The article is part of the research topic “Improving the resistance of legumes to pathogens and pests”, view all 5 articles
       The causal agent of the fungal plant disease necrosis Sclerotinia sclerotiorum (Lib.) de Bary uses a multi-tiered strategy to infect different host plants. This study proposes the use of the diamine L-ornithine, a non-protein amino acid that stimulates the synthesis of other essential amino acids, as an alternative management strategy to enhance the molecular, physiological and biochemical responses of Phaseolus vulgaris L. to white mold caused by Pseudomonas sclerotiorum. In vitro experiments showed that L-ornithine significantly inhibited the mycelial growth of S. pyrenoidosa in a dose-dependent manner. Moreover, it could significantly reduce the severity of white mold under greenhouse conditions. Furthermore, L-ornithine stimulated the growth of the treated plants, indicating that the tested concentrations of L-ornithine were not phytotoxic to the treated plants. In addition, L-ornithine enhanced the expression of non-enzymatic antioxidants (total soluble phenolics and flavonoids) and enzymatic antioxidants (catalase (CAT), peroxidase (POX), and polyphenol oxidase (PPO)), and increased the expression of three antioxidant-related genes (PvCAT1, PvSOD, and PvGR). Furthermore, in silico analysis revealed the presence of a putative oxaloacetate acetylhydrolase (SsOAH) protein in the S. sclerotiorum genome, which was highly similar to the oxaloacetate acetylhydrolase (SsOAH) proteins of Aspergillus fijiensis (AfOAH) and Penicillium sp. (PlOAH) in terms of functional analysis, conserved domains, and topology. Interestingly, the addition of L-ornithine to potato dextrose broth (PDB) medium significantly decreased the expression of SsOAH gene in S. sclerotiorum mycelia. Similarly, exogenous application of L-ornithine significantly decreased the expression of SsOAH gene in fungal mycelia collected from treated plants. Finally, L-ornithine application significantly decreased oxalic acid secretion in both PDB medium and infected leaves. In conclusion, L-ornithine plays a key role in maintaining the redox status as well as enhancing the defense response of infected plants. The results of this study may help in developing innovative, environmentally friendly methods to control white mold and mitigate its impact on bean production and other crops.
       White mold, caused by the necrotrophic fungus Sclerotinia sclerotiorum (Lib.) de Bary, is a devastating, yield-reducing disease that poses a serious threat to global bean (Phaseolus vulgaris L.) production (Bolton et al., 2006). Sclerotinia sclerotiorum is one of the most difficult soil-borne fungal plant pathogens to control, with a broad host range of over 600 plant species and the ability to rapidly macerate host tissues in a non-specific manner (Liang and Rollins, 2018). Under unfavourable conditions, it undergoes a critical phase of its life cycle, remaining dormant for long periods of time as black, hard, seed-like structures called ‘sclerotia’ in the soil or as white, fluffy growths in the mycelium or stem pith of infected plants (Schwartz et al., 2005). S. sclerotiorum is capable of forming sclerotia, which allows it to survive in infected fields for long periods of time and to persist during disease (Schwartz et al., 2005). Sclerotia are rich in nutrients, can persist in the soil for long periods, and serve as the primary inoculum for subsequent infections (Schwartz et al., 2005). Under favorable conditions, sclerotia germinate and produce airborne spores that can infect all above-ground parts of the plant, including but not limited to flowers, stems, or pods (Schwartz et al., 2005).
       Sclerotinia sclerotiorum uses a multi-tiered strategy to infect its host plants, involving a series of coordinated events from sclerotial germination to symptom development. Initially, S. sclerotiorum produces suspended spores (called ascospores) from mushroom-like structures called apothecia, which become airborne and develop into non-motile sclerotia on infected plant debris (Bolton et al., 2006). The fungus then secretes oxalic acid, a virulence factor, to control plant cell wall pH, promote enzymatic degradation and tissue invasion (Hegedus and Rimmer, 2005), and suppress the host plant’s oxidative burst. This acidification process weakens the plant cell wall, providing a favorable environment for the normal and efficient operation of fungal cell wall degrading enzymes (CWDEs), allowing the pathogen to overcome the physical barrier and penetrate the host tissues (Marciano et al., 1983). Once penetrated, S. sclerotiorum secretes a number of CWDEs, such as polygalacturonase and cellulase, which facilitate its dissemination in infected tissues and cause tissue necrosis. The progression of lesions and hyphal mats leads to the characteristic symptoms of white mold (Hegedus and Rimmer, 2005). Meanwhile, host plants recognize pathogen-associated molecular patterns (PAMPs) through pattern recognition receptors (PRRs), triggering a series of signaling events that ultimately activate defense responses.
       Despite decades of disease control efforts, shortages of adequate resistant germplasm remain in bean, as in other commercial crops, due to the resistance, survival, and adaptability of the pathogen. Disease management is therefore extremely challenging and requires an integrated, multifaceted strategy that includes a combination of cultural practices, biological control, and chemical fungicides (O’Sullivan et al., 2021). Chemical control of white mold is the most effective because fungicides, when applied correctly and at the right time, can effectively control the spread of the disease, reduce the severity of infection, and minimize yield losses. However, overuse and overreliance on fungicides can lead to the emergence of resistant strains of S. sclerotiorum and negatively impact non-target organisms, soil health, and water quality (Le Cointe et al., 2016; Ceresini et al., 2024). Therefore, finding environmentally friendly alternatives has become a top priority.
       Polyamines (PAs), such as putrescine, spermidine, spermine, and cadaverine, can serve as promising alternatives against soil-borne plant pathogens, thereby completely or partially reducing the use of hazardous chemical fungicides (Nehela et al., 2024; Yi et al., 2024). In higher plants, PAs are involved in many physiological processes including, but not limited to, cell division, differentiation, and response to abiotic and biotic stresses (Killiny and Nehela, 2020). They can act as antioxidants, help scavenge reactive oxygen species (ROS), maintain redox homeostasis (Nehela and Killiny, 2023), induce defense-related genes (Romero et al., 2018), regulate various metabolic pathways (Nehela and Killiny, 2023), modulate endogenous phytohormones (Nehela and Killiny, 2019), establish systemic acquired resistance (SAR), and regulate plant-pathogen interactions (Nehela and Killiny, 2020; Asija et al., 2022; Czerwoniec, 2022). It is worth noting that the specific mechanisms and roles of PAs in plant defense vary depending on plant species, pathogens, and environmental conditions. The most abundant PA in plants is biosynthesized from the essential polyamine L-ornithine (Killiny and Nehela, 2020).
