Manual on Biotic Stress Resistance Evaluation First Edition (2025) INTERNATIONAL RICE RESEARCH INSTITUTE 2025 IRRI is the world’s premier research organization dedicated to reducing poverty and hunger through rice science, improving the health and welfare of rice farmers and consumers, and protecting the rice- growing environment for future generations. Headquartered in the Philippines and with offices in 17 countries, IRRI is a global, independent, nonprofit research and training institute supported by public and private donors. IRRI is a member of CGIAR, a global initiative uniting international agricultural research organizations to address interconnected challenges in food, land, and water systems. IRRI's approach to research and development emphasizes collaboration. It forges alliances with advanced research institutions, collaborates with governments and national agricultural research and extension systems, and partners with development organizations. https://www.irri.org/ https://www.facebook.com/IRRI.Official About IRRI Authors: Suggested Citation: Yanoria, M. J., Atienza-Grande, G., Jonson, G., Almazan, M. L., Pacia, J., Castilla, N., Vera Cruz, C., & Schepler-Luu, V. (2025). Manual on Biotic Stress Resistance Evaluation (First Edition). International Rice Research Institute. Los Baños, Philippines Contact Information: Physical Address: Pili Drive, Los Baños, Laguna 4031, Philippines Mailing Address: DAPO Box 7777, Metro Manila 1301, Philippines Phone: +63 (2) 8580 5600, +63 (2) 8845 0563 Email: irri-education@cgiar.org Web: education.irri.org This manual is an output of the Hands-on Training on Biotic Stress Resistance Evaluation, implemented by IRRI Education and the group of Plant Pathology and Host Plant Resistance, Entomology, and Biotic Stress Resistance Evaluation Center (BSREC). ©International Rice Research Institute All Rights Reserved 2025 Copyright: First Edition (2025)Edition Number: Acknowledgement: IRRI Education Mary Jeanie Telebanco-Yanoria Genelou Atienza-Grande Gilda Jonson Maria Liberty Almazan Carmencita Bernal Nurmi Pangesti Jocelyn Pacia Nancy Castilla Raphael Carvajal Kendrix Ortega Rafael Luis Natulan Ma Michelle Valenzuela Casiana Vera Cruz Van Schepler-Luu DOI: 10.5281/zenodo.15787885 https://doi.org/10.5281/zenodo.15787885 Table of Contents ......................................................................................1 .....................................................74 ...................................................................86 .............................................94 Overview Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases .........................................................................................................................................................................................i Bacterial blight Bacterial leaf streak Rice Blast Sheath Blight Brown Spot False Smut Module 2: Insect Collection and Rearing in the Greenhouse, Infestation Techniques and Damage Assessments Brown Planthoppers (BPH) and Green Leafhoppers (GLH) Yellow Stemborer Module 3: Virus Propagation, Inoculation and Evaluation Tungro Virus Module 4: Evaluation of resistance to the rice root-knot nematode Rice root-knot nematode 1.1 1.2 1.3 1.4 1.5 1.6 2.1 2.2 3.1 4.1 .........................................................................................................................................................1 ..............................................................................................................................................16 .....................................................................................................................................................................23 ............................................................................................................................................................38 .................................................................................................................................................................49 ..................................................................................................................................................................64 ...............................................................................................................74 ...................................................................................................................................................79 .............................................................................................................................................................86 ...............................................................................................................................94 Biotic stresses due to pests and diseases are dynamic, and their occurrence is influenced by several factors, such as host genotype, cropping practices, and extreme weather events brought about by climate change. Major diseases caused by pathogens, and nematodes and insect pests, such as brown planthoppers, green leafhoppers, and stemborers, reduce rice yields in different ecosystems, if these are not properly assessed and managed. The use of host plant resistance is generally considered as the most economical, practical, and environment-friendly strategy for pest management. It enables farmers to reduce yield losses and improve productivity and profitability without reliance on pesticides, which becomes increasingly important in the face of climate change. However, robust screening protocols are often lacking or not standardized. Hence, capacity development on the protocols used at IRRI for the evaluation of resistance to biotic stresses will facilitate the identification and selection of resistant genotypes. A standard protocol will facilitate collaboration among different institutes. Acquiring skills in evaluating rice genotypes to biotic stresses is crucial for developing pest management strategies. Overview i Bacterial blight, caused by Xanthomonas oryzae pv. oryzae (Xoo), is a serious rice disease that leads to wilting of seedlings and yellowing and drying of leaves. It thrives in areas with weeds and infected plant stubble, and is common in both tropical and temperate regions, especially in irrigated and rainfed lowland areas. The disease is favored by temperatures between 25−34°C and high humidity levels above 70%. It spreads easily during strong winds and continuous heavy rains through bacterial ooze from infected lesions. Susceptible rice varieties under high nitrogen fertilization are particularly vulnerable, and in such conditions, yield losses can reach up to 70%. Early infection leads to severe yield reduction, while infection at the booting stage does not impact yield but results in poor grain quality and more broken kernels. 