The infected M. sextelata fruiting bodies were collected from the Morchella cultivation base in Chengxian, Longnan City, Gansu Province. In total, 30 samples were collected, 20 diseased samples, and 10 non-diseased samples, packed in clean self-sealing bags, and stored in a 4°C refrigerator at the laboratory for pathogen isolation. The samples were first rinsed with tap water to remove the soil attached and then rinsed with sterile water. The samples were first immersed in 75% ethanol for approximately 10 s and then in 0.1% mercuric chloride for 10 s, and washed 3 times with sterile water. Then, the materials were cut into 1–2-mm tissue slices with sterile blades and inoculated on Luria–Bertani (LB) solid medium. The LB plates with the slices were incubated in a thermostatic electric incubator at 30°C for 24 h. The bacteria were isolated from the LB agar plates using the scratch method. Single colonies with different morphologies were picked for isolation and purification. A single colony was picked and incubated on the LB plate again by streaking for 24 h until a single colony was obtained. A part of the purified strain was used for subsequent experiments, and the remaining part was stored in 20% glycerol at −80°C for future use.

Pathogenicity tests were conducted according to Koch’s postulates. Morchella sextelata was selected as the main cultivar of Morchella. The pathogenic strains to be tested were inoculated in LB liquid medium for activation and incubated at 30°C with 160 r/min oscillation. Overnight bacterial cultures (1 × 10⁸ CFU/mL) grown in LB medium were injected into the ascocarps of morels by using a 1-mL sterile syringe. The experiment consisted of three treatment groups and three control groups. The first treatment group was only inoculated with M-B bacterial cultures, the second treatment group was only inoculated with M-5 bacterial cultures, the third treatment group was inoculated with both M-B and M-5 bacterial cultures, the fourth group was inoculated with sterilized M-B cultures as a killed bacterial control; the fifth group was inoculated with sterilized M-5 cultures as a killed bacterial control. The LB medium treatment was used as the blank control. Three replicates were established for each treatment group. In order to simulate field cultivation conditions in the laboratory, the inoculated M. sextelata was cultivated in a constant temperature incubation room at 17°C and 85% humidity. The disease incidence rate and symptoms were recorded. Two days after inoculation, some symptoms were observed on the surface. The pathogenic bacteria were reisolated from the infected M. sextelata fruiting bodies on LB plates, and the characteristics of the reisolated bacteria were compared with those of the original culture.

The purified pathogenic bacteria were inoculated on LB plates and incubated at 30°C for 24 h. After observing the growth and morphological characteristics of the colonies, individual colonies were picked for microscopic examination. The isolates were stained using the Gram staining kit. The physiological and biochemical indicators of the isolated pathogenic bacteria isolated were determined by referring to the bacterial identification method presented in the Manual of Identification of Common Bacterial Systems (Dong and Cai, 2001). The physiological indicators included colony morphology, growth temperature, pseudocyanine production, fluorochrome production, utilization of carbon sources (glucose, mannitol, sucrose, trehalose, and fructose), and utilization of nitrogen sources (yeast powder, beef paste, ammonium sulfate, peptone, and sodium nitrate). The biochemical tests conducted included gelatin liquefaction, starch hydrolysis, H₂S production, Voges–Proskauer test, methyl red test, and catalase and urease determination.

All strains isolated were subjected to phylogenetic analysis. DNA was extracted using the bacterial DNA extraction kit. The 16S rRNA gene fragment of the strain was amplified through PCR by using bacterial universal primers 27F (5′-AGAGTTTGATCATGGCTCAG-3′) and 1492R (5′-GGYTACCTTGTTACGACTT-3′). PCR reaction conditions were pre-denaturation at 95°C for 5 min, denaturation at 95°C for 1 min, annealing at 55°C for 30 s, extension at 72°C for 1 min, 34 cycles, extension at 72°C for 5 min amplification (Dou et al., 2018). Then, 1% agarose gel electrophoresis was performed to detect the PCR products. The PCR products were then selected and sent to Jinweizhi Biotechnology Co., Ltd., for sequencing. The sequences were compared with the known sequences in the NCBI database by using BLAST. The phylogenetic tree was constructed using the overlay method (neighbor joining) with MEGA 11.0 software and subjected to the bootstrap test (1,000). The outgroups were Azotobacter chroococcum and Aeribacillus pallidus, respectively.

Bacterial morphology was observed under scanning electron microscopy (SEM). The actively growing bacterial solution was centrifuged at 8,000 rpm for 3–5 min, and the supernatant was discarded and poured into 2.5% glutaraldehyde and fixed overnight. The samples were then washed 3 times with phosphate buffered saline (PBS), fixed in 1% osmium solution for 4–6 h, washed 3 times with phosphate buffered saline, then dehydrated once in 30, 50, 70, 85, and 95% ethanol, and treated twice with 100% ethanol for 15–20 min each. The samples are then subjected to critical-point drying and gold spraying before being ready for SEM (ULTRA Plus-type, Zeiss Company, Germany) (Xie et al., 2005).