       L-ornithine plays multiple roles in plant growth and development. For example, previous studies have shown that in rice (Oryza sativa), ornithine may be associated with nitrogen recycling (Liu et al., 2018), rice yield, quality and aroma (Lu et al., 2020), and water stress response (Yang et al., 2000). Furthermore, exogenous application of L-ornithine significantly enhanced drought tolerance in sugar beet (Beta vulgaris) (Hussein et al., 2019) and alleviated salt stress in onion (Allium Cepa) (Çavuşoǧlu and Çavuşoǧlu, 2021) and cashew (Anacardium occidentale) plants (da Rocha et al., 2012). The potential role of L-ornithine in abiotic stress defense may be due to its involvement in proline accumulation in treated plants. For example, genes related to proline metabolism, such as ornithine delta aminotransferase (delta-OAT) and proline dehydrogenase (ProDH1 and ProDH2) genes, have previously been reported to be involved in the defense of Nicotiana benthamiana and Arabidopsis thaliana against non-host Pseudomonas syringae strains (Senthil-Kumar and Mysore, 2012). On the other hand, fungal ornithine decarboxylase (ODC) is required for pathogen growth (Singh et al., 2020). Targeting ODC of Fusarium oxysporum f. sp. lycopersici via host-induced gene silencing (HIGS) significantly enhanced the resistance of tomato plants to Fusarium wilt (Singh et al., 2020). However, the potential role of exogenous ornithine application against biotic stresses such as phytopathogens has not been well studied. More importantly, the effects of ornithine on disease resistance and the associated biochemical and physiological phenomena remain largely unexplored.
       Understanding the complexity of S. sclerotiorum infection of legumes is important for the development of effective control strategies. In this study, we aimed to identify the potential role of the diamine L-ornithine as a key factor in enhancing the defense mechanisms and resistance of legume plants to Sclerotinia sclerotiorum infection. We hypothesize that, in addition to enhancing the defense responses of infected plants, L-ornithine also plays a key role in maintaining the redox status. We propose that the potential effects of L-ornithine are related to the regulation of enzymatic and non-enzymatic antioxidant defense mechanisms and interference with fungal pathogenicity/virulence factors and associated proteins. This dual functionality of L-ornithine makes it a promising candidate for a sustainable strategy to mitigate the impact of white mold and enhance the resistance of common legume crops to this potent fungal pathogen. The results of the present study may help in the development of innovative, environmentally friendly methods to control white mold and mitigate its impact on legume production.
       In this study, a susceptible commercial variety of common bean, Giza 3 (Phaseolus vulgaris L. cv. Giza 3), was used as experimental material. Healthy seeds were kindly provided by the Legume Research Department, Field Crops Research Institute (FCRI), Agricultural Research Center (ARC), Egypt. Five seeds were sown in plastic pots (inner diameter 35 cm, depth 50 cm) filled with S. sclerotiorum-infected soil under greenhouse conditions (25 ± 2 °C, relative humidity 75 ± 1%, 8 h light/16 h dark). At 7–10 days after sowing (DPS), the seedlings were thinned to leave only two seedlings with uniform growth and three fully expanded leaves in each pot. All potted plants were watered once every two weeks and fertilized monthly at the recommended rate for the given variety.
       To prepare a 500 mg/L concentration of L-ornithinediamine (also known as (+)-(S)-2,5-diaminopentanoic acid; Sigma-Aldrich, Darmstadt, Germany), 50 mg was first dissolved in 100 mL of sterile distilled water. The stock solution was then diluted and used in subsequent experiments. Briefly, six series of L-ornithine concentrations (12.5, 25, 50, 75, 100, and 125 mg/L) were tested in vitro. In addition, sterile distilled water was used as a negative control (Mock) and commercial fungicide “Rizolex-T” 50% wettable powder (toclofos-methyl 20% + thiram 30%; KZ-Kafr El Zayat Pesticides and Chemicals Company, Kafr El Zayat, Gharbia Governorate, Egypt) was used as a positive control. Commercial fungicide “Rizolex-T” was tested in vitro at five concentrations (2, 4, 6, 8 and 10 mg/L).
       Samples of common bean stems and pods showing typical symptoms of white mold (infestation rate: 10–30%) were collected from commercial farms. Although most of the infected plant materials were identified by species/variety (susceptible commercial variety Giza 3), others, especially those obtained from local markets, were of unknown species. The collected infected materials were first surface disinfected with 0.5% sodium hypochlorite solution for 3 minutes, then rinsed several times with sterile distilled water and wiped dry with sterile filter paper to remove excess water. The infected organs were then cut into small pieces from the middle tissue (between healthy and infected tissues), cultured on potato dextrose agar (PDA) medium and incubated at 25 ± 2 °C with a 12 h light/12 h dark cycle for 5 days to stimulate sclerotia formation. The mycelial tip method was also used to purify fungal isolates from mixed or contaminated cultures. The purified fungal isolate was first identified based on its cultural morphological characteristics and then confirmed to be S. sclerotiorum based on microscopic features. Finally, all purified isolates were tested for pathogenicity on the susceptible common bean cultivar Giza 3 to meet Koch’s postulates.