1 Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases MODULE 1: 1.1 Bacterial Blight (BB) Materials I. Sampling strategies for disease collection in the field Label tags Shipping tags or pot labels Bamboo sticks (1 meter long) Abaca twine, string or wire Sterile pair of scissors Latex gloves Glassine bags or net bags or coin envelope Plastic bags Packaging or masking tape Ice chest with (fresh) ice Marking pens and/or pencils Plot layout Clipboard Authors: Genelou Atienza-Grande, Casiana M. Vera Cruz, Van Schepler-Luu Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases Procedure 1. Identify the field plot to be assessed. 2.Define the effective plot size, varieties of rice grown, and age of the plants. Perform a quick survey interview with the farmer to obtain the following information: (a) historical occurrences of diseases and varieties grown; (b) control tactics applied to manage the diseases; (c) source of irrigation; and (d) crop establishment. Obtain permission to do diseased sample collection from the field owners. 3.Check for the target disease symptoms (bacterial leaf blight symptoms). a.On seedlings, check for yellowing of leaves and wilting (also called kresek). Infected leaves turn grayish-green and roll up. As the disease progresses, the leaves turn yellow to straw-colored and wilt, leading whole seedlings to dry up and die. To distinguish kresek symptoms from stem borer damage, squeeze the lower end of infected seedlings between the fingers. Kresek symptoms should show a yellowish bacterial ooze coming out of the cut ends. Unlike plants infested with stem borer, rice plants with kresek are not easily pulled out from the soil. b.On older plants, check for lesions. Lesions usually develop as water-soaked to yellow-orange stripes on leaf blades or leaf tips or on mechanically injured parts of leaves. The lesions have a wavy margin and progress toward the leaf base. On young lesions, bacterial ooze resembling a milky dew drop can be observed early in the morning. The bacterial ooze later on dries up and becomes small yellowish beads underneath the leaf. Old lesions turn yellow to grayish-white with black dots due to the growth of various saprophytic fungi. On severely infected leaves, lesions may extend to the leaf sheath. Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases Figure 1. Yellowing and wilting (Kresek) symptoms in seedlings infected by bacterial blight 2 4. To quickly check for bacterial blight signs: Cut a young lesion across the leaf and place it in a clear glass container with clear water. Wait for a few minutes. Observe for thick or turbid liquid coming from the cut end of the leaf. Figure 2. Lesions on older rice plants caused by bacterial blight Photo courtesy of Genelou Grande and Eula Oreiro, PP&HPR-IRRI Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases Figure 3. Bacterial ooze test for Xoo Photo illustration courtesy of Dr. Nancy Castilla and Genelou Grande, IRRI PP&HPR-IRRI 3 5. To determine the size where the survey will be conducted, use stratified sampling techniques and hierarchical scheme within the selected hotspot target areas. This includes four levels of sampling, to divide the target sites into logical categories (refer to Figure 4). 1. Fill transparent glass with water. 2. Cut the leaf at the advancing part of young lesion. 3. Dip cut lesion in water 4. Check if the water becomes turbid. 4 Figure 4. Diagrammatic map illustrating the concept of a four level scheme of sampling. Figure 5. W sampling pattern for sampling hills (represented by circles) in a farmer’s field.(A) Five hills are sampled in a field with an area of less than one hectare and (B) eight hills are sampled in a field that is one hectare or larger. Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases a.Level 1 (Field): From an identified symptomatic field, determine the sampling units using the W pattern (refer to Figure 4). Sampling unit is defined as a hill. Five hills are sampled from fields with an area of less than one hectare and eight hills are sampled from larger fields (refer to Figure 5). b.Level 2 (Village): In cases where infections are spread in neighboring fields, identify 3-4 nearby fields that are 1 km apart from each other. Prioritize the diversity of rice varieties grown and select 3-4 different rice varieties. c.Level 3 (District): Search for other villages nearby that have infected fields. Follow the village collection scheme. d.Level 4 (Division): Search for 1-3 districts nearby that have infected fields. Follow the district collection scheme. 5 Note If possible, do not sample hills near the border of a field (approximately within the two- meter border) to avoid bias caused by interplot interference. This occurs when the pathogen or disease intensity is affected by adjacent plots or external factors. Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases From each sampled hill, collect five (5) leaves with young lesions observed as water-soaked to yellow-orange stripes on leaf blades or leaf tips. Cut the infected leaf at least 1‒2 inches below the advancing lesion using a pair of clean scissors. Make sure you are wearing gloves to prevent contamination from sweat or other bacteria.factors. 6. 7.Fold the leaves twice and place them neatly in a dry, properly labeled glassine bag or coin envelop, with the proper sample tag name (code or passport data). Take care not to wet the glassine bags or coin envelopes to avoid cross-contamination between samples. Figure 6. Example of sample tag for identification of collected diseased specimens from the field. Place the coin envelops inside a plastic bag and seal with tape. Immediately place the plastic bags (with coin envelopes) inside an ice chest filled with ice while in the field. Give an ID per sample and fill in the passport data table (Table 1). In the lab, store the samples at -20 °C. 8. 9. 10. 11. Sample ID Example: 2024_IRRI_BB_F1P1 Disease Example: Bacterial blight Name of sample collector Institute Date of leaf sample collection (dd-month-yyyy) Date of panicle sample collection (dd-month-yyyy) Longitude (preferably in decimal degrees) Latitude (preferably in decimal degrees) Division District Upazilla Union Village Variety Estimated field area (square meters) Crop development stage during sample collection Estimated leaf blast severity in the collected field (low, moderate, severe) Estimated yield loss in the collected field (in percentage) Estimated neck blast incidence in the collected field (low, moderate, severe) Name of farmer (if available) Remarks Table 1. Passport data sheet of the collected samples. 6 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases Materials Infected leaf samples A sterile pair of scissors Wire-loop or sterile wooden sticks Ceramic spot plate or mortar and pestle or 1.5 ml eppy tubes Alcohol lamp/Bunsen burner 70% ethanol Sterile distilled water Wakimoto’s agar Latex gloves Dipping glass Beaker Sterile paper towel Growth media preparation for Xoo Procedure 1.Disinfect the work area with 70% ethanol. 2.Turn on the Bunsen burner or light up the alcohol lamp. 3.Select an area of the leaf where the lesion is advancing. 