The Role of Biofilm Formation by Paracidovorax citrulli in the Infection Process of Hami Melon

Abstract

It has been well established that biofilm formation plays a critical role in the pathogenesis of various plant pathogenic bacteria. However, research on this process in Paracidovorax citrulli, the causal agent of bacterial fruit blotch (BFB) in cucurbits, remains limited. Through screening of the infection pathways of P. citrulli in sweet melon leaves, observing biofilm formation morphology at bacterial colonization sites, and detecting the activities of pathogenicity-related enzymes, this study revealed that P. citrulli readily colonizes Hami melon vascular tissues following inoculation via petiole immersion, petiole dipping, or vine injection. Dense biofilms were observed within the vascular bundles of symptomatic leaf veins. Furthermore, P. citrulli was confirmed to secrete cellulase and pectinase, with enzymatic activities increasing progressively as disease severity intensified. These findings suggest that BFB development in Hami melon is likely associated with the synergistic action of P. citrulli, biofilm-mediated occlusion of xylem vessels and hydrolytic degradation of plant cell walls, which may contribute to initial water-soaked lesions and subsequent vein-associated necrosis in leaf tissues. This study provides a theoretical foundation for further elucidation of the pathogenic mechanisms of P. citrulli.

1. Introduction

A biofilm is a structured, membrane-like microbial community formed when bacteria adhere to biotic or abiotic surfaces and secrete extracellular polymeric substances (EPSs) to enclose themselves, which includes extracellular polysaccharides, proteins, extracellular DNA (eDNA), and other components [1]. Extracellular polysaccharides play a central role in the formation of the three-dimensional architecture of biofilms. They confer adhesive properties to biofilms, not only facilitating nutrient acquisition by bacterial cells but also protecting against the penetration of antimicrobial agents from the external environment into the biofilm interior, thereby serving as a core component of the biofilm’s protective barrier [2]. Biofilms contain a diverse array of proteins, including lectins that mediate bacterial attachment to EPS and hydrolytic enzymes with functional roles, some of which exhibit toxic activity. These proteins are primarily involved in the dynamic regulation of biofilm structure and the expansion of its functional capabilities. Extracellular DNA (eDNA) carries a negative charge and interacts with receptors on the substrate surface, promoting microbial adhesion [3]. Moreover, eDNA binds to proteins and participates in the formation of the three-dimensional architecture of biofilms [4].

It represents the predominant mode of bacterial existence in natural environments and serves as a key survival strategy and virulence determinant for pathogenic bacteria during host colonization. Biofilms not only enhance bacterial resistance to chemical agents and host immune defenses, but also coordinate the expression of virulence factors through collective behavior, thereby serving as a critical contributor to pathogenesis during infection [5,6]. Erwinia amylovora is a typical vascular pathogen capable of infecting multiple Rosaceae host species, including Malus domestica and Pyrus pyrifolia. Curli fibers, as components of the biofilm’s extracellular matrix, facilitate bacterial adhesion to and invasion of host cells. Amylovoran and Levan, both integral components of the biofilm, function as key virulence factors. Mutant strains unable to produce amylovoran lose pathogenicity and fail to disseminate within the plant xylem vessels. Similarly, Levan-deficient mutants exhibit delayed symptom development in host plants [7]. E. amylovora mutants with reduced biofilm-forming capacity fail to colonize the xylem vessels, and the pathogen remains localized at the inoculation site when introduced into leaves, significantly impeding systemic spread within the host plant [8]. In susceptible potato plants, the extracellular polysaccharides of Clavibacter michiganensis subsp. sepedonicus strongly bind to protoplast surfaces and microsomal membranes, promoting pathogen adhesion [9]. The bacterium proliferates extensively within tomato xylem vessels, forming a biofilm that induces degradation of vascular bundle tissues, leading to stem cankers and leaf wilting, and ultimately resulting in whole-plant death [10,11]. Furthermore, the extracellular polysaccharides exacerbate wilting symptoms. When introduced into wounds, these polysaccharides induce distinct ulceration symptoms in tomato stems [12].