       In addition, the most invasive S. sclerotiorum isolate (isolate #3) was further confirmed based on internal transcribed spacer (ITS) sequencing as described by White et al., 1990; Baturo-Ciesniewska et al., 2017. Briefly, isolates were cultured in potato dextrose broth (PDB) and incubated at 25 ± 2 °C for 5–7 days. Fungal mycelium was then collected, filtered through cheesecloth, washed twice with sterile water, and dried with sterile filter paper. Genomic DNA was isolated using the Quick-DNA™ Fungal/Bacterial Miniprep Kit (Kuramae-Izioka, 1997; Atallah et al., 2022, 2024). The ITS rDNA region was then amplified using the specific primer pair ITS1/ITS4 (TCCGTAGGTGAACCTGCGG TCCTCCGCTTATTGATATGC; expected size: 540 bp) ( Baturo-Ciesniewska et al., 2017 ). The purified PCR products were submitted for sequencing (Beijing Aoke Dingsheng Biotechnology Co., Ltd.). The ITS rDNA sequences were sequenced bidirectionally using the Sanger sequencing method. The assembled query sequences were then compared with the latest data in GenBank and the National Center for Biotechnology Information (NCBI, http://www.ncbi.nlm.nih.gov/gene/) using BLASTn software. The query sequence was compared with 20 other S. sclerotiorum strains/isolates retrieved from the latest data in NCBI GenBank (Supplementary Table S1) using ClustalW in the Molecular Evolutionary Genetics Analysis Package (MEGA-11; version 11) (Kumar et al., 2024). Evolutionary analysis was performed using the maximum likelihood method and the general time-reversible nucleotide substitution model (Nei and Kumar, 2000). The tree with the highest log-likelihood is shown. The initial tree for the heuristic search is selected by choosing the tree with the higher log-likelihood between the neighbor-joining (NJ) tree (Kumar et al., 2024) and the maximum parsimony (MP) tree. The NJ tree was constructed using a pairwise distance matrix calculated using the general time-reversible model (Nei and Kumar, 2000).
       The antibacterial activity of L-ornithine and the bactericide “Rizolex-T” was determined in vitro by the agar diffusion method. Method: Take the appropriate amount of the stock solution of L-ornithine (500 mg/L) and mix it thoroughly with 10 ml of the PDA nutrient medium to prepare solutions with final concentrations of 12.5, 25, 50, 75, 100 and 125 mg/L, respectively. Five concentrations of the fungicide “Rizolex-T” (2, 4, 6, 8 and 10 mg/L) and sterile distilled water were used as a control. After the medium had solidified, a freshly prepared mycelial plug of Sclerotinia sclerotiorum culture, 4 mm in diameter, was transferred to the center of the Petri dish and cultured at 25±2°C until the mycelium covered the entire control Petri dish, after which the fungal growth was recorded. Calculate the percentage inhibition of radial growth of S. sclerotiorum using equation 1:
       The experiment was repeated twice, with six biological replicates for each control/experimental group and five pots (two plants per pot) for each biological replicate. Each biological replicate was analyzed twice (two technical replicates) to ensure the accuracy, reliability and reproducibility of the experimental results. In addition, probit regression analysis was used to calculate the half-maximal inhibitory concentration (IC50) and IC99 (Prentice, 1976).
       To evaluate the potential of L-ornithine under greenhouse conditions, two consecutive pot experiments were conducted. Briefly, pots were filled with sterilized clay-sand soil (3:1) and inoculated with a freshly prepared culture of S. sclerotiorum. First, the most invasive isolate of S. sclerotiorum (isolate #3) was cultured by cutting one sclerotium in half, placing it face down on a PDA and incubating at 25°C in constant darkness (24 h) for 4 days to stimulate mycelial growth. Four 5 mm diameter agar plugs were then taken from the leading edge and inoculated with 100 g of a sterile mixture of wheat and rice bran (1:1, v/v) and all flasks were incubated at 25 ± 2 °C under a 12 h light/12 h dark cycle for 5 days to stimulate sclerotia formation. The contents of all flasks were thoroughly mixed to ensure homogeneity before adding soil. Then, 100 g of the colonizing bran mixture was added to each pot to ensure a constant concentration of pathogens. The inoculated pots were watered to activate fungal growth and placed in greenhouse conditions for 7 days.
       Five seeds of the Giza 3 variety were then sown in each pot. For the pots treated with L-ornithine and the fungicide Rizolex-T, the sterilized seeds were first soaked for two hours in an aqueous solution of the two compounds with a final IC99 concentration of about 250 mg/L and 50 mg/L, respectively, and then air-dried for one hour before sowing. On the other hand, the seeds were soaked in sterile distilled water as a negative control. After 10 days, before the first watering, the seedlings were thinned out, leaving only two neat seedlings in each pot. Additionally, to ensure infection with S. sclerotiorum, bean plant stems at the same developmental stage (10 days) were cut at two different locations using a sterilized scalpel and approximately 0.5 g of the colonizing bran mixture was placed into each wound, followed by high humidity to stimulate infection and disease development in all inoculated plants. Control plants were similarly wounded and an equal amount (0.5 g) of sterile, uncolonized bran mixture was placed into the wound and maintained under high humidity to simulate the environment for disease development and ensure consistency between treatment groups.
       Treatment method: Bean seedlings were watered with 500 ml of an aqueous solution of L-ornithine (250 mg/l) or the fungicide Rizolex-T (50 mg/l) by irrigating the soil, then the treatment was repeated three times at an interval of 10 days. The placebo-treated controls were irrigated with 500 ml of sterile distilled water. All treatments were carried out under greenhouse conditions (25 ± 2°C, 75 ± 1% relative humidity, and a photoperiod of 8 h light/16 h dark). All pots were watered fortnightly and treated monthly with a balanced NPK fertilizer (20-20-20, with 3.6% sulfur and TE microelements; Zain Seeds, Egypt) at a concentration of 3–4 g/l by foliar spraying according to the recommendations for the specific variety and the manufacturer’s instructions. Unless otherwise stated, fully expanded mature leaves (2nd and 3rd leaves from top) were collected from each biological replicate at 72 h post-treatment (hpt), homogenized, pooled and stored at -80 °C for further analyses including, but not limited to, in situ histochemical localization of oxidative stress indicators, lipid peroxidation, enzymatic and non-enzymatic antioxidants and gene expression.
