Samples were collected from the Lienhuachi Experimental Forest (Permission no.: 1082272516), an evergreen broadleaf forest in central Taiwan (23°55'7''N 120°52'58''E) in 2020. Ant cadavers infected by O. pseudolloydii were carefully removed by cutting the leaf and placing it into a 50-mL conical centrifuge tube, which was then transported to the laboratory. Only cadavers in which the fungal growth stage preceded the development of pale yellow perithecia, which theoretically has the highest biological activity, were collected (Fig. 1). Altogether, 24 infected D. thoracicus samples were collected and examined in this study.
Bacterial isolation and cultivation
The protocols used for isolating bacteria were described in our previous work (Tu et al. 2021). In brief, sample surfaces were cleaned by agitation in 600 μL of sterilized water using a vortex mixer (AL-VTX3000L, 114 CAE Technology Co., Ltd., Québec, Canada), soaked with 600 μL of 70% ethanol, washed twice with 600 μL of sterilized water, then vortexed in 400 μL of sterilized water. Two hundred microliters of the supernatant was spread homogeneously onto a Luria-Bertani (LB) agar plate (25 g LB broth and 15 g agar per liter) to confirm the absence of live bacteria. The cleaned samples were homogenized in 200 μL of water to release the internal bacteria and cultured on LB agar plates at 28°C for 2 days. Approximately equal numbers of the bacterial isolates from each of the ant individuals were picked randomly with sterile toothpicks and suspended individually in the LB medium supplemented with 15% (v/v) glycerol and maintained at -80°C until examination. In total, 137 bacterial isolates were collected. In addition to the bacterial isolates from the ant bodies, 60 bacterial isolates from soil, leaves, and air in the same locality were sampled to examine the tolerance to naphthoquinones (see below) using the procedure mentioned above but without initial cleaning and sterilizing of the sample surface.
Genomic DNA was extracted from the bacterial isolates cultured in LB medium at 28°C overnight. Randomly amplified polymorphic DNA (RAPD) analysis with the primer 5′-GAGGGTGGCGGTTCT-3′ (Huey and Hall 1989) was used to determine the bacterial strain. The PCR condition followed our previous work: initial denaturation at 95°C for 5 min, 40 cycles of amplification, including denaturation at 95°C for 1 min, annealing at 42°C for 30 s, and extension at 72°C for 1 min, followed by a final extension at 72°C for 10 min. The RAPD pattern was checked by gel electrophoresis of the PCR products on a 2% agarose gel. Bacterial isolates with the same RAPD pattern were determined as the same strain and coded with "JYCB," followed by a series of numbers (e.g., JYCB1491).
The taxonomic status of each strain was determined by sequencing the V3/V4 hypervariable regions of the 16S rDNA gene. The sequence was first amplified by PCR with the primer set (8F: 5′-AGAGTTTGATCCTGGCTCAG-3′ and 1541R: 5′-AAGGAGGTGATCCAGCCGCA-3′) under the following condition: initial denaturation at 95°C for 5 min, 40 cycles of amplification, including denaturation at 95°C for 1 min, annealing at 55°C for 30 s, and extension at 72°C for 1 min 45 s, followed by a final extension at 72°C for 10 min (Edwards et al. 1989; Turner et al. 1999), and sequenced at Genomics, Inc. (New Taipei City, Taiwan).
To identify the taxon, the bacterial strains were first clustered into the same clades according to the sequences dissimilarity (<0.01) calculated by the unweighted pair group method with arithmetic mean (UPGMA) using MEGA X (Kumar et al. 2018). Each clade was considered as an operational taxonomic unit in this study. The species was judged by the basic local alignment search tool (BLAST) method against nucleotide sequences in the National Center for Biotechnology Information (NCBI) nucleotide database (https://ftp.ncbi.nlm.nih.gov/blast/db/), updated May 17, 2021.
Each strain was first labeled with the species of the sequence with the highest BLAST identity, which was ranked by expect value (E-value), bit score, percentage of identical matches, and alignment length (https://www.ncbi.nlm.nih.gov/BLAST/tutorial/Altschul-1.html). If multiple sequences from the database were found to be the same in the identity indexes, the bacterial strain was labeled by the species that appeared most frequently. In the cases of the clades containing several bacterial strains, the species were judged by the bacterial species found most frequently in the strains belonging to the clade.
