The bacterial strains and plasmids used in this study are listed in Table 1. B. glumae BGR1 was used as the wild-type strain. The ntrC deletion mutant (ΔntrC) and complemented strain (CntrC) were generated in this study. Escherichia coli DH5α λpir and E. coli S17-1 λpir were used for plasmid construction and mobilization, respectively.

All B. glumae and E. coli strains were routinely cultured in Luria-Bertani (LB) broth or on LB agar at 37°C. When required, antibiotics were added at the following concentrations: rifampicin, 100 μg mL⁻¹; kanamycin, 30 or 50 μg mL⁻¹; and benomyl, 50 μg mL⁻¹. Antibiotics were used individually or in combination for plasmid selection or strain maintenance. For long-term preservation, bacterial cultures were stored at −80°C in LB broth containing 20% (v/v) glycerol.

Genomic DNA was extracted from B. glumae BGR1 using the Wizard^® Genomic DNA Purification Kit (Promega, Madison, WI, USA) according to the manufacturer’s instructions. Plasmid DNA was isolated with the HiGene™ Plasmid Mini Prep Kit (BIOFACT, Daejeon, Korea). DNA concentration and purity were measured using a NanoDrop ND-2000 spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). Polymerase chain reaction (PCR) amplifications were performed using Pfu-X DNA Polymerase (Solgent, Daejeon, Korea) in an Eppendorf Pro S vapo.protect thermocycler. Primer sequences and their corresponding target regions are listed in Table 2.

The ΔntrC mutant and its complemented strain were generated following standard molecular cloning techniques (Green and Sambrook, 2012). For gene deletion, the upstream (L) and downstream (R) flanking regions of ntrC were amplified from B. glumae BGR1 genomic DNA using primers carrying appropriate restriction sites (Table 2). The two fragments were fused by overlap-extension PCR to obtain a continuous LR fragment, which was cloned into the suicide vector pK18mobsacB (Schäfer et al., 1994). The recombinant plasmid was first propagated in E. coli DH5α λpir and screened on kanamycin-containing LB agar using blue-white selection. Verified constructs were transferred to E. coli S17-1 λpir, which served as the donor strain for biparental conjugation with B. glumae BGR1 (Simon et al., 1983). Transconjugants were selected on LB agar containing rifampicin (for B. glumae) and kanamycin (for the plasmid).

Single-crossover recombinants were obtained via homologous recombination at the ntrC locus. Double-crossover events leading to an unmarked deletion were selected by repeated subculturing on LB agar supplemented with 20% sucrose for sacB-based counterselection. Sucrose-resistant, kanamycin-sensitive colonies were screened by PCR to confirm the loss of ntrC.

For genetic complementation, the full open reading frame of ntrC was amplified and cloned into the broad-host-range vector pBBR1MCS-2 (Kovach et al., 1995). The recombinant plasmid was introduced into the ΔntrC mutant by electroporation. Transformants were selected on LB agar containing rifampicin and kanamycin, and successful complementation was verified by PCR amplification and Sanger sequencing (Macrogen, Seoul, Korea).

To evaluate the role of NtrC in growth regulation under different nutrient conditions, B. glumae BGR1 (wild-type), ΔntrC, and CntrC strains were cultured in nutrient-rich and nitrogen-limited media. For nutrient-rich conditions, bacterial growth was monitored in LB broth at 37°C with shaking at 200 rpm.

For nitrogen-limited conditions simulating the rice apoplastic environment, a minimal medium (MM) was used as the basal formulation containing 30 g L⁻¹ sucrose, 1 g L⁻¹ KH₂PO₄, 0.5 g L⁻¹ MgSO₄·7H₂O, 0.5 g L⁻¹ KCl, and 0.2 mL L⁻¹ of a trace element solution (Table 3). Nitrogen limitation was imposed by supplementing MM with a single nitrogen source (3 mM), either ammonium sulfate (MM-AS) or glutamine (MM-Gln).