       White mold infection intensity was assessed weekly 21 days post inoculation (dpi) using a scale of 1–9 (Supplementary Table S2) based on the Petzoldt and Dickson scale (1996) modified by Teran et al. (2006). Briefly, the stems and branches of bean plants were examined starting at the point of inoculation to follow the progression of lesions along internodes and nodes. The distance of the lesion from the point of inoculation to the furthest point along the stem or branch was then measured and a score of 1–9 was assigned based on the location of the lesion, where (1) indicated no visible infection near the point of inoculation and (2–9) indicated a gradual increase in lesion size and progression along nodes/internodes (Supplementary Table S2). White mold infection intensity was then converted to percentage using formula 2:
       In addition, the area under the disease progression curve (AUDPC) was calculated using the formula (Shaner and Finney, 1977), which was recently adapted for white rot of common bean (Chauhan et al., 2020) using equation 3:
       Where Yi = disease severity at time ti, Yi+1 = disease severity at next time ti+1, ti = time of first measurement (in days), ti+1 = time of next measurement (in days), n = total number of time points or observation points. Bean plant growth parameters including plant height (cm), number of branches per plant, and number of leaves per plant were recorded weekly for 21 days in all biological replicates.
       In each biological replicate, leaf samples (second and third fully developed leaves from the top) were collected on day 45 after treatment (15 days after the last treatment). Each biological replicate consisted of five pots (two plants per pot). About 500 mg of the crushed tissue was used for the extraction of photosynthetic pigments (chlorophyll a, chlorophyll b and carotenoids) using 80% acetone at 4 °C in the dark. After 24 h, the samples were centrifuged and the supernatant was collected for the determination of chlorophyll a, chlorophyll b and carotenoid contents colorimetrically using a UV-160A spectrophotometer (Shimadzu Corporation, Japan) according to the method of (Lichtenthaler, 1987) by measuring the absorbance at three different wavelengths (A470, A646 and A663 nm). Finally, the content of photosynthetic pigments was calculated using the following formulas 4–6 described by Lichtenthaler (1987).
       At 72 h post-treatment (hpt), leaves (second and third fully developed leaves from the top) were collected from each biological replicate for in situ histochemical localization of hydrogen peroxide (H2O2) and superoxide anion (O2•−). Each biological replicate consisted of five pots (two plants per pot). Each biological replicate was analyzed in duplicate (two technical replicates) to ensure the accuracy, reliability and reproducibility of the method. H2O2 and O2•− were determined using 0.1% 3,3′-diaminobenzidine (DAB; Sigma-Aldrich, Darmstadt, Germany) or nitroblue tetrazolium (NBT; Sigma-Aldrich, Darmstadt, Germany), respectively, following the methods described by Romero-Puertas et al. (2004) and Adam et al. (1989) with minor modifications. For histochemical localization of H2O2 in situ, leaflets were vacuum infiltrated with 0.1% DAB in 10 mM Tris buffer (pH 7.8) and then incubated at room temperature in the light for 60 min. Leaflets were bleached in 0.15% (v/v) TCA in 4:1 (v/v) ethanol:chloroform (Al-Gomhoria Pharmaceuticals and Medical Supplies, Cairo, Egypt) and then exposed to light until they darkened. Similarly, valves were vacuum infiltrated with 10 mM potassium phosphate buffer (pH 7.8) containing 0.1 w/v % HBT for histochemical localization of O2•− in situ. The leaflets were incubated in the light at room temperature for 20 min, then bleached as above, and then illuminated until dark blue/violet spots appeared. The intensity of the resulting brown (as an H2O2 indicator) or blue-violet (as an O2•− indicator) color was assessed using the Fiji version of the image processing package ImageJ (http://fiji.sc; accessed 7 March 2024).
       Malondialdehyde (MDA; as a marker of lipid peroxidation) was determined according to the method of Du and Bramlage (1992) with slight modifications. Leaves from each biological replicate (second and third fully developed leaves from the top) were collected 72 h post-treatment (hpt). Each biological replicate included five pots (two plants per pot). Each biological replicate was analyzed in duplicate (two technical replicates) to ensure the accuracy, reliability and reproducibility of the method. Briefly, 0.5 g of ground leaf tissue was used for MDA extraction with 20% trichloroacetic acid (TCA; MilliporeSigma, Burlington, MA, USA) containing 0.01% butylated hydroxytoluene (BHT; Sigma-Aldrich, St. Louis, MO, USA). The MDA content in the supernatant was then determined colorimetrically by measuring the absorbance at 532 and 600 nm using a UV-160A spectrophotometer (Shimadzu Corporation, Japan) and then expressed as nmol g−1 FW.
       For assessment of non-enzymatic and enzymatic antioxidants, leaves (second and third fully developed leaves from the top) were collected from each biological replicate at 72 h post-treatment (hpt). Each biological replicate consisted of five pots (two plants per pot). Each biological sample was analyzed in duplicate (two technical samples). Two leaves were ground with liquid nitrogen and used directly for determination of enzymatic and non-enzymatic antioxidants, total amino acids, proline content, gene expression, and oxalate quantification.
       Total soluble phenolics were determined using Folin-Ciocalteu reagent (Sigma-Aldrich, St. Louis, MO, USA) with slight modifications of the method described by Kahkonen et al. (1999). Briefly, approximately 0.1 g of homogenized leaf tissue was extracted with 20 ml 80% methanol in the dark for 24 h and the supernatant was collected after centrifugation. 0.1 ml of the sample extract was mixed with 0.5 ml Folin-Ciocalteu reagent (10%), shaken for 30 s and left in the dark for 5 min. Then 0.5 ml of 20% sodium carbonate solution (Na2CO3; Al-Gomhoria Pharmaceuticals and Medical Supplies Company, Cairo, Egypt) was added to each tube, mixed thoroughly and incubated at room temperature in the dark for 1 h. After incubation, the absorbance of the reaction mixture was measured at 765 nm using a UV-160A spectrophotometer (Shimadzu Corporation, Japan). The concentration of total soluble phenols in the sample extracts was determined using a gallic acid calibration curve (Fisher Scientific, Hampton, NH, USA) and expressed as milligrams of gallic acid equivalent per gram of fresh weight (mg GAE g-1 fresh weight).