The 60 bacterial isolates collected from the environment were examined using the RAPD method and a Bacillus-specific primer set (5′-CTTGCTCCTCTGAAGTTAGCGGCG-3′ and 5′-TGTTCTTCCCTAATAACAGAGTTTTACGACCCG-3′), with the PCR conditions suggested by Nakano et al. (Nakano et al. 2004). Twenty of the bacterial isolates (10 Bacillus and 10 non-Bacillus) with different RAPD patterns were collected for their resistance to naphthoquinones (see below).
Biological properties of bacterial isolates from infected ants
All the bacterial strains were selected for examining the hemolysis reaction. Among multiple isolates belonging to a strain, one isolate was selected randomly for the examination. To characterize the biological properties, including hydrolytic activity, repellence against entomopathogenic fungi, and resistance to naphthoquinone derivatives, 20 bacterial strains (Fig. S1) were selected according to the UPGMA analysis of the sequences, which maximally cover the diverging clades. The 20 bacterial strains included 13 strains of the predominant species (B. cereus/thuringiensis, clade 7), and one strain from each of the clades 1, 6, 8, 9, 12, 17, and 18 with relatively lower abundance (Table 1, Fig. S1). We selected one hemolytic strain (JYCB1618) and one non-hemolytic strain (JYCB1543) to preliminary examine the effect of co-cultured nematodes because the antagonistic effect might have the potential to be a repellent against the soil pests.
For hemolysis reaction tests, one 3-µL drop of the log-phase bacterial suspension was placed onto tryptic soy agar (TSA) plates (15 g pancreatic digest of casein, 5 g soybean meal, 5 g NaCl, and 15 g agar, with final pH of 7.3), which was mixed with 5% defibrinated sheep blood after it had cooled down to approximately 50°C. The hemolysis reaction was determined by the formation of clean (β-hemolysis) or greenish (α-hemolysis) hemolytic zones, or no such zone (γ-hemolysis, non-hemolytic) around the bacterial colonies after incubation at 28°C for 1–2 days (Baumgartner et al. 1998).
Resistance to naphthoquinones
To examine the resistance of bacterial isolates to naphthoquinones, the growth of 13 bacterial strains from the predominant clade (clade 7) and seven low-abundant clades (clade 1, 6, 8, 9, 12, 17, 18) isolated from the ant host, and the 20 environmental bacterial isolates (10 Bacillus and 10 non-Bacillus) was compared using two naphthoquinones, respectively. As the fungal naphthoquinones are currently not purified and commercialized, the two naphthoquinones prepared for the experiment, plumbagin (de Paiva et al. 2003) and lapachol (Eyong et al. 2006), were those found in plants. The two naphthoquinones prepared for the experiment were dissolved in 30% dimethyl sulfoxide (DMSO)-70% water solution. Naphthoquinone concentrations were determined from the serial dilutions in which three randomly selected bacteria from the ant host and three from the environment had the most distinctive growth rate. Based on these results, concentrations of 45 µg/mL (plumbagin) and 64.5 µg/mL (lapachol) were used.
In this experiment, the bacterial isolates were first inoculated in LB medium at 20°C (the mean annual temperature in Lianhuachi Research Center, where the infected ants were collected) overnight, then refreshed to the exponential phase with LB medium for 3 h. The bacterial concentration was adjusted to ~1.5×108 cells/mL. Next, 10 μL of the bacterial suspension and 180 μL of Mueller-Hinton broth medium (Sigma-Aldrich) were added to either 10 μL of the naphthoquinone solution or 10 μL of the 30% DMSO-70% water solution for the control. The growth of bacterial isolates at 20°C was monitored by measuring the optical density at 600 nm (OD600) with a Multiskan GO microplate spectrophotometer (Thermo Scientific) every hour for 12 h. Each combination of bacterial isolate and naphthoquinone or control was replicated twice.
The resistance index of each bacterial isolate was calculated by the following formula: [naphthoquinone – DMSO] / [naphthoquinone + DMSO]) (Tu et al. 2021). The resistance index was analyzed using a linear mixed model with the resource (Bacillus from the ant host, Bacillus and non-Bacillus from the environment) as the fixed effect, the bacterial isolate as a random effect, and growth time (5-12 h) as a nest effect. The significance of the fixed effect was examined by the likelihood ratio test with the model removing the fixed-effect term. Post hoc tests were undertaken using Tukey's all-pair comparisons. The model building and hypothesis tests were conducted using the "lme4" and "multcomp" packages in R.
