the containers. All were maintained for 15 days under experimental conditions. Adult mortality, number of eggs laid, eggs hatching, larval development, and adult emergence were recorded in both treated and control seeds.
Molecular identification of the potent bacterial strains
Genomic DNA from the most potent bacterial strain was extracted as reported by Hyronimus et al. (1998). Bacterial DNA was subjected to the polymerase chain reaction (PCR), using universal primers; F (5-AGAGTTTGATCCTGGCTCAG-3′) and R (5-GGTTACCTTGTTACGACTT-3′) and recommended thermal cycling conditions (activation 95 °C for 10 min and 35 cycles of 95 °C for 30 s, 65 °C for 1 min, 72 °C for 1 min and 30 s, extension 72 °C for 10 min) to amplifying 16S rDNA gene fragments.
The PCR products were cleaned with the QIAquick gel extraction kit (Qiagen Inc., Valencia, CA) then, purified PCR product was sequenced by Applied Biosystems 3130 automated DNA Sequencer (ABI, 3130, USA). Using a ready-reaction BigDye Terminator v3.1 Cycle Sequencing Kit (Perkin-Elmer/Applied Biosystems, Foster City, CA), with Cat. No. 4336817. A BLAST (Basic Local Alignment Search Tool) analysis was initially performed to establish sequence identity to GenBank accessions (Altschul et al., 1990).
Phylogenetic analysis
A comparative analysis of sequences was performed, using the Clustal W multiple sequence alignment program, version 1.83 of MegAlign module of DNASTAR Lasergene software Pairwise, which was designed by Thompson et al. (1994), and phylogenetic analyses were done, using maximum likelihood, neighbor joining and maximum parsimony in MEGA6 (Tamura et al., 2013).