The isolated pathogenic bacteria were inoculated on LB plates and incubated at 30°C for 24 h for activation. The laboratory-preserved M. sextelata strain ZW111 was inoculated on potato dextrose agar (PDA) plates and incubated at 20°C for 4 days for activation. The plate standoff method (Wang W. X. et al., 2014) was used to detect the inhibitory effect of the pathogenic bacteria. M. sextelata ZW111’s cake was placed in the center of the PDA plate. The antagonistic bacterial cake (diameter = 8 mm) was placed on three points at equal distance (2.5 cm) from the center of the M. sextelata. Three parallels were run for each treatment. The control group plate only contained M. sextelata ZW111. Two other B. subtilis preserved in this laboratory were selected for testing to preliminarily investigate whether all B. subtilis species inhibited M. sextelata growth or only the strain M-5, isolated in this study, inhibited M. sextelata. P. chlororaphis subsp. aureofaciens exhibits a strong antifungal activity (Raio et al., 2011; Zhang et al., 2021), and so, other strains were not tested. All plates were incubated at 25°C for 4 days in a constant temperature incubator. The pathogenic bacteria in the plates were observed for their inhibitory effect on M. sextelata, and observed morphological changes in the mycelium under SEM. The samples were taken from the mycelium of M. sextelata located in the marginal region of the inhibition zone, and the subsequent fixation, dehydration, drying and spraying gold treatments of the samples were referred to in 2.3.

The occurrence of bacterial diseases of M. sextelata during cultivation is closely related to the influence of temperature, especially high temperature and high humidity, we determined the effect of temperature on the growth of pathogenic bacteria to provide a better theoretical reference for preventing and controlling field diseases of morels. The prepared LB liquid medium was divided into test tubes, with each tube containing 10 mL LB medium. Then, 0.1 mL of bacterial solution (1 × 10⁸ CFU/mL) was added to each of the test tubes. The inoculated tubes were placed on a shaker at 4, 10, 15, 20, 25, 28, 30, 35, 37, 40, and 42°C, 160 r/min and incubated with shaking for 24 h. Three replicates were established set for each group of temperature, and the culture medium without any inoculum was used as the control. After 24 h of incubation, the OD_600nm values of the bacterial broth incubated at different temperatures were measured using a ultraviolet-visible spectrophotometer and recorded (Zhou, 2013).

The isolated pathogenic bacteria were inoculated in carboxymethyl cellulose (CMC) agar medium (0.5% CMC-Na, 0.1% NaNO₃, 0.1% K₂HPO₄, 0.1% NaCl, 0.05% MgSO₄, 0.05% yeast extract, 1.5% agar) and incubated at a constant temperature of 37°C. After incubation, the CMC agar plates were flooded with 0.2% Congo red for 30 min, and destained by washing twice with 1 mol/L NaCl solution for 20 min. Standard cellulase solution (MAKLIN Reagent Co.) was used as a positive control and inactivated enzyme solution as a negative control. The activity zones had observed against a red background after Congo red staining (Zhang et al., 2022).

The filter paper enzyme activity (FPase) of cellulase was measured through the amount of reducing sugars liberated during hydrolysis using filter paper as substrate by the DNS (3,5-dinitrosalicylic acid) method (Ghose, 1987; Xie et al., 2023). The strains to be tested were inoculated into cellulase production broth (1% CMC-Na, 1% peptone, 0.00075% FeSO₄⋅7H₂O, 0.00025% MnSO₄⋅H₂O, 0.0002% ZnSO₄), incubated at 30°C for 3–4 days at 160 r/min, and centrifuged at 6,000 r/min for 10 min, and the supernatant was the crude enzyme solution. Cut the filter paper into 1 cm × 3 cm strips, roll them into small rolls and place them in a 5 mL test tube, add 1.75 mL of acetic acid-sodium acetate buffer (pH = 4.8) and 0.25 mL of enzyme solution, respectively. Place the tube in a water bath at 50°C for 1 h, then the reaction was terminated by 1 mL DNS. The color of the mixture was developed by incubating it for 10 min at 100°C. The inactivated crude enzyme solution was treated as above and used as a control. The absorbance was measured at 540 nm by using an ultraviolet-visible spectrophotometer. A standard curve was plotted using glucose. The equation of the standard curve is y = 0.5515 × +0.0206 and the correlation coefficient is R² = 0.9996 (Supplementary Table 3). One unit (U) is defined as the amount of enzyme required to produce 1 μmol of glucose equivalent released per minute under the conditions described above (Li et al., 2020; Sarangthem et al., 2023).