Production of hydrolytic enzymes
The productions of chitinase, proteinase, lipase, and esterase were examined with the four different types of plated media, including chitinase detection medium (solid medium with 0.3 g MgSO4.7H2O, 3 g (NH4)2SO4, 2 g KH2PO4, 1 g citric acid monohydrate, 0.15 g bromocresol purple, 200 μL Tween 80, 4.5 g colloidal chitin, and 1 L deionized water with 1.5% [w/v] agar, with final pH of 4.7), skim milk agar (solid medium with 2% [w/v] agar, 28 g skim milk powder, 5 g casein enzymic hydrolysate (Tryptone), 2.5 g yeast extract, 1 g dextrose, and 1 L deionized water), lipase agar (solid medium with 2% [w/v] agar, 0.1 g phenol red, 1 g CaCl2, 10 mL olive oil, and 1 L deionized water, with final pH of 7.4), and esterase agar (solid medium with 2% [w/v] agar, 0.1 g phenol red, 1 g CaCl2, 10 mL tributyrin, and 1 L deionized water, with final pH of 7.4). A 3-µL drop of the exponential-phase bacterial suspension was cultured on each of the media, and the production of the hydrolytic enzymes was determined by purple zones for chitinase activity (Chi et al. 2009; Agrawal and Kotasthane 2012), clearance zones for proteinase activity (Cattelan et al. 1999), and yellow zones for lipase and esterase activity (Ramnath et al. 2017). All the experiments were carried out in the dark at 20°C with three replicates.
Antagonism against entomopathogenic fungi
Three entomopathogenic fungi, including Aspergillus nomius (isolated from D. thoracicus), Trichoderma asperellum (isolated from the litchi stink bug, Tessaratoma papillosa), and Purpureocillium lilacinum (isolated from T. papillosa), were cultured on potato dextrose agar (PDA) plates for 4 (A. nomius, T. asperellum) or 10 (P. lilacinum) days at 28°C, until the mycelia covered approximately 80% of the plate. A piece of mycelium (approximately 5 × 5 mm2) was seeded in the center of a TSA plate and surrounded by three equidistant 3-μL drops of the exponential-phase bacterial suspension. After incubation at 20°C for 7-10 days, photographs were taken. The growth of the entomopathogenic fungi was assessed by calculating the areas of the mycelium on the digital images using ImageJ. The control (a piece of mycelium with blank LB suspension) and each pair of bacteria and entomopathogenic fungi were replicated 3–4 times.
Antagonism was expressed as the percentage of mycelial growth inhibition calculated by the formula ([Rmc – Rexp] / Rmc) × 100%, where Rmc represents the mean mycelial area of the control fungus, and Rexp is the area of the examined entomopathogenic fungus co-cultured with the examined Bacillus (Michereff et al. 1994). For each of the three entomopathogenic fungi, the significance of antagonism of each of the bacteria was examined by comparing the mycelial area with the respective control using Student's t-test, and P-values were adjusted by the Holm–Bonferroni method.
Adverse effects of bacterial isolates on nematode
To simulate the effect of bacteria on the free-living stage of the plant-pathogenic nematodes, the model nematode, Caenorhabditis elegans strain N2, was used for the examination. Daily mortality of the nematode was examined in response to hemolytic and non-hemolytic bacterial strains. Synchronized L4 nematodes were cultured on nematode growth medium (NGM; 3 g NaCl, 2.5 g peptone, 17 g agar, 5 mg cholesterol, 1 mL of 1 M CaCl2, 1 mL of 1 M MgSO4, 25 mL of 1 M KH2PO4, and H2O to 1 L) agar plates seeded with the regular food, Escherichia coli OP50. The examined bacterial isolates were prepared by inoculating in 3 mL of LB liquid broth at 20°C overnight and then adjusting the OD600 value to 0.2. Five treatments were used to examine the survival rate by co-culturing the nematodes with a 20 μL bacterial suspension of 1) the hemolytic bacterial strain, 2) the non-hemolytic bacterial strain, 3) mix of the hemolytic strain + E. coli OP50, 4) mix of the non-hemolytic strain + E. coli OP50, and 5) E. coli OP50 only (control). For each treatment, 30 L4 larvae were cultured on the NGM agar plate, and the daily survival rate was monitored for 5 days. Each treatment was replicated three times. Survival curves were compared by survival analysis with treatment as the fixed effect and replication as the block. The significance of the fixed effect was assessed by model reduction and the likelihood ratio test. Post hoc multiple comparisons were conducted with Tukey's all-pair comparisons. The model building and hypothesis tests were conducted using the "survival" and "multcomp" packages in R.