4.Cut this area into small pieces, about 25mm , onto a ceramic spot plate with sterile buffer or distilled water. 5.Allow the bacteria to ooze out of the lesion and into the suspending medium. 6.Streak a subsample onto a pre-poured solid medium following the streak for an isolation pattern. Do this in duplicates per leaf sample. 7. Incubate the plates at 28°C and observe for growth of colonies exhibiting a yellow, mucoidal morphology. 8.Pick up a single colony and re-streak onto the plated medium. 9.Check for contamination. 10.Transfer a single colony onto a slant. II. Isolation and purification of Xoo from infected leaves Isolation of Xoo from leaves Cut a section from an area where the lesion is still advancing. Disinfect the cut leaf section by dipping it in 70% ethanol and rinse it in sterile distilled water. Cut the leaf sample into a smaller, 1-mm2 section and place it on a ceramic spot plate with sterile distilled water. a. b. c. 7 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases Xoo requires a growth medium containing sucrose (Noda and Kaku 1999). The most used media are peptone sucrose agar (Ou 1985), Suwa’s agar (sodium glutamate 2.0 g L‒1, MgCl2•6H2O 1.0 g L‒1, KH2PO4 0.1 g L‒1, peptone 10.0 g L‒1, sucrose 5.0 g L‒1, Fe-EDTA 1.0 mL L‒1) (Suwa 1962), and modified Wakimoto’s medium (sucrose 20.0 g L‒1, peptone 5.0 g L‒1, Ca(NO3)2•4H2O 0.5 g L‒1, FeSO4 0.05 g L‒1, agar 17 g L‒1) (Karganilla et al. 1973).This laboratory recommends modified Wakimoto’s medium to isolate, grow, and store working stocks of the bacterium. Let stand to allow the bacteria to ooze out into the suspending medium. Dip a sterile stick or loop into the suspension and streak onto pre-poured agar plates. Check the plates after 3‒5 days of incubation at 28°C. Pick three to four colonies from the plate. Streak into agar slants to obtain pure cultures of the isolate. d. e. f. g. h Figure 7. Step-by-step procedure on the isolation of Xoo from infected leaf tissues. Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 8 III. Maintenance of a viable and virulent bacterial culture collection Methods of culture preservation Preservation of Xoo and Xoc are done by cold storage in the appropriate medium: skim milk (skim milk (nonfat milk) 1 g L‒1, sodium glutamate 0.15 g L‒1, in distilled water; sterilize by autoclaving at 15 psi for 10 min) or glycerol (30%). For long-term storage of cultures, lyophilization of skimmed milk stocks is highly recommended. A. Periodic subculture Maintain cultures in slants or plates at 4°C for temporary storage for three (3) months. Temporary Storage Skimmed milk (nonfat) Small flask with cotton plugs Sodium glutamate Microtubes Sterile distilled water B. Culture suspension in skimmed milk Materials Short-term preservation 1.Dissolve all ingredients thoroughly and dispense in small flasks with cotton plugs. 2.Autoclave at 15 psi for 10 minutes. 3.Dispense aseptically in sterile empty microtubes. 4.Scrape bacterial growth from one test tube slant and transfer it into one microtube and homogenize by vortexing. 5.Store the cultures at –20 °C. Storage will take up to 5 years. Procedure 30% glycerol Sterile distilled water Screw-capped microtubes Flask or bottle 10% skim milk (nonfat milk) C. Culture suspension in glycerol Materials Procedure 1.Mix 30% glycerol and water thoroughly and dispense in a bottle or flask. 2.Autoclave at 15 psi for 15 minutes. 3.Dispense the 30% glycerol solution into 2 ml screw-capped microtubes aseptically. 4.Scrape the bacterial growth from the test tube or petri plate and transfer it into the sterilized 30% glycerol solution. (The bacterial growth in one test tube slant is usually transferred into one microtube containing the sterile 30% glycerol solution.) 5.Store the cultures at –80° C for up to 10 years. Freeze-dryer Ampoules Constriction machine Sealing torch Pipette (0.1 ml, 2.0 ml) Test tubes (5 ml capacity) D. Lyophilization (freeze-drying) Materials Long-term preservation Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 9 Scissors Forceps UV-protected eye goggles Face shield Procedure 1. Transfer bacteria into test tube slants and incubate for 48 hours at 30 °C. 2. Prepare 10% skimmed milk and autoclave at 15 psi for 10 minutes. 3. Prepare ampoules with small labels (of isolate) and seal with cotton plugs. 4. Autoclave the sealed ampoules at 15 psi for 15 minutes. 5. Pour 1−1.5 ml of 10% skimmed milk into the 48-hour-old bacterial culture and scrape the bacterial growth. 6. Transfer the scraped bacterial suspension into a sterile empty tube and homogenize by vortexing. 7. Aseptically dispense 0.2 ml of the homogenized bacterial suspension into the individual ampoule. Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 10 Skimmed milk and glycerol stocks 1. Pipette 50 µL of the bacterial suspension from the stock culture onto modified Wakimoto’s agar slants. 2. Incubate at 28°C for two to three days. 3. Check for growth and/or contamination. 4. Transfer by streaking onto modified Wakimoto’s agar slants and incubate for another two to three days at 28°C. 1. Heat the ampoule tip. 2. Place a drop of sterile water on the heated tip to break it open. 3. Remove the cotton plug. 4. Add 200 µL of sterile peptone water. 5. Pour onto a petri dish or disposable plate with modified Wakimoto’s agar. 6. Incubate at 28°C and check for growth after three to four days of incubation. 7. Transfer pure colonies onto modified Wakimoto’s agar slants. 8. Incubate at 28°C for three to four days. Freeze-dried cultures Revival of isolates from storage Inoculum Preparation IV. Inoculation of Xoo using leaf-clip method Materials Culture in slant or plate medium (two to three days old) Sterile distilled water Sterilized wooden stick or wire loop Test tubes or beakers Vortex Latex gloves Ice bucket (with ice) Cut the excess cotton plug above the tip of the ampoule and push down the remaining cotton plug inside the ampoule just above the label. Constrict the ampoule using a constriction machine. Load constricted ampoules in the freeze-dryer and run it for 12 to 14 hours until the cultures are completely dried. Seal the ampoules after freeze-drying with a sealing torch. Store freeze-dried cultures at 4 ºC. 8. 9. 10. 11. 12. Procedure 1. For small-scale inoculation (less than 50 plants for inoculation), one slant culture is enough as an inoculum. 2. After two to three days of incubation of the slant culture, pour 10 ml of distilled water into the single slant culture. 3. Scrape bacterial growth with a sterilized wooden stick or wireloop. 4. Pour the suspension into another clean, empty test tube or falcon tube. 5.Mix the suspension thoroughly by vortexing to obtain a homogeneous mixture. 6. Standardize the concentration of the bacterial inoculum (use 1.0 O.D. at 600nm). 7. Place the tube inside an ice chest with ice. The bacterial suspension is now ready for inoculation. For small-scale inoculation (greenhouse) 11 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 1.For large-scale inoculation in the field, pour the scraped bacterial growth from a single slant culture (48 hours old) into square bottles containing Wakimoto’s medium. 