Biofilms are structured, multicellular bacterial communities that form through coordinated cellular cooperation. This process is primarily regulated by quorum sensing (QS), a mechanism that facilitates intercellular communication via the secretion and detection of self-produced signaling molecules, enabling population density-dependent coordination of gene expression [1,13]. In Pseudomonas syringae pv. tomato, the coordination between biofilm formation and virulence factor expression is primarily fine-tuned by the second messenger c-di-GMP, with the bifunctional protein Chp8 serving as a central regulatory component of this pathway. Upon plant infection, the host produces the flavonoid phloretin, which activates chp8 expression. Activation of Chp8 drives c-di-GMP synthesis through the HrpSR-HrpL signaling cascade, leading to enhanced production of extracellular polysaccharides and robust biofilm development, while concurrently suppressing fliC expression. This regulatory strategy enables the pathogen to evade salicylic acid -mediated host immunity by reducing pathogen-associated molecular pattern (PAMP) recognition and facilitates a transition to the biofilm lifestyle, thereby promoting successful host colonization and full virulence expression [14]. In Pectobacteria, bacterial aggregation leads to elevated levels of acyl-homoserine lactones (AHLs), which are recognized and bound by the transcriptional regulator ExpR to form the AHL-ExpR complex. This complex suppresses the expression of rsmA, thereby reducing the abundance of the negative regulatory protein RsmA and relieving its repression on mRNAs encoding plant cell wall-degrading enzymes (PCWDEs). Consequently, the translation of PCWDEs is activated, facilitating pathogenicity [15]. In Xanthomonas oryzae pv. oryzae, DSF-family signals that regulate biofilm formation similarly enhance xylanase activity in the pathogen [16].

Paracidovorax citrulli is the causal agent of bacterial fruit blotch (BFB), a severe disease affecting cucurbit crops. In 1992, Willems et al. [17] designated the bacterium as Acidovorax avenae subsp. citrulli based on rRNA-rRNA and DNA-DNA hybridization data. In 2008, it was reclassified as A. citrulli due to distinct phenotypic and genetic characteristics [18]. In 2022, Du et al. [19] conducted a comparative genomic analysis of the genus Acidovorax and proposed two novel genera: Paracidovorax gen. nov. and Paenacidovorax gen. nov. Phylogenomic evidence revealed that A. citrulli clusters within the Paracidovorax clade, leading to its formal reclassification into this newly established genus, with the updated nomenclature P. citrulli [20,21,22]. Leaves of seedlings infected with P. citrulli exhibit characteristic water-soaked lesions, with necrotic lesions developing along leaf veins. Water-soaked lesions on fruits typically coalesce into large fruit spots or are characterized by small, sunken lesions and internal fruit rot [23]. In infectious plant diseases, water-soaked symptoms are associated with enzymes produced by pathogenic bacteria and lower fungi. Pathogen-secreted enzymes degrade plant cell wall structures, leading to cytoplasmic leakage and accumulation of cellular contents in the apoplastic space, which results in the characteristic semi-transparent, water-soaked appearance of infected tissues. Notably, pectinases secreted by Dickeya and Pectobacterium species are key virulence factors responsible for the development of these symptoms [24]. Enzymes are key components of the pathogenic bacterial arsenal, enabling the direct degradation of plant physical and chemical defense barriers, thereby promoting pathogen invasion, colonization, and spread within host tissues, and ultimately driving the development of disease symptoms. The pectate lyase, polygalacturonase, and protease secreted by E. amylovora [25], along with the cellulases (ClsA and CbsA) and xylanase (Xyn) produced by Xanthomonas oryzae pv. oryzae [26], are key virulence factors during pathogen infection. A comprehensive understanding of the types and functions of enzymes secreted by pathogenic bacteria, their regulatory mechanisms, and their interactions with the plant defense system is crucial for the development of novel disease control strategies.

It has been well established that biofilm formation in pathogenic bacteria plays a crucial role in the pathogenesis process and is coordinately regulated with the expression of virulence factors through QS [13,27]. However, studies on P. citrulli biofilms have predominantly focused on assessing their in vitro formation capacity, and, to date, no reports have documented biofilm formation by this pathogen within plant hosts [28,29,30,31]. Li Siqi et al. [32] demonstrated that the severity of leaf disease is positively correlated with the amount of biofilm formed by the pathogen on the leaf surface, as revealed by crystal violet staining. However, the study was based solely on macroscopic assessment and did not include microscopic visualization of biofilm structures. To date, no studies have reported whether this pathogen forms biofilms in Hami melon leaves during infection, the association between biofilm formation and disease progression—given that previous research has been restricted to in vitro systems—or whether P. citrulli biofilms regulate the secretion of hydrolytic enzymes during infection, thereby influencing pathogenicity. Building on these knowledge gaps, this study employed Hami melon leaves as the experimental system to investigate the role of P. citrulli biofilms during infection by examining: (i) the in planta colonization morphology of P. citrulli; (ii) the association between biofilm formation and disease development; and (iii) the activity of pathogenicity-related enzymes during leaf infection.