       Total soluble flavonoid content was determined according to the method of Djeridane et al. (2006) with slight modifications. Briefly, 0.3 ml of the above methanol extract was mixed with 0.3 ml of 5% aluminum chloride solution (AlCl3; Fisher Scientific, Hampton, NH, USA), vigorously stirred and then incubated at room temperature for 5 min, followed by the addition of 0.3 ml of 10% potassium acetate solution (Al-Gomhoria Pharmaceuticals and Medical Supplies, Cairo, Egypt), thoroughly mixed and incubated at room temperature for 30 min in the dark. After incubation, the absorbance of the reaction mixture was measured at 430 nm using a UV-160A spectrophotometer (Shimadzu Corporation, Japan). The concentration of total soluble flavonoids in sample extracts was determined using a rutin calibration curve (TCI America, Portland, OR, USA) and then expressed as milligrams of rutin equivalent per gram of fresh weight (mg RE g-1 fresh weight).
       The total free amino acid content of bean leaves was determined using a modified ninhydrin reagent (Thermo Scientific Chemicals, Waltham, MA, USA) based on the method proposed by Yokoyama and Hiramatsu (2003) and modified by Sun et al. (2006). Briefly, 0.1 g of ground tissue was extracted with pH 5.4 buffer, and 200 μL of the supernatant was reacted with 200 μL of ninhydrin (2%) and 200 μL of pyridine (10%; Spectrum Chemical, New Brunswick, NJ, USA), incubated in a boiling water bath for 30 min, then cooled and measured at 580 nm using a UV-160A spectrophotometer (Shimadzu Corporation, Japan). On the other hand, proline was determined by the Bates method (Bates et al., 1973). Proline was extracted with 3% sulfosalicylic acid (Thermo Scientific Chemicals, Waltham, MA, USA) and after centrifugation, 0.5 ml of the supernatant was mixed with 1 ml glacial acetic acid (Fisher Scientific, Hampton, NH, USA) and ninhydrin reagent, incubated at 90°C for 45 min, cooled and measured at 520 nm using the same spectrophotometer as above. Total free amino acids and proline in the leaf extracts were determined using glycine and proline calibration curves (Sigma-Aldrich, St Louis, MO, USA), respectively, and expressed as mg/g fresh weight.
       To determine the enzymatic activity of antioxidant enzymes, approximately 500 mg of homogenized tissue was extracted with 3 ml of 50 mM Tris buffer (pH 7.8) containing 1 mM EDTA-Na2 (Sigma-Aldrich, St. Louis, MO, USA) and 7.5% polyvinylpyrrolidone (PVP; Sigma-Aldrich, St. Louis, MO, USA), centrifuged at 10,000 × g for 20 min under refrigeration (4 °C), and the supernatant (crude enzyme extract) was collected (El-Nagar et al., 2023; Osman et al., 2023). Catalase (CAT) was then reacted with 2 ml of 0.1 M sodium phosphate buffer (pH 6.5; Sigma-Aldrich, St. Louis, MO, USA) and 100 μl of 269 mM H2O2 solution to determine its enzymatic activity according to the method of Aebi (1984) with slight modifications (El-Nagar et al., 2023; Osman et al., 2023). Guaiacol-dependent peroxidase (POX) enzymatic activity was determined using the method of Harrach et al. (2009). (2008) with minor modifications (El-Nagar et al., 2023; Osman et al., 2023) and the enzymatic activity of polyphenol oxidase (PPO) was determined after reaction with 2.2 ml of 100 mM sodium phosphate buffer (pH 6.0), 100 μl of guaiacol (TCI chemicals, Portland, OR, USA) and 100 μl of 12 mM H2O2. The method was slightly modified from (El-Nagar et al., 2023; Osman et al., 2023). The assay was performed after reaction with 3 ml of catechol solution (Thermo Scientific Chemicals, Waltham, MA, USA) (0.01 M) freshly prepared in 0.1 M phosphate buffer (pH 6.0). CAT activity was measured by monitoring the decomposition of H2O2 at 240 nm (A240), POX activity was measured by monitoring the increase in absorbance at 436 nm (A436), and PPO activity was measured by recording absorbance fluctuations at 495 nm (A495) every 30 s for 3 min using a UV-160A spectrophotometer (Shimadzu, Japan).
       Real-time RT-PCR was used to detect the transcript levels of three antioxidant-related genes, including peroxisomal catalase (PvCAT1; GenBank Accession No. KF033307.1), superoxide dismutase (PvSOD; GenBank Accession No. XM_068639556.1), and glutathione reductase (PvGR; GenBank Accession No. KY195009.1), in bean leaves (the second and third fully developed leaves from the top) 72 h after the last treatment. Briefly, RNA was isolated using Simply P Total RNA Extraction Kit (Cat. No. BSC52S1; BioFlux, Biori Technology, China) according to the manufacturer’s protocol. Then, cDNA was synthesized using TOP script™ cDNA Synthesis Kit according to the manufacturer’s instructions. The primer sequences of the above three genes are listed in Supplementary Table S3. PvActin-3 (GenBank accession number: XM_068616709.1) was used as the housekeeping gene and the relative gene expression was calculated using the 2-ΔΔCT method (Livak and Schmittgen, 2001). Actin stability under biotic stress (incompatible interaction between common legumes and the anthracnose fungus Colletotrichum lindemuthianum) and abiotic stress (drought, salinity, low temperature) was demonstrated (Borges et al., 2012).
       We initially performed a genome-wide in silico analysis of oxaloacetate acetylhydrolase (OAH) proteins in S. sclerotiorum using the protein-protein BLAST tool (BLASTp 2.15.0+) (Altschul et al., 1997, 2005). Briefly, we used OAH from Aspergillus fijiensis CBS 313.89 (AfOAH; taxide: 1191702; GenBank accession number XP_040799428.1; 342 amino acids) and Penicillium lagena (PlOAH; taxide: 94218; GenBank accession number XP_056833920.1; 316 amino acids) as query sequences to map the homologous protein in S. sclerotiorum (taxide: 5180). BLASTp was performed against the most recently available S. sclerotiorum genome data in GenBank on the National Center for Biotechnology Information (NCBI) website, http://www.ncbi.nlm.nih.gov/gene/.