2.Spread the bacterial suspension evenly throughout the medium in bottles to allow uniform bacterial growth. 3. Incubate the bottle at room temperature for three to four days. 4.After incubation, pour one (1) liter of distilled water into each bottle and shake vigorously to remove bacterial masses from the agar surface. 5.Pour the bacterial suspension into another plastic container using a strainer or nylon mesh. This serves as the inoculum in the field. For large-scale inoculation (field) Inoculation of Xoo using leaf-clip method Materials Plants at maximum tillering stage (45-50 DAS) Sterile scissors or clippers 10ml bacterial suspension inside ice chest Test tube rack Paper towels Nitrile or latex gloves Procedure 1. Preferably inoculate fully expanded green leaves (2nd and 3rd position starting from the youngest leaf) of the main tiller per plant. 2. Dip sterile scissors into 10 ml bacterial suspension. 3. Cut the leaf 1-2 cm from the tip. 4. Allow the bacterial suspension to drip along the scissor blades. 5. One dip of scissors will cut 1-2 leaf tips. 6. In the field, clippers are used to cut more leaves per plant. 7. Use a separate pair of scissors for different Xoo isolates. 8. Incubate the inoculated plants in the greenhouse for symptom development. Note In all pathogenicity tests for race designation of Xoo isolates, the standard set of near-isogenic lines (NILs) plus IR24 as susceptible control should be used. The length of BB lesion is measured at 14 d after inoculation, and these data will serve as the basis for identifying the Xoo race existing in a certain locality or rice-growing area. Scoring system for disease assessment BB Assessment in greenhouse condition 1.Assessment of BB is done 14 days after inoculation (DAI) at full symptom expression of the disease. Procedure 12 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases Figure 8. Measuring bacterial blight lesions Note Notice where the measurement should end. Table 2. Prescribed table format of score sheet for BB assessment. 1.Assessment of BB is done 14 days after inoculation (dai) at full symptom expression of the disease. 2.Score the inoculated leaves (2nd and 3rd leaf positions). 3.Determine the disease severity based on lesion length measurement (in cm) or estimation of percent diseased leaf area following the Standard Evaluation System (SES) for Rice, IRRI (2013). Procedure for Data Analysis 1.Calculate the mean and standard deviation per replicate. Another option: consider leaves as replicates (instead of three reps, a total of 12 reps). 2.Prepare a boxplot per isolate per replicate to determine which data points are on the extremes (option to remove). 3.Calculate mean and standarddeviation again. 4.Proceed to prepare clusteredcolumn bar graphsif measures are OK. 5.Convert numerical data (lesion lengths) to categorical data (R, MR, MS, S) (refer to Table 3). 13 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases BB disease assessment in the field 1.Timing of assessment: Natural infection of BLB can be assessed at tillering to the heading stage, usually at 45 to 60 days after transplanting (DAT). Procedure Lesion length (cm) Disease Reaction 0-5 R 6-10 MR 11-14 MS ≥ 15 S Table 3. BB disease assessment scheme under greenhouse test. Lesion length measurements with corresponding disease reaction. Table 4. Prescribed table format of Assessment Form 14 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases Figure 9. Diagrammatic illustration for estimation of BB disease severity Score 1 3 5 7 9 % DLA 1-5% 6-12% 13-15% 26-50% 51-100% Reaction R MR MS S S Selection of samples for assessment: The assessment for disease severity is done by randomly selecting five to seven hills, depending on your plot size using the desired sampling pattern. In each hill, 5 diseased leaves are randomly selected and recorded for the percentage diseased leaf area (DLA) following the IRRI, Standard Evaluation System (SES) for Rice, IRRI (2013). A sample data sheet is provided. Disease severity is assessed based on the estimation of percent diseased leaf area. 2. 3. Scale Diseased leaf area (%) 1 1-5 3 6-12 5 13-25 7 26-50 9 51-100 Table 5. Scale for BB disease assessment under field test adapted from SES manual (IRRI, 2013). Berner D, Viernstein H. 2006. Effects of protective agents on the viability of Lactococcus latis subjected to freeze-thawing and freezing drying. Sci Pharm 74: 137–149. Bureau of Plant Industry, Philippine Rice Research Institute, International Rice Research Institute. 2022. Manual for Crop Health Assessment in the Philippines. Philippine Rice Research Institute, Muñoz, Nueva Ecija, Philippines. Cartwright RD, Groth DE, Wamishe YA, Greer CA, Calvert LA, Vera Cruz CM, Verdier V, Way MO, eds. 2018. Compendium of Rice Diseases and Pests. 2nd Ed. The American Phytopathological Society, St. Paul, Minnesota, U.S.A. IRRI. 2013. Standard Evaluation System for Rice. 5th Edition. International Rice Research Institute. McMaugh T. 2005. Guidelines for surveillance for plant pests in Asia and the Pacific. ACIAR Monograph No. 119, 192p. Mew TW. 1993. Xanthomonas oryzae pathovars on rice: cause of bacterial blight and bacterial leaf streak. London: Chapman and Hall. Mew TW, Hibino H, Savary S, Vera Cruz CM, Opulencia R, Hettel GP, eds. 2018. Rice diseases: Biology and selected management practices. Los Baños (Philippines): International Rice Research Institute. PDF e- book. rice-diseases.irri.org. Mew TW, Misra JK, eds.1994. A Manual of Rice Seed Health Testing. International Rice Research Institute. 113p. Mew TW, Reyes, Vera Cruz CM. 1989. Screening for bacterial blight resistance in rice. In Methods in Phytobacteriology, Z. Klement, K. Rudolph, D.C. Sands (eds.), Akademiae Kiado, Budapest. pp. 338–341. Morgan CA, Herman N, White PA, Vesey G. 2006. Preservation of microorganisms by drying; a review. J Microbiol Methods 66:183–193. Nakasone KK, Peterson SW,Jong SC. 2004. Preservation and distribution of fungal cultures. Biodiversity of Fungi: Inventory and Monitoring Methods (Bills G, Muller GM & Foster MS, eds), pp. 37–47. Elsevier, Amsterdam Niño-Liu DO, Ronald PC, Bogdanove AJ. 2006. Xanthomonas oryzae pathovars: model pathogens of a model crop. Mol. Plant Pathol. 