Overnight bacterial cultures were washed twice with sterile 1 mM MgSO₄ and adjusted to an initial optical density of 0.05 at 600 nm (OD₆₀₀). Cultures were incubated at 37°C with shaking, and optical density was recorded at designated time points over a 48 h period using a spectrophotometer. Each experiment was performed in triplicate with three independent biological replicates.

The virulence of B. glumae BGR1 wild-type, ΔntrC mutant, and complemented (CntrC) strains was evaluated on rice (Oryza sativa L. cv. Dongjin) at the flowering stage. Bacterial suspensions were prepared in 1 mM MgSO₄ and adjusted to an optical density (OD₆₀₀) of 0.8. Rice panicles were immersed in 35 mL of the bacterial suspension for 1 min, while 1 mM MgSO₄ served as the negative control. Plants were maintained under greenhouse conditions at 37°C and nearly 100% relative humidity at the Department of Southern Area Crop Science, National Institute of Crop Science (Miryang, Korea).

Disease symptoms were assessed 14 days post-inoculation (dpi). Disease severity was determined using the panicle blight index (Kim et al., 2023), rated on a 0-5 scale: 0 = no symptoms, 5 = severe panicle blight (80-100% of the panicle affected). Thirty panicles were used per treatment, and disease severity was calculated as:

To assess virulence during early infection, O. sativa L. cv. Dongjin seeds (Korea Seed & Variety Service, Gimcheon, Korea) were used in a seedling blight assay. Bacterial suspensions (OD₆₀₀ = 0.8 in 1 mM MgSO₄) were prepared, and 30 seeds were soaked in 15 mL of suspension for 24 h at 37°C with shaking (200 rpm). Seeds were then transferred to sterile containers and germinated at 30°C for 3 days, followed by growth for 7 days under a 12 h light/12 h dark cycle (25°C day/22°C night). Control seedlings were treated with 1 mM MgSO₄ only. Disease severity was assessed by measuring shoot and root lengths relative to controls. Each treatment included 30 seedlings.

The rice seedling infection assay was repeated under controlled growth conditions, and disease severity was recorded at 10 dpi. Shoot and root lengths were measured to quantify growth inhibition relative to the control. To examine whether the virulence attenuation of the ΔntrC mutant was associated with reduced colonization, bacterial population densities were quantified in infected seedlings at 0 and 10 dpi. Ten seedlings per treatment were homogenized in 20 mL sterile 1 mM MgSO₄ using a pre-chilled mortar and pestle. Homogenates (100 μL) were serially diluted, and 3 μL from each dilution was spotted on LB agar plates containing rifampicin (100 μg mL⁻¹), kanamycin (50 μg mL⁻¹), and benomyl (50 μg mL⁻¹) to selectively recover B. glumae BGR1. Plates were incubated at 37°C for 24 h, and colony-forming units (CFU) were counted to calculate bacterial populations (CFU mL⁻¹). Each assay was performed in three independent experiments with three biological replicates.

Swimming motility was assessed on MM supplemented with 3 mM glutamine (MM-Gln) and solidified with 0.3% (w/v) agar. Overnight cultures were grown in LB broth at 37°C with shaking (200 rpm) and adjusted to OD₆₀₀ = 0.5. Ten milliliters of each culture were centrifuged at 7,000 × g for 3 min, washed twice with sterile 1 mM MgSO₄, and resuspended in 100 μL of the same solution. A 3 μL aliquot was inoculated onto the center of each MM-Gln motility plate (Kim et al., 2023; Nickzad et al., 2015). Plates were incubated at 37°C for 48 h, and motility zone diameters were measured in millimeters. Each experiment was performed in triplicate.