2. Materials and Methods

2.1. Bacterial Strains and Growth Conditions

The test strain P. citrulli XJ05-1 was provided by the Laboratory of Biological Control of Plant Diseases at Shihezi University. Pathogenic bacteria preserved at −80 °C in 20% glycerol were inoculated onto KBA medium and incubated at 28 °C for 2 days. A single colony was then transferred to a 50 mL conical flask containing 20 mL of LB medium and cultured at 28 °C with shaking at 160 rpm for 24 h, during which the culture reached the stationary growth phase with maximum cell density for subsequent use [33].

2.2. Plant Growth Conditions and Inoculation Methods

The Xinmi Za No. 11 honeydew melon variety, provided by the Agricultural Science Institute of the 6th Division, Xinjiang Production and Construction Corps, was used as the host plant for P. citrulli. Healthy and plump Hami melon seeds were fully submerged in 5% sodium hypochlorite solution for 10 min and subsequently rinsed five times with sterile water (2 min per rinse). The disinfected seeds were then air-dried under a laminar flow hood and reserved for use. A substrate mixture of nutrient soil and vermiculite (3:1, v/v) was prepared and transferred into plastic pots, each equipped with a bottom tray to facilitate capillary watering. Seeds were sown in the substrate and grown in a controlled-environment growth chamber at 26 °C under a 12 h light/12 h dark photoperiod. Plants were irrigated every four days. Plants at the five-true-leaf stage were used for subsequent experiments.

P. citrulli cultures were centrifuged at 8000 r/min for 5 min to pellet the bacterial cells. The bacterial pellet was resuspended in sterile water, centrifuged again, and resuspended. Then, the OD₆₀₀ of the bacterial suspension was measured using a microplate reader (Multiskan FC, Thermo Fisher Scientific, Waltham, MA, USA), and the cell density was adjusted with sterile water to an OD₆₀₀ of 0.3 (≈10⁸ CFU/mL) [34] for subsequent experiments.

Inoculation methods were conducted in 8 distinct ways: Treatment a: Injection inoculation—The pathogen suspension was injected into the leaf tissue of melon leaves using a sterile micro-syringe; Treatment b: Needle prick inoculation—The leaf surface was gently pricked with a sterile needle, and 5 μL of bacterial suspension was aspirated and dotted onto the micro-wound; Treatment c: Infiltration inoculation—A microsyringe loaded with 5 μL of pathogen suspension was used without a needle; the tip was gently pressed against the adaxial leaf surface while slight counterpressure was applied on the abaxial side with a finger, allowing the inoculum to be introduced into the leaf tissue. Treatment d: Droplet inoculation—A 5 μL droplet of pathogen suspension was placed onto the surface of the melon leaf. Following both treatments, plants were maintained under high humidity for 12 h. Treatment e: Petiole immersion—The leaf petiole was immersed in the bacterial suspension. Treatment f: Petiole dip inoculation—The wounded end of the petiole was dipped into the bacterial suspension, then wrapped with moist sterile cotton and placed in a sterile Petri dish lined with wet filter paper. Treatment g: Foliar spray inoculation—The leaf surface was uniformly sprayed with bacterial suspension using a fine mist sprayer (Bluethin, Shenzhen, China) until visibly wet. Treatment h: Vine injection inoculation method—Select a vine shoot with five true leaves. Using a micro syringe, inject 5 μL of bacterial suspension into the vascular tissue of the vine shoot between the first and second true leaves. Inoculated melon plants were transferred to a 28 °C incubator and maintained under a 12 h light/12 h dark photoperiod for observation. All treatments included a control group treated with sterile water. Each treatment group was subjected to six replicates per trial, with the experiment repeated three independent times.

2.3. Relationship Between Pathogenic Bacterial Concentration and Disease Onset

The leaf petiole immersion method was employed. Leaves exhibiting consistent growth status and appearing fresh and healthy were selected, and their petioles were immersed in bacterial suspensions of P. citrulli at concentrations of 10², 10⁴, 10⁶ and 10⁸ CFU/mL, respectively. The samples were incubated at a constant temperature of 28 °C, and disease development on the leaves was monitored continuously. With sterile water as the control, each group was set up with six replicates, and the experiment was conducted three independent times.

2.4. Determination of Biofilm Formation in Infected Melon Leaves

2.4.1. Preliminary Observation of Biofilm Formation on Diseased Leaves

At 24 and 96 h post-inoculation, the melon leaves were completely immersed in a 1% crystal violet solution (Solarbio, G1062, Beijing, China) for 20 min. The leaves were rinsed three times with water to remove excess dye solution, and biofilm formation on the inoculated leaves was then observed. Concurrently, leaf tissue from the margin between diseased and healthy regions was excised and placed on a water droplet at the center of a microscope slide. A coverslip was carefully applied and gently pressed to eliminate air bubbles within the tissue. The samples were then observed and imaged using an Olympus BX61 microscope (Olympus Corporation, Tokyo, Japan).