The isolated pathogenic bacteria were inoculated on colloidal chitin agar medium [1% colloidal chitin, 0.5% NaCl, 0.05% MgSO₄⋅7H₂O, 0.07% K₂HPO₄, 0.03% KH₂PO₄, 1% (NH₄)₂SO₄, 0.3% yeast extract powder, and 1.5% agar] and incubated at a constant temperature of 37°C. Colloidal chitin was prepared according to the method by Sandhya et al. (2004). After incubation, the plates were stained with 0.2% Congo red using the same method as in 2.6. Standard chitinase solution (MAKLIN Reagent Co.) was used as a positive control and inactivated enzyme solution as a negative control (Linda et al., 2018).

The chitinase activity of the strains was also determined by the DNS (3,5-dinitrosalicylic acid) method as in 2.6. The strains were inoculated into colloidal chitin broth (1% colloidal chitin, 0.05% MgSO₄⋅7H₂O, 1.2% urea, 0.03% KH₂PO₄, 0.5% NaCl, 0.07% K₂HPO₄, 0.3% yeast extract powder), the supernatant was the crude enzyme solution. Mix 0.5 ml crude enzyme solution with 0.5 ml colloid chitin and 0.5 ml PBS by volume. The reaction was carried out in a water bath at 37°C for 30 min, then the reaction was terminated by 1 mL DNS. The absorbance was measured at 540 nm. The inactivated crude enzyme solution was treated as above and used as a control. A standard curve was plotted using N-acetyl-D-glucosamine. The equation of the standard curve is y = 0.4063 × −0.0384 and the correlation coefficient is R² = 0.9994 (Supplementary Table 4). One unit (U) is defined as the amount of enzyme required to catalyze the production of 1 μmol of N-acetyl-D-glucosamine from the substrate in 1 min under suitable conditions as one enzyme activity unit (U) (Du et al., 2021).

All experiments were repeated in triplicate. The data were processed using the SPASS statistical software to perform an ANOVA [least-significant difference (LSD) test and Duncan’s multiple range test, p < 0.05]. Origin software was used for statistical analysis and graph drawing.

The collected infected M. sextelata samples were reddish in color and had a foul odor (Figure 1A). We isolated two types of bacteria, named M-B and M-5, from the diseased M. sextelata, and we did not isolate these bacteria from the undiseased M. sextelata. Strain M-B was inoculated on LB solid medium and incubated at 30°C for 24 h. The colonies appeared orange, smooth, and opaque with a round, slightly elevated surface (Figure 1B). Strain M-5 was inoculated on LB solid medium and incubated at 30°C for 24 h. After incubation, the colonies appeared creamy white, subcircular, with irregular edges, rough and opaque, and a dry, wrinkled surface (Figure 1C).

The treated M. sextelata were placed in the incubation room at 17°C and 85% humidity. At 36 h after inoculation, reddish spots started to appear from the base of the stipe of M. sextelata in all bacterial solution treatment groups (Figure 2). After 60 h, the whole stipe began to turn red, and the red color of the stipe was the deepest in the mixed treatment group, and the M-5-treated M. sextelata started to produce a foul odor (Figure 2). At 84 h, the whole mushroom body in all bacterial solution treatment groups began to turn red and the red color continued to deepen (Figure 2). A total of 108 h later, the mushroom body began to turn reddish brown and soft rot (Figure 2). At 132 h, the whole body was dark red, wilted, and crumpled, accompanied by a large amount of unpleasant odor (Figure 2). Similar to field symptoms. The three control groups inoculated with sterilized M-B bacterial solution, sterilized M-5 bacterial solution, and LB blank medium showed slight browning over time, with no softening of the fruiting body and without symptoms of disease. Pathogenic bacteria were reisolated from these infected M. sextelata. These isolates exhibited similar morphologies on LB solid plates, satisfying Koch’s postulates.

According to the results of Gram staining, strain M-5 (Supplementary Figure 1A) was stained purple and exhibited a rod-shaped body, with bluntly rounded ends and budding spores. It was tentatively determined to be Gram-positive. Morphological analysis through SEM (Supplementary Figure 1B) revealed that the bacterium was rod-shaped with no pods and a size of (0.6–0.7) μm × (2.3–1.4) μm. Its endospore was located in the center of the body or slightly deviated and did not expand after the bacterial cell became oval. Strain M-B (Supplementary Figure 1C) was stained red and exhibited a rod-shaped body. It was tentatively determined to be Gram-negative. SEM (Supplementary Figure 1D) revealed that the bacterium was rod-shaped, with a size of (0.4–0.5) μm × (1–1.3) μm.