7:303‒324. Noda T, Kaku H. 1999. Growth of Xanthomonas oryzae pv. oryzae in planta and in the guttation fluid of rice. Ann. Phytopathol. Soc. Jpn. 65:9‒14. Reddy PR, Mohanti S. 1981. Epidemiology of the kresek phase of bacterial blight of rice. Plant Dis. 657:578‒ 580. Suwa T. 1962. Studies on the culture media of Xanthomonas oryzae (Uyeda et Ishiyama) Dowson, the causal organism of bacterial blight of rice plant. Ann. Phytopathol. Soc. Jpn. 27(1962):165‒171. Vera Cruz CM, Cottyn B, Nguyen MH, Lang J, Verdier V, Mew TW, Leach JE. 2016. Detection of Xanthomonas oryzae pv. oryzae and X. oryzae pv. oryzicola in rice seeds (Chapter 8). In APS Manual on Detection of Plant-Pathogenic Bacteria in Seed and Other Planting Material, 2nd Ed., M’Barek Fatmi, Ron R. Walcott, and Norman W. Schaad (eds.). APS Press, Minneapolis, MN, ISBN 978-0-89054-539-3. pp. 45‒ 55. Winters RD & Winn WC Jr. 2010. A simple effective method for bacterial culture storage: a brief technical report. J Bacteriol Virol 40:99–101. References We acknowledge previous members of the Plant Pathology and Host Plant Resistance Group for their contributions to the development and optimization of this BB protocol: Marian Hanna R. Nguyen, Pauline C. Capistrano, Jonas J. Padilla, Eula Gems Oreiro, Epifania F. Garcia, Ana Cope, and Ruby Burgos. Acknowledgements Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 15 16 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 1.2 Bacterial leaf streak (BLS) Bacterial leaf streak is caused by Xanthomonas oryzae pv. oryzicola. Infected plants show browning and drying of leaves. Under severe conditions, this could lead to reduced grain weight due to loss of photosynthetic area. Bacterial leaf streak occurs in areas with high temperature and high humidity. It is transmitted through seeds and infected stubbles to the next planting season. It can occur in fields where X. oryzae pv. oryzicola bacteria is present on leaves, in the water, or in the debris left after harvest. Particularly, the disease is common in tropical and subtropical regions of Asia, Africa (including Madagascar), South America, and Australia. It can affect the plant during early stages, from maximum tillering to panicle initiation. Mature rice plants can easily recover from leaf streaks and have minimal grain yield loss. I. Inoculation of Xoc using spraying method in greenhouse condition Inoculum Preparation Culture in slant or plate medium (two to three days old) Sterile distilled water Sterilized wooden stick or wire loop Test tubes or beakers Vortex Latex gloves Ice bucket (with ice) Materials Authors: Genelou Atienza-Grande, Ma. Michelle Valenzuela, Van Schepler-Luu 17 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 1. Inoculation is preferably done between 10:00 am to 1:00 pm at high sun exposure. 2.Transfer bacterial suspension (10ml is enough for one plant) into a spray bottle. 3.Spray inoculum generously onto the adaxial (upper side) and abaxial (lower side) surfaces of the leaf blade covering the entire area from the tip down to its base. 4. Inoculum should be sprayed as a fine mist to facilitate absorption by stomata on the leaf surface. 5.Cover the inoculated plants with plastic or maintain a high relative humidity within the inoculation area. 6. Incubate the inoculated plants in the greenhouse for symptom development. Procedure Note To avoid contamination (for multiple isolates) and to ensure all plants receive the same amount of inoculum, maintaining approximately one meter distance between samples. Upon the appearance of initial symptoms (dark green and translucent streaks), employ additional spraying of water onto the leaf surface in order to facilitate the spread of the bacteria. 1. For small-scale inoculation, one plate culture is enough for inoculation of 10 plants (one plate = 100ml bacterial inoculum). 2. After two to three days of incubation of the culture, pour 20 ml of distilled water into the single plate culture. 3. Scrape bacterial growth with a sterilized wooden stick or wireloop. 4. Pour the suspension into another clean, empty flask or falcon tube. 5. Add the remaining 80 ml of sterile distilled water into the bacterial suspension. 6. Mix the suspension thoroughly by vortexing to obtain a homogeneous mixture. 7. Standardize the concentration of the bacterial inoculum (use 1.0 O.D. at 600nm) 8. Keep the bacterial suspension on ice until inoculation. Procedure Inoculation of Xoc using spraying method Plants at maximum tillering stage (45-50 DAS) Spray bottles 100ml bacterial suspension inside ice chest Paper towels Nitrile or latex gloves Materials A. Disease estimation of percent diseased leaf area using SES 1.Assessment of BLS is done 11 to 14 days after inoculation (DAI) at full symptom expression of the disease. 2.Score the inoculated leaves with the most visible symptoms (2nd or 3rd leaf position). 3.Determine the disease severity based on the estimation of the percent diseased leaf area according to the assessment scheme used in the Standard Evaluation System (SES) for Rice (IRRI, 2013). Figure 10. Index value and the corresponding levels if severity for a leaf streak disease. SES for Rice, IRRI (2013) II. BLS assessment for greenhouse test 18 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases B. Step-by-step protocol for quantification of diseased leaf areas using ImageJ software Preparation of the diseased samples 1.Fourteen days post-inoculation, collect the leaf with the most visible symptoms. 2.Keep the leaves in a test tube with water to prevent them from wilting. 3.Cut the tip of the leaves. Measure 25 cm, keeping the portion with the most symptoms. Prepare the laminating film (A4 size). Put a double-sided tape (as small as possible) on the upper side of the film to keep the leaves from moving (refer to Figure 11). Figure 11. Recommended layout of the samples. 19 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases Place the leaves inside the laminating film. Use a small portion of double- sided tape to secure the bottom of the leaves. Turn on the laminator and wait for it to heat up. Once ready, place the film on the laminator. Pass the film containing the samples through the machine twice to ensure they are airtight. This prevents molds from growing on the samples during storage. To prepare the samples for scoring, scan the laminated mounted leaf samples together with a 12-inch ruler beside the sample and save as .jpeg file. Alternatively, take a photo of the laminated mounted samples placed on top of a lightbox. 4. 5. 6. 7. 8. Quantification of diseased leaf area using ImageJ software 1.Download and install ImageJ. 2.Under "File", click "Open" and choose the captured photo of the sample. Calibration 1.Use the "Zoom" function under the "Image". 2.Using the "Line" tool and draw a line measuring 1 cm on the ruler. 3.Go to "Analyze" then "Set Scale". 4.Change the "known distance" to "1.00" and the "unit of length" to "cm". 5.Select "Global" and press "OK". Determining percentage of leaf disease damage 1.To measure the total leaf area, go back to the photo. Using the "Rectangle" tool, select one leaf sample. Use "Duplicate" under the "Image" or simply use the function Ctrl+Shift+D. Go to "Image" then "Adjust". Choose "Color Threshold". Adjust the "Hue" (0-255), "Saturation" (30-255), and "Brightness" (0-255) such that only the whole leaf is colored in red. 2. 3. Click "Select". You will see the whole leaf area is outlined in white. Close the window. 4. Go to "Process" and choose Binary, then choose "Make Binary". Remove outliers by going to "Process" > "Noise" > "Remove Outliers" with Radius between 1-5. Tick "Preview" and adjust the "Radius" accordingly. 