2.4.2. Scanning Electron Microscope Observation

Scanning electron microscopy (SEM) was employed for ultrastructural analysis to visualize the morphology of pathogenic bacteria colonizing the vascular tissues of melon leaves. Leaves exhibiting disease symptoms following inoculation with a 1 × 10⁸ CFU/mL bacterial suspension in Section 2.3 were selected. Leaf surfaces were rinsed with sterile water and air-dried. Diseased vascular tissues were excised horizontally into 1 mm × 5 mm segments using a sterile blade. Samples were fixed at room temperature for 2 h in 5% glutaraldehyde prepared in 0.1 mol/L phosphate buffer (pH 7.2). Following fixation, samples were washed three times with the same buffer (15 min per wash), then dehydrated through a graded ethanol series (30%, 50%, 70%, 80%, 90%, and 95%; 15 min each), and finally subjected to two changes of absolute ethanol (20 min each). Critical point drying was carried out using a critical point dryer (EM CPD300, Leica, Wetzlar, Germany). Dried tissue sections were mounted on aluminum stubs and sputter-coated with gold. Scanning electron microscopy (SEM) imaging was performed using an SU8010 microscope (Hitachi, Tokyo, Japan) at an accelerating voltage of 2.0 kV under various magnifications.

2.4.3. Pathogen Verification from Symptomatic Leaves

PCR amplification was performed using the specific primers WFB1 (5′-GACCAGCCACACTGGAC-3′) and WFB2 (5′-CTGCCGCACTCCAGCGA-3′) [35] to confirm the presence of the putative pathogen in symptomatic leaves. These primers target a 360 bp fragment within the 16S rDNA region of P. citrulli, corresponding to positions 293–310 and 652–669 of the reference sequence AF137506 (1481 bp). Diseased leaf tissue was surface-sterilized with 75% ethanol for 3 min, rinsed three times with sterile water, and transferred to a centrifuge tube containing 700 µL of sterile water. The tissue was thoroughly ground, allowed to stand for 20 min, and then centrifuged at 8000 r/min for 5 min. Given the simple cellular structure of bacteria, genomic DNA is readily released during the pre-denaturation step of PCR. Consequently, DNA extraction and purification are not required prior to amplification; instead, the centrifuged supernatant can be used directly as a template in PCR assays. A 1 µL aliquot of the sample was transferred into a PCR tube and mixed with 1 µL of primer WFB1, 1 µL of primer WFB2, 5 µL of 2 × PCR Mix (GDSBio, Guangzhou, China), and 2 µL of ddH₂O. The thermal cycling program on a T100 PCR instrument (BIO-RAD) was set as follows: initial denaturation at 94 °C for 3 min; 30 cycles of 94 °C for 30 s, 55 °C for 30 s, and 72 °C for 1 min; followed by a final extension at 72 °C for 10 min. After amplification, 5 µL of the PCR product was loaded onto a 1% (w/v) agarose gel in 1 × TBE buffer. Electrophoresis was carried out at 120 V for 15 min, using a 2 kb DNA Marker (MD114, Tiangen, Beijing, China) as molecular weight standard. Amplified bands were visualized under Universal Hood II Gel Doc System (Bio-Rad Laboratories, Hercules, CA, USA).

Additionally, the grinding suspension was serially diluted and plated onto AacSM selective agar medium (P. citrulli selective medium, Na₂PO₄∙12H₂O 2.5 g/L, KH₂PO₄ 0.5 g/L, (NH₄) ₂SO₄ 2 g/L, MgSO₄∙7H₂O 0.029 g/L, CaCl₂∙2H₂O 0.067 g/L, Na₂HMo₄∙H₂O 0.025 g/L, Yeast extract 0.01 g/L, Ammonium adipate 10 g/L, Bromothymol blue 0.0125 g/L, Agar 15 g/L, Ampicillin 0.02 g/L, Cycloheximide 0.03 g/L, phenethicillin potassium 0.1g/L, novobiocin 0.03 g/L) [36]. The plates were incubated at 40 °C for 4 days, and colony morphology, particularly pigmentation, was observed.