       In addition, the predicted OAH gene from S. sclerotiorum (SsOAH) and the evolutionary analysis and phylogenetic tree of AfOAH from A. fijiensis CBS 313.89 and PlOAH from P. lagena were inferred using the maximum likelihood method in MEGA11 (Tamura et al., 2021) and the JTT matrix-based model (Jones et al., 1992). The phylogenetic tree was combined with the multiple alignment analysis of protein sequences of all predicted OAH genes (SsOAH) from S. sclerotiorum and the query sequence using the Constraint-Based Alignment Tool (COBALT; https://www.ncbi.nlm.nih.gov/tools/cobalt/re_cobalt.cgi) (Papadopoulos and Agarwala, 2007). In addition, the best matching amino acid sequences of SsOAH from S. sclerotiorum were aligned with the query sequences (AfOAH and PlOAH) (Larkin et al., 2007) using ClustalW (http://www.genome.jp/tools-bin/clustalw), and conserved regions in the alignment were visualized using the ESPript tool (version 3.0; https://espript.ibcp.fr/ESPript/ESPript/index.php).
       Furthermore, the predicted functional representative domains and conserved sites of S. sclerotiorum SsOAH were interactively classified into different families using the InterPro tool (https://www.ebi.ac.uk/interpro/) (Blum et al., 2021). Finally, three-dimensional (3D) structure modeling of the predicted S. sclerotiorum SsOAH was performed using the Protein Homology/Analogy Recognition Engine (Phyre2 server version 2.0; http://www.sbg.bio.ic.ac.uk/~phyre2/html/page.cgi?id=index) (Kelley et al., 2015) and validated using the SWISS-MODEL server (https://swissmodel.expasy.org/) (Biasini et al., 2014). The predicted three-dimensional structures (PDB format) were interactively visualized using the UCSF-Chimera package (version 1.15; https://www.cgl.ucsf.edu/chimera/ ) (Pettersen et al., 2004).
       Quantitative real-time fluorescence PCR was used to determine the transcriptional level of oxaloacetate acetylhydrolase (SsOAH; GenBank accession number: XM_001590428.1) in the mycelia of Sclerotinia sclerotiorum. Briefly, S. sclerotiorum was inoculated into a flask containing PDB and placed in a shaking incubator (model: I2400, New Brunswick Scientific Co., Edison, NJ, USA) at 25 ± 2 °C for 24 h at 150 rpm and in constant darkness (24 h) to stimulate mycelial growth. Thereafter, the cells were treated with L-ornithine and the fungicide Rizolex-T at final IC50 concentrations (approximately 40 and 3.2 mg/L, respectively) and then cultured for another 24 h under the same conditions. After incubation, the cultures were centrifuged at 2500 rpm for 5 min and the supernatant (fungal mycelium) was collected for gene expression analysis. Similarly, fungal mycelium was collected at 0, 24, 48, 72, 96, and 120 h post-infection from infected plants that had formed white mold and cottony mycelium on the surface of infected tissues. RNA was extracted from the fungal mycelium and then cDNA was synthesized as described above. The primer sequences for SsOAH are listed in Supplementary Table S3. SsActin (GenBank accession number: XM_001589919.1) was used as the housekeeping gene, and relative gene expression was calculated using the 2-ΔΔCT method (Livak and Schmittgen, 2001).
       Oxalic acid was determined in potato dextrose broth (PDB) and plant samples containing the fungal pathogen Sclerotinia sclerotiorum according to the method of Xu and Zhang (2000) with slight modifications. Briefly, S. sclerotiorum isolates were inoculated into flasks containing PDB and then cultured in a shaking incubator (model I2400, New Brunswick Scientific Co., Edison, NJ, USA) at 150 rpm at 25 ± 2 °C for 3–5 days in constant darkness (24 h) to stimulate mycelial growth. After incubation, the fungal culture was first filtered through Whatman #1 filter paper and then centrifuged at 2500 rpm for 5 min to remove residual mycelium. The supernatant was collected and stored at 4°C for further quantitative determination of oxalate. For the preparation of plant samples, approximately 0.1 g of plant tissue fragments were extracted three times with distilled water (2 ml each time). The samples were then centrifuged at 2500 rpm for 5 min, the supernatant was dry filtered through Whatman No. 1 filter paper and collected for further analysis.
       For quantitative analysis of oxalic acid, the reaction mixture was prepared in a glass stoppered tube in the following order: 0.2 ml of sample (or PDB culture filtrate or oxalic acid standard solution), 0.11 ml of bromophenol blue (BPB, 1 mM; Fisher Chemical, Pittsburgh, PA, USA), 0.198 ml of 1 M sulfuric acid (H2SO4; Al-Gomhoria Pharmaceuticals and Medical Supplies, Cairo, Egypt) and 0.176 ml of 100 mM potassium dichromate (K2Cr2O7; TCI chemicals, Portland, OR, USA), and then the solution was diluted to 4.8 ml with distilled water, vigorously mixed and immediately placed in a 60 °C water bath. After 10 min, the reaction was stopped by adding 0.5 ml of sodium hydroxide solution (NaOH; 0.75 M). The absorbance (A600) of the reaction mixture was measured at 600 nm using a UV-160 spectrophotometer (Shimadzu Corporation, Japan). PDB and distilled water were used as controls for the quantification of culture filtrates and plant samples, respectively. The oxalic acid concentrations in the culture filtrates, expressed as micrograms of oxalic acid per milliliter of PDB medium (μg.mL−1), and in the leaf extracts, expressed as micrograms of oxalic acid per gram of fresh weight (μg.g−1 FW), were determined using an oxalic acid calibration curve (Thermo Fisher Scientific Chemicals, Waltham, MA, USA).