5. Go to "Analyze", then “Set measurements”, then tick “Area” and “Limit to threshold” Go to “Analyze” and select "Measure". Alternatively, use "Ctrl+M". To measure the damaged area/area with lesions, follow the same steps used in measuring the total leaf area. However, you now need to adjust the color thresholds with "Hue" (25-45), "Saturation" (120-255), and "Brightness" (adjustable) such that only the areas with lesions are selected. 6. 7. 8. 20 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases To compute the leaf disease damage percentage, use following formula:9. 21 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases Leaf disease damage percentage (%) = ( )damaged area total leaf area x 100 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases Bureau of Plant Industry, Philippine Rice Research Institute, International Rice Research Institute. 2022. Manual for Crop Health Assessment in the Philippines. Philippine Rice Research Institute, Muñoz, Nueva Ecija, Philippines. IRRI. 2013. Standard Evaluation System for Rice. 5th Edition. International Rice Research Institute. Mew TW, Hibino H, Savary S, Vera Cruz CM, Opulencia R, Hettel GP, eds. 2018. Rice diseases: Biology and selected management practices. Los Baños (Philippines): International Rice Research Institute. PDF e- book. rice-diseases.irri.org. Mew TW, Misra JK, eds.1994. A Manual of Rice Seed Health Testing. International Rice Research Institute. 113p. Niño-Liu DO, Ronald PC, Bogdanove AJ. 2006. Xanthomonas oryzae pathovars: model pathogens of a model crop. Mol. Plant Pathol. 7:303‒324. Reddy PR, Mohanti S. 1981. Epidemiology of the kresek phase of bacterial blight of rice. Plant Dis. 657:578‒ 580. Vera Cruz CM, Cottyn B, Nguyen MH, Lang J, Verdier V, Mew TW, Leach JE. 2017. Detection of Xanthomonas oryzae pv. oryzae and Xanthomonas oryzae pv. oryzicola in rice seeds. Chapter 8 in APS Manual on Detection of Plant Pathogenic Bacteria in Seed and Planting Material. APS Press. References We acknowledge previous members of the Plant Pathology and Host Plant Resistance Group for their contributions to the development and optimization of this BLS protocol: Marian Hanna R. Nguyen, Pauline C. Capistrano, Jonas J. Padilla, Eula Gems Oreiro, Epifania F. Garcia, Ana Cope, and Ruby Burgos. Acknowledgements 22 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 1.3 Rice Blast 23 Screening Protocol I. Field sampling protocol for leaf and neck blast 1.Before carrying out the collection, make sure that you obtain official documents that allow collecting biological materials from farmers’ fields. 2. Identify an infected field, talk to farmers to collect the passport information (Table 1). 3.To determine the size where the survey will be conducted, use stratified sampling techniques and hierarchical scheme within the selected hotspot target areas. This includes four levels of sampling, to divide the target sites into logical categories (refer to Figure 4 and 5). a.Level 1 (Field): From an identified symptomatic field, determine the sampling units using the W pattern. The sampling unit is defined as a hill. Five hills are sampled from fields with an area of less than one hectare and eight hills are sampled from larger fields. b.Level 2 (Village): In cases where infections are spread in neighboring fields, identify 3-4 nearby fields that are 1 km apart from each other. Prioritize the diversity of rice varieties grown and select 3-4 different rice varieties. c.Level 3 (District): Search for other villages nearby that have infected fields. Follow the village collection scheme. d.Level 4 (Division): Search for 1-3 districts nearby that have infected fields. Follow the district collection scheme. 4.From each sampled hill, collect five leaves and three panicles with young typical lesions with gray center and infected panicle base 5.Place the leaves and panicles from one hill in separate envelopes to avoid mixing them with samples from other hills Authors: Mary Jeanie Telebanco-Yanoria, Genelou Grande, Casiana Vera Cruz, Van Schepler- Luu Rice blast is one of the most destructive diseases of rice, caused by the fungal pathogen Magnaporthe oryzae. It was first reported in China and Japan over 300 years ago and has since spread globally, affecting rice production in both temperate and tropical regions. The disease manifests as diamond-shaped lesions with gray or whitish centers and brown margins on leaves, while it can also infect nodes, panicles, and collars, leading to severe plant damage. Under favorable conditions, rice blast can cause significant yield losses, ranging from 10% to 30% in moderate cases and up to 100% in severe outbreaks, posing a major threat to global food security. II. Single spore isolation 1. Cut 3.5 cm from the infected leaf/panicle samples with a typical blast lesion. 2. Place two to three cut sections in a sterile glass slide in a sterile petri dish with moistened filter paper. 3. Incubate the plates in a plastic box for about 18 to 24 hours at room temperature (25°C to 28°C). 4. Examine the rice blast lesions under a dissecting microscope to observe the sporulating lesion. 5. Use a 9-inch Pasteur pipette with a very fine tip (tip melted by heating with a gas burner, pulled and cooled). 6. Pick lightly the top of the conidial masses by touching it with the tip of the Pasteur pipette and then spread it gently on a water agar plate; distances between spores should allow you to pick up a single spore. 7. Pick a single spore and transfer it to another spot on the water agar plate. 8. Incubate the plates at 28°C for three to four days in order to see the growth of macroscopic mycelium. 9.Cut a fungal mycelium and transfer it to a prune slant/plate. 10.Incubate for 5-7 days prior to stock culture for long-term storage. Remove filled or unfilled spikelets, and cut panicles near the infected neck to fit into the envelope. Do not sample hills near the border of a field to avoid bias caused by interplot interference. This occurs when the pathogen or disease intensity is affected by adjacent plots or external factors. Properly label each envelope with the sample ID (refer to Figure 6) Place the coin envelopes inside a plastic bag and seal with tape. Immediately place the plastic bags (with coin envelopes) inside a chest filled with ice while in the field. In the lab, aerate or air-dry all samples collected for 4-8 hours and kept in -20°C. 6. 7. 8. 9. 10. 11. Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 24 Figure 12. Macroscopic mycelium 5-6 days after single spore isolation III. Preparation of stock culture for long-term storage (-20°C) 1. Place two pieces of pre-cut sterile Whatman no. 1 filter paper (equally cut into four pieces) on the surface of a prune agar plate. 2. Cut a fungal mycelium block 5x15 cm of a 6-7 day old culture of M. oryzae using a transfer needle/spatula. 3. Transfer it into a sterile test tube with 5 ml sterile distilled water. 