Biofilm formation was quantified using a modified crystal violet assay (Ball et al., 2022; Vestby et al., 2020). Overnight cultures were centrifuged at 7,000 ×g for 3 min, washed twice with 1 mM MgSO₄, and resuspended in MM-Gln medium to OD₆₀₀ = 0.05. A 250 μL aliquot of each suspension was added to sterile 96-well polystyrene plates and incubated at 37°C for 2 days without shaking. After incubation, planktonic cells were removed, and wells were gently washed twice with sterile distilled water. Plates were air-dried at 50°C for 30 min and stained with 0.1% (w/v) crystal violet for 15 min. Excess stain was removed by four washes with distilled water, followed by air drying for 20 min. Bound crystal violet was solubilized in 30% (v/v) acetic acid for 15 min, and absorbance was recorded at 570 nm (Kamimura et al., 2022; O’Toole, 2011). Biofilm biomass was expressed as the mean absorbance from three independent experiments, each with three technical replicates. To verify that differences in biofilm formation were not due to reduced growth under nitrogen-limited conditions, an additional assay was performed using cultures adjusted to a high initial density. Overnight LB cultures were washed twice with sterile MgSO₄ and normalized to OD₆₀₀ = 1.0 before inoculation into MM-Gln medium. Biofilm formation was assessed as described above.

Proteolytic activity was measured on MM + 3 mM Gln supplemented with 1% (w/v) skim milk (BD Difco, Franklin Lakes, NJ, USA). Overnight cultures were adjusted to OD₆₀₀ = 1.0, and 10 μL aliquots were inoculated into 4 mm wells made in the agar using a sterile cork borer. Plates were incubated at 37°C for 3 days, and protease activity was quantified by measuring the diameter of the clear zones around inoculation sites (Jung et al., 2025).

Toxoflavin production was quantified using a modified thin-layer chromatography (TLC) method (Kim et al., 2004; Kim et al., 2023; Lee et al., 2016). Strains were cultured in 4 mL MM-Gln for 12 h at 37°C with shaking (200 rpm). Cultures were centrifuged at 7,000 ×g for 3 min, and supernatants were extracted twice with equal volumes of chloroform. The organic phase was collected, centrifuged at 13,000 ×g for 10 min, and evaporated under vacuum for 20 min using a centrifugal concentrator. Dried extracts were resuspended in methanol and spotted onto silica-gel TLC plates (Merck, Germany). Toxoflavin was visualized under UV light (365 nm), and bands were imaged. For quantitative analysis, methanol-dissolved extracts were measured spectrophotometrically at 393 nm (Li et al., 2019). The OD₆₀₀ of each culture was recorded in parallel, and toxoflavin levels were expressed as Abs₃₉₃/OD₆₀₀ to normalize for differences in growth. Data represent mean ± standard deviation (SD) from three independent experiments.

All experiments were conducted at least twice, with three biological replicates per treatment unless otherwise specified. Statistical analyses were performed using SAS (Statistical Analysis System; SAS Institute, Cary, NC, USA). Analysis of variance was carried out using the General Linear Model procedure, and mean separation was performed using Tukey’s Honestly Significant Difference (HSD) test at P < 0.05. Data are presented as the mean ± standard deviation.

The ΔntrC mutant exhibited a pronounced growth defect under nitrogen-limited conditions compared with the wild-type and complemented (CntrC) strains. In nutrient-rich LB medium, all strains displayed similar growth kinetics, indicating that NtrC is dispensable for proliferation when nitrogen is readily available (Fig. 1A). In contrast, under nitrogen-limiting conditions in MM supplemented with a single nitrogen source, the mutant showed a marked impairment. When either ammonium sulfate (MM-AS) or glutamine (MM-Gln) was supplied at 3 mM, the ΔntrC strain consistently failed to reach the optical densities achieved by the WT and CntrC strains during 48 h of incubation (Fig. 1B and C). Restoration of growth in the complemented strain confirmed that this phenotype resulted specifically from ntrC disruption. These findings demonstrate that NtrC is indispensable for efficient assimilation of ammonium and glutamine—key substrates in bacterial nitrogen metabolism—and highlight its critical role in sustaining growth under nitrogen scarcity that mimics the host apoplastic environment.