       Throughout the study, all experiments were designed in a completely randomized design (CRD) with six biological replicates per treatment and five pots per biological replicate (two plants per pot) unless otherwise stated. Biological replicates were analyzed in duplicate (two technical replicates). Technical replicates were used to check the reproducibility of the same experiment but were not used in the statistical analysis to avoid spurious replicates. Data were statistically analyzed using analysis of variance (ANOVA) followed by Tukey-Kramer honestly significant difference (HSD) test (p ≤ 0.05). For in vitro experiments, IC50 and IC99 values ​​were calculated using the probit model and 95% confidence intervals were calculated.
       A total of four isolates were collected from different soybean fields in El Ghabiya Governorate, Egypt. On PDA medium, all isolates produced creamy white mycelium that quickly turned cottony white (Figure 1A) and then beige or brown at the sclerotium stage. Sclerotia are usually dense, black, spherical or irregular in shape, 5.2 to 7.7 mm long and 3.4 to 5.3 mm in diameter (Figure 1B). Although four isolates developed a marginal pattern of sclerotia at the edge of the culture medium after 10–12 days of incubation at 25 ± 2 °C (Fig. 1A), the number of sclerotia per plate was significantly different among them (P < 0.001), with isolate 3 having the highest number of sclerotia (32.33 ± 1.53 sclerotia per plate; Fig. 1C). Similarly, isolate #3 produced more oxalic acid in PDB than other isolates (3.33 ± 0.49 μg.mL−1; Fig. 1D). Isolate #3 showed typical morphological and microscopic characteristics of the phytopathogenic fungus Sclerotinia sclerotiorum. For example, on PDA, colonies of isolate #3 grew rapidly, were creamy white (Figure 1A), reverse beige or light salmon yellow-brown, and required 6-7 days at 25 ± 2°C to completely cover the surface of a 9 cm diameter plate. Based on the above morphological and microscopic characteristics, isolate #3 was identified as Sclerotinia sclerotiorum.
       Figure 1. Characteristics and pathogenicity of S. sclerotiorum isolates from common legume crops. (A) Mycelial growth of four S. sclerotiorum isolates on PDA medium, (B) sclerotia of four S. sclerotiorum isolates, (C) number of sclerotia (per plate), (D) oxalic acid secretion on PDB medium (μg.mL−1), and (E) disease severity (%) of four S. sclerotiorum isolates on susceptible commercial legume cultivar Giza 3 under greenhouse conditions. Values ​​represent the mean ± SD of five biological replicates (n = 5). Different letters indicate statistically significant differences between treatments (p < 0.05). (F–H) Typical white mold symptoms appeared on aboveground stems and siliques, respectively, 10 days post inoculation with isolate #3 (dpi). (I) Evolutionary analysis of the internal transcribed spacer (ITS) region of S. sclerotiorum isolate #3 was performed using the maximum likelihood method and compared with 20 reference isolates/strains obtained from the National Center for Biotechnology Information (NCBI) database (https://www.ncbi.nlm.nih.gov/). The numbers above the clustering lines indicate the region coverage (%), and the numbers below the clustering lines indicate the branch length.
       Furthermore, to confirm the pathogenicity, four obtained S. sclerotiorum isolates were used to inoculate the susceptible commercial bean cultivar Giza 3 under greenhouse conditions, which is consistent with Koch’s postulates (Fig. 1E). Although all the obtained fungal isolates were pathogenic and could infect the green bean (cv. Giza 3), causing typical white mold symptoms on all above-ground parts (Fig. 1F), especially on the stems (Fig. 1G) and pods (Fig. 1H) at 10 days post inoculation (dpi), isolate 3 was the most aggressive isolate in two independent experiments. Isolate 3 had the highest disease severity (%) on bean plants (24.0 ± 4.0, 58.0 ± 2.0, and 76.7 ± 3.1 at 7, 14, and 21 days post-infection, respectively; Figure 1F).
       The identification of the most invasive S. sclerotiorum isolate #3 was further confirmed based on internal transcribed spacer (ITS) sequencing (Fig. 1I). Phylogenetic analysis between isolate #3 and 20 reference isolates/strains showed high similarity (>99%) between them. It is worth noting that the S. sclerotiorum isolate #3 (533 bp) has a high similarity to the American S. sclerotiorum isolate LPM36 isolated from dry pea seeds (GenBank accession number MK896659.1; 540 bp) and the Chinese S. sclerotiorum isolate YKY211 (GenBank accession number OR206374.1; 548 bp), which causes violet (Matthiola incana) stem rot, all of which are grouped separately at the top of the dendrogram (Figure 1I). The new sequence has been deposited in the NCBI database and named “Sclerotinia sclerotiorum – isolate YN-25” (GenBank accession number PV202792). It can be seen that isolate 3 is the most invasive isolate; therefore, this isolate was chosen for study in all subsequent experiments.
       The antibacterial activity of the diamine L-ornithine (Sigma-Aldrich, Darmstadt, Germany) at different concentrations (12.5, 25, 50, 75, 100 and 125 mg/L) against S. sclerotiorum isolate 3 was investigated in vitro. It is noteworthy that L-ornithine exerted an antibacterial effect and gradually inhibited the radial growth of S. sclerotiorum hyphae in a dose-dependent manner (Figure 2A, B). At the highest concentration tested (125 mg/L), L-ornithine demonstrated the highest mycelial growth inhibition rate (99.62 ± 0.27%; Figure 2B), which was equivalent to the commercial fungicide Rizolex-T (inhibition rate 99.45 ± 0.39%; Figure 2C) at the highest concentration tested (10 mg/L), indicating similar efficacy.
       Figure 2. In vitro antibacterial activity of L-ornithine against Sclerotinia sclerotiorum. (A) Comparison of the antibacterial activity of different concentrations of L-ornithine against S. sclerotiorum with the commercial fungicide Rizolex-T (10 mg/L). (B, C) Inhibition rate (%) of S. sclerotiorum mycelial growth after treatment with different concentrations of L-ornithine (12.5, 25, 50, 75, 100 and 125 mg/L) or Rizolex-T (2, 4, 6, 8 and 10 mg/L), respectively. Values ​​represent the mean ± SD of five biological replicates (n = 5). Different letters denote statistical differences between treatments (p < 0.05). (D, E) Probit model regression analysis of L-ornithine and commercial fungicide Rizolex-T, respectively. The probit model regression line is shown as a solid blue line, and the confidence interval (95%) is shown as a dashed red line.