4. Macerate the fungal mycelium block and pour about 1.5 ml into an agar prune plate lined with sterile filter paper. See to it that the filter paper is covered with the suspension then incubate for 5-7 days at 28°C until the filter paper is covered by the growth of the fungal mycelium. 5. Remove or lift the colonized filter paper carefully and transfer it to a dry sterile plate lined with filter paper. 6. Stack the plates in a box and dried for two weeks under continuous light (fluorescent lamp) in a customized cabinet chamber. 7. When the filter paper is completely dried, cut aseptically into small paper disks at 3-5 mm . 8. Transfer the small filter papers in sterilized 2 ml microcentrifuge tubes using alcohol-flamed sterilized forceps. Make at least two tubes for each isolate one is for working (used for reviving) and the other one for storage only. 9. Tubes must be labeled properly (isolate name and date of isolation). 10. Keep the microcentrifuge tubes in a plastic DNA box and store them at -20°C or -80°C freezer. 2 25 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases IV. Medium Preparation Yeast extract, 2g Sucrose, 5g Agar, 30g Distilled water, 1000 ml Water Agar 1.Soak 20g of rabbit food in 500 ml distilled water for five minutes. 2.Mix and stir the mixture and add 500 ml of distilled water. 3.Get the extract by filtering it with nylon mesh. Volume it up to 1000 ml and add 20 g of agar. 4.Plug the flask with cotton and then autoclave the media at 121°C for 20 minutes at 15 psi. 5.Before plating add a pinch of Streptomycin for every liter of media then dispense the melted media in a plate (approximately 20 plates/liter or about 50 ml/plate) and in slant (9 ml). Procedure *Autoclaved at 121°C for 20 minutes V. Plant Preparation and Maintenance 1.Soak seeds three days before sowing. Sow seeds in sterilized soil with basal fertilizer (complete) and grow for 14 days. If directly seeded, allow 17 days for growing seedlings (4th leaf stages). All plants are maintained in a greenhouse condition and seedlings are maintained healthy and free from any disease and insect damage. 2.Two days before inoculation (i.e. 12 DAS), apply ammonium sulfate to promote vegetative growth. Figure 13. Sample growing seedling set up 26 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases VI. Fungal Culture Preparation 1. Revive fungal isolate from filter paper stock for seven to 10 days in prune agar plates (this should be done concurrently with the seed soaking). 2. After seven to 10 days, subculture the revived isolate into prune agar plates and allow fungal growth for seven to 10 days. 3. Three days before inoculation, scrape the mycelial growth from the fungal culture and expose it under white/fluorescent light for three days. Remove the cover of the Petri plates. 4. Prepare the inoculum by dispensing about 15 ml of sterile distilled water with Tween 20 (500-600 ul Tween 20 to 1L of sterile distilled water) and scrape gently the surface of the plates using a sterile glass slide. 5. Filter the spore suspension in sterile miracloth. 6. Adjust the spore concentration to 1.5x 105 spores/ml. Place the suspension in ice to prevent or delay spore germination. Table 6. Sample inoculation schedule 27 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases How to use the hemacytometer? 1.Mix spore solution well. 2.Add 10 µl of spore solution to each side of the hemacytometer. 3.Count number of spores in zones A, B, C, D and E on both sides of the hemacytometer, record them, and calculate the average of the two sides. a. If a spore falls on the left or bottom line DO NOT count it. b. If a spore falls on the right or top line DO count it Figure 14. How to use the hemacytometer 4.To determine the number of spores per mL a. (A + B + C + D + E) x 2000 (dilution factor) 28 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 5. Dilute the spore solution to desired concentration (spores/mL) a. Use C1V1= C2V2 formula 67,000 x = 40 * 50,000 x = 2,000,000 = 29.85 ~=30mL 67,000 Add 30 mL spore solution to 10 mL of sterile dH20 (40-30=10) to give a final concentration of 50,000 spores/mL in 40 mL. How to calculate the concentration of spores in a fungal spore suspension?. Available from: https://www.researchgate.net/post/How_to_calculate_the_concentration_of_spor es_in_a_fungal_spore_suspension [ September 30, 2024] https://www.researchgate.net/post/How_to_calculate_the_concentration_of_spores_in_a_fungal_spore_suspension https://www.researchgate.net/post/How_to_calculate_the_concentration_of_spores_in_a_fungal_spore_suspension VII. Leaf blast inoculation by spraying method 1.Spray the spore suspension using the airbrush attached to a compressor (see Figure 14). Clean the airbrush using 70% Ethanol then rinse with sterile distilled water. 2.Make sure to cover the leaves evenly with a fine mist to mimic natural blast infection. 3.Place inoculated plants in a large plastic box and cover with the box lid. 4. Incubate the plants inside the moist jute sack chamber for 24 hours at ~25°C. 5.After 24 hours, take out the plants and place them inside the mist room (~23 °C- 24 °C) and replace the box lid with plastic film until disease assessment (6 dpi). Reference: https://olympos-airbrush.ocnk.net/product/807 Figure 15. Compressor and airbrush used in spray inoculation VIII. Disease Assessment 29 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases Evaluate the plants at six days post-inoculation (dpi) using the scoring method by JIRCAS (based on lesion types; please refer to Figure 15). 1. Figure 16. Scoring method by JIRCAS 30 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases Table 7. Sample data sheet for leaf blast 31 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 32 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases Take a representative leaf sample from each line and mount it in a double- sided tape (see Figure 17) for future reference. 2. Figure 17. Sample data sheet for mounting leaf sample from each line 33 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases IX. Neck Blast Artificial Disease Screening 1.Seven to ten days in a plate with prune agar medium. 2.For culture multiplication, cut a 0.5 x 0.5 cm portion of the mycelial growth from a grown stock culture and transfer it into a new plated prune agar medium. 3.Incubate the plates at 28°C for eight to ten days. 4.Scrape gently the surface using a sterile glass slide and expose the opened plate to a continuous light for three days to induce the sporulation. 5.Three days after exposure to light, collect the plates and put back their cover. 6.Pour about 10 ml of sterile distilled water per plate and then harvest the spores by gently scraping the surface with a glass slide to dislodge the spores and strain it using a miracloth or a double-layered gauze cloth. 