Loss of ntrC significantly compromised the virulence of B. glumae in both flowering and seedling infection assays. In the panicle assay, plants inoculated with the ΔntrC mutant exhibited visibly reduced symptoms compared with those inoculated with the wild-type, whereas the complemented strain restored symptom severity to wild-type levels (Fig. 2A-C). Quantitative scoring using the panicle blight index confirmed a substantial reduction in disease severity for the mutant, indicating that NtrC is required for effective colonization and symptom development in rice spikelets.

Consistent results were obtained in the seedling blight assay. Wild-type and CntrC infections caused pronounced inhibition of shoot and root elongation, while seedlings inoculated with the ΔntrC mutant showed only mild stunting and limited root necrosis (Fig. 2D-F). Collectively, these observations establish that NtrC is indispensable for the full virulence of B. glumae BGR1 during both early and late stages of infection. The restoration of pathogenicity in the complemented strain further confirms that the observed attenuation results directly from ntrC deletion rather than secondary mutations.

Although the ΔntrC mutant caused markedly milder disease symptoms, it was unclear whether this attenuation resulted from impaired bacterial proliferation within host tissues. To address this, bacterial population dynamics were quantified in infected rice seedlings.

However, enumeration of bacterial populations revealed no significant differences in CFU counts between the wild-type and ΔntrC strains at either the initial inoculation (0 dpi) or at 10 dpi (Fig. 3A and B). These results indicate that NtrC does not affect bacterial survival or multiplication in planta. The reduced virulence of the ΔntrC mutant therefore reflects a regulatory impairment that disrupts the expression or activation of virulence-associated traits, rather than a defect in colonization or persistence within host tissues.

Efficient colonization of host tissues by B. glumae relies on motility for surface exploration and biofilm formation for stable attachment and persistence. When tested in MM-Gln, the ΔntrC mutant exhibited reduction in swimming motility compared with the wild-type strain (Fig. 4A and B). Motility was restored in the complemented strain (CntrC), reaching levels comparable to wild-type.

Similarly, biofilm formation was reduced in the ΔntrC mutant compared with the wild-type under nitrogen-limited conditions (Fig. 4C and D). Because the initial inoculum (OD₆₀₀ = 0.05) might allow growth defects to influence attachment, we repeated the assay using cultures adjusted to an OD₆₀₀ of 1.0 prior to inoculation to minimize growth-dependent effects. The ΔntrC mutant still showed significantly reduced biofilm formation under these conditions, confirming that this phenotype is not solely attributable to impaired growth (Supplementary Fig. 1). These results demonstrate that NtrC positively regulates both swimming motility and biofilm formation under nitrogen-limited conditions. Given the importance of these traits during the early stages of rice infection, the observed attenuation of virulence in the ΔntrC mutant likely results, at least in part, from impaired colonization capacity.

To determine whether NtrC also affects virulence factors associated with later stages of infection, extracellular protease secretion and toxoflavin biosynthesis were examined. The ΔntrC mutant exhibited protease activity comparable to that of the wild-type and CntrC strains, as indicated by similar clear zone diameters on skim milk agar plates (Fig. 5A and B). These findings suggest that NtrC does not contribute to the regulation of extracellular protease production, a function commonly linked to host tissue maceration.

In contrast, toxoflavin synthesis was markedly reduced in the ΔntrC mutant compared with wild-type. The TLC analysis revealed substantially weaker toxoflavin bands in mutant extracts, whereas complementation restored the band intensity to near WT levels (Fig. 5C). Quantitative spectrophotometric measurements at 393 nm further confirmed a significant decrease in toxoflavin accumulation in the mutant (Fig. 5D). Together, these results identify NtrC as a positive regulator of toxoflavin biosynthesis but not of extracellular protease secretion, underscoring its selective role in modulating key secondary metabolites essential for B. glumae virulence.