2.5. Determination of Enzymes Activities

The enzyme profiles of P. citrulli were determined using specific enzyme-screening media through the halo formation assay. The fermentation broth of P. citrulli was centrifuged at 8000 r/min for 10 min at 4 °C. The supernatant was collected and filtered through a 0.22 µm bacterial filter to obtain the planktonic extracellular crude enzyme fraction. The pellet was resuspended in 20 mL of sterile water and centrifuged again under the same conditions (8000 r/min, 10 min, 4 °C). This washing step was repeated once more prior to cell disruption. Cells were disrupted using a sonicator (SCIENTZ-IID, Ningbo Xinzhi Biotechnology Co., Ltd., Ningbo, China) set at 200 W with cycles of 5 s pulse and 5 s rest for a total duration of 10 min on ice. Following disruption, the lysate was centrifuged at 8000 r/min for 10 min at 4 °C, and the resulting supernatant was passed through a 0.22 µm filter to yield the planktonic intracellular crude enzyme fraction. A 1% (v/v) inoculum of the P. citrulli culture from step 2.1 was transferred into a 24-well plate containing 1 mL of fresh LB medium per well and statically incubated at 28 °C for 24 h. After incubation, the supernatant was carefully removed, and the wells were gently rinsed twice with sterile water to remove non-adherent planktonic cells. Biofilm-associated cells attached to the well walls were then detached by vigorous washing with sterile water and collected as the biofilm-state cell fraction.

Using a 6 mm diameter punch, make holes in the cellulase detection medium (CMC-Na as substrate, visualized by Congo red staining) [37], pectinase detection medium (pectin as substrate, visualized by Congo red staining) [38], xylanase detection medium (xylan, visualized by Congo red staining) [39], β-glucanase detection medium (β-glucan as substrate, visualized by Congo red staining) [40], chitinase detection medium (colloidal chitin as substrate, no staining required) [41], protease detection medium (skimmed milk powder as substrate, no staining required) and amylase detection medium (soluble starch as substrate, no staining required) [42]. The planktonic extracellular crude enzyme solution and biofilm-state cells of P. citrulli were separately inoculated into the corresponding wells. Additionally, 200 µL of the planktonic intracellular crude enzyme solution was added to Oxford cups placed on the assay plates. All plates were incubated at 28 °C for 4 days. Formation of a transparent halo around the wells or cups was recorded as a positive enzymatic reaction. For amylase detection, plates were stained with iodine solution after incubation to visualize residual starch, followed by ethanol decolorization to enhance contrast prior to observation.

2.6. Measurement of Enzyme Activity in Diseased Leaf Tissues Following Inoculation

Leaf tissues from diseased areas were collected using a 6 mm diameter cork borer, with three leaf discs pooled as one biological replicate. The samples were transferred to 1.5 mL centrifuge tubes containing 800 µL of sterile water and homogenized thoroughly with sterile grinding rods. After incubation at room temperature for 20 min, the suspensions were centrifuged at 8000 r/min and 4 °C for 10 min. The resulting supernatants were filtered through a 0.22 µm bacterial filter to obtain crude enzyme extracts for enzymatic activity assays. Crude enzyme extracts from the control group were prepared using the identical procedure.

2.6.1. Cellulase Activity Assay

Cellulase activity was assayed using the 3,5-dinitrosalicylic acid (DNS) [43] method. A series of glucose standard solutions were prepared by adding 0.2, 0.4, 0.6, 0.8, 1.0, 1.2, 1.4, and 1.6 mL of 1 mg/mL glucose solution to separate 50 mL centrifuge tubes, respectively, and adjusting the final volume to 2 mL with deionized water. The mixtures were vortexed thoroughly, followed by the addition of 1.5 mL DNS reagent (Solarbio, D7800). The reaction mixtures were incubated in a boiling water bath for 15 min, cooled rapidly under running tap water, then supplemented with 10 mL deionized water and mixed well. Aliquots of 200 μL were transferred to a microplate for measurement of absorbance at 540 nm using a Varioskan Flash multimode microplate reader (ThermoFisher Scientific, Waltham, MA, USA). A blank control containing no glucose was included for baseline correction. A standard curve was constructed by plotting glucose concentration (mg/mL) on the x-axis against corresponding absorbance values.

0.5 mL of crude enzyme solution was mixed with 1.5 mL of CMC-Na solution, and the mixture was incubated at 50 °C for 30 min. After incubation, 1.5 mL of DNS reagent was added, followed by boiling water bath treatment for 15 min. The reaction was cooled rapidly, then 10 mL of deionized water was added, and the solution was mixed thoroughly. Absorbance at 540 nm was measured using the Varioskan Flash multimode microplate reader. The inactivated samples, prepared by boiling for 20 min, were used as the blank control. Each sample was analyzed in six replicates, and the experiment was independently repeated three times. Enzyme activity was defined as the amount of cellulase required to release 1 μg of glucose per minute from the substrate under standard assay conditions, and one unit of enzyme activity was expressed as U/mL.

U = A × N × 1000/(V × T)

(1)

A—glucose yield (mg); N—sample dilution factor; V—reaction mixture volume (mL); T—reaction time (min).