       In addition, probit regression analysis was performed and the corresponding plots are shown in Table 1 and Figures 2D,E. Briefly, the acceptable slope value (y = 2.92x − 4.67) and associated significant statistics (Cox & Snell R2 = 0.3709, Nagelkerke R2 = 0.4998 and p < 0.0001; Figure 2D) of L-ornithine indicated an enhanced antifungal activity against S. sclerotiorum compared to the commercial fungicide Rizolex-T (y = 1.96x − 0.99, Cox & Snell R2 = 0.1242, Nagelkerke R2 = 0.1708 and p < 0.0001) (Table 1).
       Table 1. Values ​​of half-maximum inhibitory concentration (IC50) and IC99 (mg/l) of L-ornithine and commercial fungicide “Rizolex-T” against S. sclerotiorum.
       Overall, L-ornithine (250 mg/L) significantly reduced the development and severity of white mold on treated common bean plants compared to untreated S. sclerotiorum-infected plants (control; Figure 3A). Briefly, although the disease severity of untreated infected control plants gradually increased (52.67 ± 1.53, 83.21 ± 2.61, and 92.33 ± 3.06%), L-ornithine significantly reduced the disease severity (%) throughout the experiment (8.97 ± 0.15, 18.00 ± 1.00, and 26.36 ± 3.07) at 7, 14, and 21 days post-treatment (dpt), respectively (Figure 3A). Similarly, when S. sclerotiorum-infected bean plants were treated with 250 mg/L L-ornithine, the area under the disease progression curve (AUDPC) decreased from 1274.33 ± 33.13 in the untreated control to 281.03 ± 7.95, which was slightly lower than that of the positive control 50 mg/L Rizolex-T fungicide (183.61 ± 7.71; Fig. 3B). The same trend was observed in the second experiment.
       Fig. 3. Effect of exogenous application of L-ornithine on the development of white rot of common bean caused by Sclerotinia sclerotiorum under greenhouse conditions. (A) Disease progression curve of white mold of common bean after treatment with 250 mg/L L-ornithine. (B) Area under the disease progression curve (AUDPC) of white mold of common bean after treatment with L-ornithine. Values ​​represent the mean ± SD of five biological replicates (n = 5). Different letters denote statistically significant differences between treatments (p < 0.05).
       Exogenous application of 250 mg/L L-ornithine gradually increased plant height (Fig. 4A), number of branches per plant (Fig. 4B), and number of leaves per plant (Fig. 4C) after 42 days. While the commercial fungicide Rizolex-T (50 mg/L) had the greatest effect on all nutritional parameters studied, exogenous application of 250 mg/L L-ornithine had the second greatest effect compared to untreated controls (Figs. 4A–C). On the other hand, L-ornithine treatment had no significant effect on the content of photosynthetic pigments chlorophyll a (Fig. 4D) and chlorophyll b (Fig. 4E), but slightly increased the total carotenoid content (0.56 ± 0.03 mg/g fr wt) compared to the negative control (0.44 ± 0.02 mg/g fr wt) and positive control (0.46 ± 0.02 mg/g fr wt; Fig. 4F). Overall, these results indicate that L-ornithine is not phytotoxic to treated legumes and may even stimulate their growth.
       Fig. 4. Effect of exogenous L-ornithine application on growth characteristics and photosynthetic pigments of bean leaves infected with Sclerotinia sclerotiorum under greenhouse conditions. (A) Plant height (cm), (B) Number of branches per plant, (C) Number of leaves per plant, (D) Chlorophyll a content (mg g-1 fr wt), (E) Chlorophyll b content (mg g-1 fr wt), (F) Total carotenoid content (mg g-1 fr wt). Values ​​are the mean ± SD of five biological replicates (n = 5). Different letters indicate statistically significant differences between treatments (p < 0.05).
       In situ histochemical localization of reactive oxygen species (ROS; expressed as hydrogen peroxide [H2O2]) and free radicals (expressed as superoxide anions [O2•−]) revealed that exogenous application of L-ornithine (250 mg/L) significantly reduced the accumulation of H2O2 (96.05 ± 5.33 nmol.g−1 FW; Fig. 5A) and O2•− (32.69 ± 8.56 nmol.g−1 FW; Fig. 5B) compared to the accumulation of both untreated infected plants (173.31 ± 12.06 and 149.35 ± 7.94 nmol.g−1 FW, respectively) and plants treated with 50 mg/L of the commercial fungicide Rizolex-T (170.12 ± 9.50 and 157.00 ± 7.81 nmol.g−1 fr wt, respectively) at 72 h. High levels of H2O2 and O2•− accumulated under hpt (Fig. 5A, B). Similarly, TCA-based malondialdehyde (MDA) assay showed that S. sclerotiorum-infected bean plants accumulated higher levels of MDA (113.48 ± 10.02 nmol.g fr wt) in their leaves (Fig. 5C). However, exogenous application of L-ornithine significantly reduced lipid peroxidation as evidenced by the decrease in MDA content in treated plants (33.08 ± 4.00 nmol.g fr wt).
       Fig. 5. Effect of exogenous L-ornithine application on major markers of oxidative stress and non-enzymatic antioxidant defense mechanisms in bean leaves infected with S. sclerotiorum at 72 h post-infection under greenhouse conditions. (A) Hydrogen peroxide (H2O2; nmol g−1 FW) at 72 hpt, (B) superoxide anion (O2•−; nmol g−1 FW) at 72 hpt, (C) malondialdehyde (MDA; nmol g−1 FW) at 72 hpt, (D) total soluble phenols (mg GAE g−1 FW) at 72 hpt, (E) total soluble flavonoids (mg RE g−1 FW) at 72 hpt, (F) total free amino acids (mg g−1 FW) at 72 hpt, and (G) proline content (mg g−1 FW) at 72 hpt. Values ​​represent the mean ± standard deviation (mean ± SD) of 5 biological replicates (n = 5). Different letters indicate statistically significant differences between treatments (p < 0.05).