Physiological and biochemical characterization of strains M-B and M-5 showed (Table 1) that both strains M-B and M-5 were able to utilize glucose, mannitol, sucrose, trehalose, fructose, yeast, beef extract, ammonium sulfate, peptone, sodium nitrate. Strain M-B was negative for starch hydrolysis, H₂S production, methyl red test, positive for gelatin liquefaction, Voges–Proskauer test, catalase and urease reaction. Strain M-5 was negative for H₂S production, methyl red test and urease reaction, and positive for gelatin liquefaction, catalase, Voges–Proskauer test and urease reaction. According to the physiological and biochemical tests, and combined with morphological characteristics, according to the Manual of Identification of Common Bacterial Systems, the strain M-B was preliminarily identified as Pseudomonas chlororaphis subsp. aureofaciens and strain M-5 was preliminarily identified as Bacillus subtilis.

After sequencing was completed, the sequences were compared with the known sequences in the NCBI database by using BLAST. The sequences with a high homology were downloaded and compared using MEGA 11.0 software to construct a phylogenetic tree by using the neighbor-joining method. The results revealed (Figure 3) that the isolated strains belonged to two main groups. M-5 and B. subtilis converged in the same species, and so, the bacterium was tentatively identified as B. subtilis. M-B and P. chlororaphis subsp. aureofaciens converged in the same species, and therefore, the bacterium was tentatively identified as P. chlororaphis subsp. aureofaciens.

The plate standoff results (Figure 4) revealed a clear inhibition circle between the pathogenic bacteria and M. sextelata. Strains M-B and M-5 severely inhibited mycelial growth in M. sextelata (Figures 4A, C). SEM showed that the mycelium of M. sextelata in the control group was healthy and full (Figure 4F), whereas that at the edge of the colonies of M. sextelata treated with the pathogenic strains exhibited an obvious distortion, surface depression, folding, drying, and rupture (Figures 4B, D). The results thus indicated that the pathogenic bacteria changed the morphology of the M. sextelata mycelium. The laboratory-preserved other strains of B. subtilis did not produce an inhibition circle around M. sextelata (Figures 4G, I), the M. sextelata mycelium covered the growth of the B. subtilis cake and the mycelium was full and strong. This indicated that the other B. subtilis strains had no inhibitory effect on the mycelial growth of M. sextelata on the plate. These results preliminarily indicate that perhaps not all B. subtilis strains inhibit M. sextelata growth. Only the B. subtilis strain M-5 isolated from the infected M. sextelata exhibited an inhibitory effect on M. sextelata. In the following research, we will continue our in-depth investigation on whether all B. subtilis will inhibit the growth of M. sextelata. The difference between different pathogenic and non-pathogenic strains among the same species also serves as a direction for subsequent research.

The temperature effect results showed (Figure 5 and Supplementary Tables 1, 2) that the growth of strain M-B was inhibited at 4°C. When the temperature was increased, the OD_600nm values were the maximum at 35°C, and the growth was inhibited when the temperature was higher than 40°C. Therefore, strain M-B did not grow at temperatures below 4°C or above 40°C. When the strain was incubated for 10 min in a water bath at 55°C and coated on the LB plate, no colonies were observed for 48 h, indicating that the lethal temperature for strain M-B was 55°C. The growth of strain M-5 was inhibited at 4°C, and the OD_600nm values were the maximum at 25°C with an increase in temperature. After strain M-5 was incubated on LB plates at 110°C for 10 min, no colonies were observed for 48 h, indicating that the lethal temperature for strain M-5 was 110°C. Strain M-B started to multiply at 10–15°C, and strain M-5 started at 15–20°C. Therefore, the temperature can be controlled below 15°C to as the lower temperature could inhibit the growth of pathogenic bacteria during M. sextelata cultivation.

The results of cellulase secretion by two bacterial pathogens inoculated into cellulose medium by spot inoculation are shown in Figure 6. Congo red binds strongly to the large polysaccharide cellulose, which is hydrolyzed by the action of cellulase into smaller sugars that Congo red cannot bind, resulting in a hyaline ring around the colony. A clear circle can be seen around the edge of the colony of strain M-5, while no clear circle is evident around strain M-B, indicating that strain M-5 can secrete cellulase. Based on the enzyme activity data (Table 2 and Supplementary Table 5), the enzyme activity value of strain M-5 is much higher than that of strain M-B, indicating strong cellulase production ability. Strain M-5 has lower enzyme activity and weaker cellulase production ability.

The two bacterial pathogens were inoculated separately into the chitinase medium by spot inoculation and the results of chitinase secretion by both pathogens are shown in Figure 7, where clear circles can be seen at the edge of the colonies of both strains M-5 and M-B. This indicates that both strains can secrete chitinase. Under the action of chitinase, chitin can be hydrolyzed from white insoluble particles to soluble small molecules, resulting in a transparent ring around the colony of the bacterium to be tested. Combined with the enzyme activity data (Table 2 and Supplementary Table 6), both strain M-5 and strain M-B had strong chitinase production ability.