2.6.2. Pectinase Activity Assay

Pectinase activity was determined using the DNS method [43]. Aliquots of 0.1, 0.2, 0.3, 0.4, 0.5, 0.6, 0.7, 0.8, and 0.9 mL of 1 mg/mL galacturonic acid solution (Solarbio, G8120) were added to separate 50 mL centrifuge tubes. Deionized water was added to adjust the volume to 1 mL in each tube, followed by thorough mixing. Then, 3 mL of DNS reagent was added to each tube. The mixtures were incubated in a boiling water bath for 10 min, cooled rapidly under running water, diluted with 21 mL of deionized water, and mixed thoroughly. A 200 μL aliquot of each solution was transferred to a microplate and the absorbance at 540 nm (OD₅₄₀) was measured using a Varioskan Flash multi-mode microplate reader (ThermoFisher Scientific, Waltham, MA, USA). The tube without galacturonic acid served as the blank control. A standard curve was constructed by plotting galacturonic acid concentration (mg/mL) on the x-axis and the corresponding absorbance values on the y-axis.

The reaction was initiated by adding 0.8 mL of pectin solution (Solarbio, P8030) to 0.2 mL of crude enzyme solution, followed by incubation at 50 °C for 30 min. After incubation, 3 mL of DNS reagent was added, followed by heating in a boiling water bath for 10 min. The reaction mixture was cooled rapidly under running water, then diluted with 21 mL of deionized water and mixed thoroughly. The absorbance at 540 nm (OD540) was measured. An inactivated crude enzyme solution (boiled for 20 min) was used as the blank control. Each sample was analyzed in six replicates, and the experiment was independently repeated three times. One enzyme activity unit (U/mL) was defined as the amount of enzyme that catalyzes the hydrolysis of pectin to release 1 μg of galacturonic acid per minute under the assay conditions, using 1 mL of enzyme solution.

U = (X₁ − X₂) × N × 1000/(K × V × T)

(2)

X₁-absorbance of the experimental group; X₂—absorbance of the control group; N—sample dilution factor; K—slope of the galacturonic acid standard curve; V—reaction liquid volume (mL); T—response time (min).

2.7. Statistical Analysis

Statistical analyses were conducted using SPSS version 24. Prior to performing the t-test, the normality of data distribution was assessed using the Shapiro–Wilk test, and homogeneity of variance was evaluated using Levene’s test. All datasets satisfied the assumptions of normality and homogeneity of variance; therefore, independent samples t-tests were applied for pairwise comparisons, with statistical significance set at p < 0.05. Data are presented as mean ± standard error (SE).

3. Results

3.1. Inoculation Method and Symptom Manifestation

As shown in Figure 1, leaves inoculated via injection (a) and droplet inoculation (d) developed small, localized brown necrotic lesions restricted to the mesophyll tissue, exhibiting mild symptoms and slow lesion expansion. Needle-prick inoculation (b) resulted in the mildest symptoms, with only small pits at the inoculation site and no lesion spread. Infiltration inoculation (c) resulted in deep brown necrotic lesions confined to the inoculation site, which often led to perforation. In contrast, inoculation through petiole immersion (e), petiole dipping (f), foliar spraying (g), and vine injection (h) led to lesions that expanded along the vascular veins and displayed a water-soaked appearance. Notably, symptoms following petiole immersion, petiole dipping, leaf spraying, and stem injection appeared rapidly, with swift disease progression. These results suggest that P. citrulli readily colonizes the vascular bundles within leaf veins and is consistent with the characteristics of a xylem-limited endophytic pathogen.

3.2. Effect of Inoculation Concentration on the Development of P. citrulli Disease

Bacterial biofilm formation is a key pathogenic factor in vascular plant diseases, and crystal violet staining effectively visualizes these biofilms by imparting a distinct purple color. As shown in Figure 2, on the first day after inoculation, no visible symptoms were observed on the leaves of any treatment group, and following crystal violet staining, only the terminal ends of the petioles exhibited a purple coloration. By 3 days post-inoculation, water-soaked lesions began to appear on the leaves and spread progressively along the vascular veins. Lesions in the high-concentration treatment groups (10⁸ and 10⁶ CFU/mL) were slightly larger than those in the low-concentration groups (10⁴ and 10² CFU/mL), and the extent of crystal violet staining corresponded closely with the lesion size.

At 6 days post-inoculation, disease symptoms on leaves in the high-concentration treatment groups were markedly more severe than those in the low-concentration groups. Leaves inoculated with 10⁸ and 10⁶ CFU/mL developed extensive brown lesions, with lesion size slightly greater in the 10⁸ CFU/mL treatment compared to the 10⁶ CFU/mL treatment. In contrast, leaves exposed to 10⁴ and 10² CFU/mL exhibited smaller lesions primarily confined to the central leaf vein, though lesion size in the 104 CFU/mL treatment was slightly larger than that in the 10² CFU/mL treatment (Figure 2). Following crystal violet staining, the leaves inoculated with 10⁸ and 10⁶ CFU/mL exhibited extensive purple staining across nearly the entire leaf blade, whereas for those inoculated with 10⁴ and 10² CFU/mL, purple staining was predominantly restricted to the leaf tissues adjacent to the central midvein. The above results suggest that higher inoculation concentrations are associated with earlier attainment of the biofilm formation threshold by the bacteria within leaf tissues, which appears to correlate with faster symptom development.

3.3. Biofilm Formation During Bacterial Invasion

Leaves collected on the 1st and 3rd days post-inoculation were stained with crystal violet, prepared as microscope slides, and observed under a light microscope. On day 1, slight purple staining was detected within the vascular bundles of leaf tissues, indicating the initial formation of biofilms. By day 3, extensive and coalesced purple deposits were evident inside the leaves, with adjacent veins also exhibiting distinct purple staining (Figure 3).

Scanning electron microscopy revealed that bacterial cells within leaf vascular bundles had already aggregated into clusters prior to symptom onset (Figure 4B). At the initial stage of symptom development, numerous bacteria had accumulated and formed biofilms within the vascular tissues (Figure 4C). In the late stage of infection, dense biofilms were observed extensively colonizing the vascular bundles, which was accompanied by severe occlusion of the xylem vessels (Figure 4D).

3.4. Pathogen Verification in Diseased Leaves

Leaf tissues were sampled from both the lesion margin (interface between diseased and healthy tissues) and non-symptomatic regions of inoculated leaves for pathogen detection (Figure 5A). When the leaf tissue homogenates were diluted and plated on the selective medium AacSM [35], yellowish-green colonies emerged (Figure 5B), morphologically consistent with P. citrulli grown on the same medium. Furthermore, PCR amplification using the species-specific primers WFB1/WFB2 successfully amplified a 360 bp fragment of P. citrulli from both tissue sources [36] (Figure 5C). These results confirm that P. citrulli was present in the infected tissues and is the causal agent of the observed disease symptoms.

3.5. Enzyme Activity Detection

The enzyme-producing screening assay revealed that both intracellular and extracellular components of planktonic cells, as well as biofilm-associated bacteria, produced clear zones on the cellulase-containing agar plate. On the pectinase-containing agar plate, only biofilm cells formed a clear zone (Figure 6), indicating that P. citrulli in the planktonic state produces cellulase, whereas the biofilm state enables production of both cellulase and pectinase.

Leaf tissues from symptomatic sites were collected at 4, 24, and 96 h post-inoculation for enzyme activity assays. Endogenous production of cellulase and pectinase by plant leaves was detected; however, total enzyme levels following P. citrulli inoculation exceeded the plant-derived baseline. The difference was attributed to pathogen-secreted enzymes. The results showed that plant-derived enzyme levels remained constant throughout infection, whereas cellulase and pectinase activities attributable to P. citrulli increased progressively (Figure 7 and Figure 8) suggest that the increasing activities of these enzymes are consistent with a role in the pathogenicity of P. citrulli during biofilm-mediated infection.

4. Discussion

Biofilms represent a fundamental pathogenic strategy employed by plant pathogenic bacteria, enabling their successful survival and colonization in host tissues such as leaves, stems, seeds, and roots [44]. The formation and structural architecture of biofilms are modulated by factors including plant species, specific tissue types, and the site of colonization [45]. Among the eight inoculation methods tested, P. citrulli induced the most rapid leaf disease development and exhibited typical vein-constrained lesion expansion when applied via petiole immersion, petiole dipping, or stem injection. These symptoms are consistent with previously reported field symptoms of leaf disease [46], suggesting that P. citrulli readily colonizes xylem vessels within vascular bundles and aligns with the characteristics of a vascular bundle-inhabiting pathogen. This finding aligns with the results reported by Wei Zhongren [47], who used GFP gene labeling to demonstrate that P. citrulli can colonize the vascular tissues of hypocotyls in melon rootstock seedlings and cause systemic plant infection.

Studies have demonstrated that the deletion of genes hrcS [29], pstS [30], and pilA [31], as well as the genes hrpW [22], glpR [28], rpoN1 [48] and furA (Ferric Uptake Regulator) [49], leads to reduced biofilm formation in P. citrulli, whereas deletion of the genes clpA [50], gptT [51], and OmpRAc/EnvZAc [52] results in enhanced biofilm production. These findings indicate the presence of a complex genetic regulatory network that fine-tunes biofilm formation in P. citrulli. However, all such studies have been conducted under in vitro conditions, with biofilm phenotypes assessed solely through quantitative measurements. To date, the role of biofilm formation by P. citrulli in plant colonization and its contribution to pathogenicity during host infection remains uncharacterized. This study demonstrates that the inoculation concentration gradient assay reveals a positive correlation between inoculum dosage and the rate of disease onset. High-concentration inocula (10⁸, 10⁶ CFU/mL) facilitated rapid biofilm formation within the vascular bundles and induced severe disease symptoms, whereas low-concentration inocula (10⁴, 10² CFU/mL) required a longer period to reach the threshold necessary for biofilm establishment. This phenomenon parallels the infection mechanisms of several well-characterized vascular plant pathogenic bacteria. For example, X. fastidiosa [53], which causes Pierce’s disease of grapevine and citrus variegated chlorosis, Ralstonia solanacearum [54], responsible for bacterial wilt in tomato, and Pantoea stewartii subsp. stewartii [55], the causal agent of Stewart’s wilt in corn, all exhibit density-dependent infectivity. These pathogens form biofilms within the plant vascular system, where high-density biofilm accumulation obstructs host xylem vessels, leading to leaf wilting and necrosis [6]. These observations support the idea that biofilm formation may play an important role in P. citrulli overcoming host defenses and establishing stable colonization sites. Microscopic observations and molecular validation clearly demonstrated the dynamic biofilm formation process of P. citrulli. During the early infection stage (1 day post-inoculation), the pathogen has already completed initial colonization within the vascular bundles and formed minimal biofilm structures, while no visible symptoms are yet apparent on the leaves. As the disease progresses (3 days post-inoculation and beyond), biofilms accumulate extensively in the vascular tissues, leading to progressive occlusion of xylem conduits, accompanied by the emergence of water-soaked lesions or leaf wilting. This pattern closely resembles the pathogenic mechanisms of established vascular pathogens [53,54,55], further underscoring the central role of biofilm formation in vascular disease development.

The formation of water-soaked lesions in plant tissues typically results from compromised cell integrity, leading to the leakage of intracellular water and cellular contents. During infection, pathogenic bacteria secrete hydrolases that degrade the host cell wall, which may contribute to the development of water-soaked lesions in plant tissues. A representative example is the genus Erwinia, which produces multiple cell wall-degrading enzymes during infection. These enzymes serve as primary virulence factors, facilitating tissue maceration and enabling bacterial invasion into degraded plant tissues. Pectin-degrading enzymes purified from the secretions of Dickeya and Pectobacterium induce water-soaked lesions on healthy host tissues that resemble those observed during natural soft rot infection [24]. It is noteworthy that hydrolase activity is directly associated with the infection progression and disease severity of plant pathogens. In Xylella fastidiosa, endoglucanase influences the development rate of Pierce’s disease in grapevines [56], while the accumulation of secreted lipase/esterase (LesA) in leaf tissues correlates positively with symptom severity [57]. Similarly, Liu Xiaoyan et al. [58] reported that following inoculation of Sclerotinia sclerotiorum into oilseed rape, the pathogen’s cellulase and pectinase activities were significantly positively correlated with lesion length—higher enzyme activity corresponded to greater lesion expansion in the host plant. This pattern is also observed in P. citrulli: enzyme activity assays revealed differences in the secretion of pathogenicity-related enzymes between biofilm and planktonic cells. Biofilm-forming cells not only secrete cellulase but also produce pectinase, whereas planktonic cells exhibit detectable cellulase activity only. This phenotypic divergence suggests the presence of a molecular regulatory mechanism governing the expression of pathogenicity-associated enzymes within the biofilm microenvironment. Following inoculation of P. citrulli onto honeydew melon leaves, cellulase and pectinase activities were found to increase progressively over time, indicating that these enzymes play a pathogenic role during the infection process. Previous studies have demonstrated that loss of cellulase activity results in a significant decrease in bacterial pathogenicity [59], further supporting the essential role of these two enzymes during the infection process.

5. Conclusions

This study demonstrates that P. citrulli readily colonizes the vascular bundle tissues of Hami melon leaves, and the symptoms observed in infected leaves are associated with biofilm formation. Furthermore, the bacterium was found to secrete cellulase and pectinase during the infection process. These findings suggest that the pathogenicity of P. citrulli toward Hami melon leaves may be linked to both its ability to form biofilms and the secretion of cell wall-degrading hydrolases. Biofilms may obstruct host vascular bundles through spatial occupation and could also influence the environmental adaptability and pathogenicity of the pathogen, potentially through modulation of virulence factor expression, including hydrolytic enzymes. This provides a theoretical foundation for elucidating the pathogenic mechanism of P. citrulli.