7.Adjust the spore concentration to 100,000 to 150,000 spores/ml. 8.Keep the prepared inoculum suspension in ice or maintain it at 4°C condition for two to three hours during the whole inoculation period. Otherwise, prepare a new inoculum suspension beyond four hours. Inoculum preparation Inoculation by Injection Method 1. Inoculation is done one to two weeks after flowering. Prior to inoculation, plants are cleaned by removing old and dried leaves to prevent insect infestation during incubation. 2.Select at least five panicles per plant for inoculation and label it properly. 3. Inject 100 to 200 µl per panicle (depending on the size of panicle) about 5-10 cm below the panicle base using a 1 ml syringe with a small needle. 4.After inoculation, the inoculated panicles are covered with a transparent lightweight plastic bag for 24 hours to maintain a relative humidity of at least 80 %. 5. Inoculated plants are kept in an air-conditioned room with a temperature of 21 ± 4 C for 10 days. 6.Ten days after inoculation (DAI), each panicle is evaluated (see Table 8 for sample data sheet for neck blast) for disease infection following the IRRI SES (2014) for rice panicle blast (refer to Figure 19). 34 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases A. B. Figure 18. (A) Injection site below the base of the panicle (5-10cm), (B) Inoculated panicles are covered with a lightweight plastic bag for 24 hours; (C) Inoculated plants incubated in an air condition room at 24 to 26⁰C for 10 days; (D) Disease assessment 10 days after inoculation. C. D. Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 35 Table 8. Sample data sheet for neck blast Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 36 Figure 19. Modified Evaluation scale for neck blast disease (IRRI Plant Pathology Laboratory, 2019) Hayashi N, Kobayashi N, Vera Cruz CM, Fukuta Y. 2009. Protocols for the sampling of diseased specimens and evaluation of blast disease in rice. JIRCAS Working Report 63:17–33. IRRI. 2013. Standard Evaluation System for Rice. 5th Edition. International Rice Research Institute. References We acknowledge previous members of the Plant Pathology and Host Plant Resistance Group (IRRI) and JIRCAS for their contributions to the development and optimization of the leaf blast inoculations protocol: Ellen Silab, Casiana Vera Cruz Yoshimichi Fukuta, Nagao Hayashi, John M. Bonman. We acknowledge members of the Plant Pathology and Host Plant Resistance Group (IRRI) for their contributions to the development and optimization of the neck blast inoculations protocol: Ellen Silab for the isolate stocks, Melencio Apostol for culture preparations and Neil Bryan Fondevilla for inoculation and evaluation. Acknowledgements Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 37 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 38 1.4 Sheath Blight (ShB) Rice sheath blight is a major fungal disease of rice caused by Rhizoctonia solani, a soilborne pathogen with a wide host range. The disease was first identified in Japan in the early 20th century and has since become a serious concern in rice-growing regions worldwide, particularly in intensive production systems. Symptoms include water-soaked, oval or irregular lesions on the leaf sheath, which enlarge and coalesce, forming large necrotic areas that can girdle the stem. Under favorable conditions, sheath blight can lead to premature senescence, reduced tillering, and poor grain filling. Yield losses typically range from 5% to 30%, but severe infections under high humidity and dense canopy conditions can cause losses exceeding 50%, significantly impacting rice productivity. Screening Protocol I. Sampling Follow the sampling protocol described for bacterial blight and blast. Briefly: 1.Before carrying out the collection, make sure that you obtain official documents that allow collecting biological materials from farmers’ fields. 2. Identify an infected field, talk to farmers to collect the passport information (Table 1). 3.To determine the size where the survey will be conducted, use stratified sampling techniques and hierarchical scheme within the selected hotspot target areas. This includes four levels of sampling, to divide the target sites into logical categories (refer to Figure 4 and 5). Authors: Mary Jeanie Telebanco-Yanoria, Van Schepler-Luu II. Isolation 1.Cut several pieces of 0.5 to 1 cm of tissue from the infected tiller samples and choose a young typical ShB lesion. 2.Surface sterilize the cut tissue with 5% Sodium Hypochlorite (NaOCI) solution or 70% Ethanol for 3 to 5 minutes and rinse it twice with sterile distilled water. 3.Blot dry the cut tissues with a sterile paper towel and keep them in a sterile petri dish. 4.Place one cut section on a PDA plate. Make five plates for each sample. 5. Incubate the plates at 28 °C for 3 to 5 days. 6.Examine the growth of typical R. solani mycelia or the presence of sclerotial bodies. 7.Cut 3 to 5 cm agar block from advancing mycelial growth and transfer into new PDA plate and incubate for 5 to 7 days until sclerotia are formed. 8.To purify, pick one sclerotium and transfer it to a new PDA plate. 9. Incubate the plates at 28 °C for 5 to 7 days until sclerotia are formed and this will be used for the stock culture for long-term storage. Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 39 a.Level 1 (Field): From an identified symptomatic field, determine the sampling units using the W pattern. The sampling unit is defined as a hill. Five hills are sampled from fields with an area of less than one hectare and eight hills are sampled from larger fields. b.Level 2 (Village): In cases where infections are spread in neighboring fields, identify 3-4 nearby fields that are 1 km apart from each other. Prioritize the diversity of rice varieties grown and select 3-4 different rice varieties. c.Level 3 (District): Search for other villages nearby that have infected fields. Follow the village collection scheme. d.Level 4 (Division): Search for 1-3 districts nearby that have infected fields. Follow the district collection scheme. From each sampled hill, collect infected tillers with typical ShB lesions. Choose a young ShB lesion. Place the tillers from one hill in separate envelopes to avoid mixing them with samples from other hills. Do not sample hills near the border of a field to avoid bias caused by interplot interference. This occurs when the pathogen or disease intensity is affected by adjacent plots or external factors. Properly label each envelope with the sample ID (refer to Figure 6) Place the coin envelopes inside a plastic bag and seal with tape. Immediately place the plastic bags (with coin envelopes) inside a chest filled with ice while in the field. In the lab, aerate or air-dry all samples collected for 4-8 hours and kept in -20°C. 4. 5. 6. 7. 8. 9. 10. III. Preparation of stock culture for long term storage (-20°C) 1. Place two pieces of pre-cut sterile Whatman no. 1 filter paper (equally cut into four pieces) on the surface of the Potato Dextrose Agar Plate. 2. Cut a fungal mycelium block 5x15 cm or scleotria of a five to seven days old culture of R. Solani using a transfer needle/spatula. 3. Transfer the agar block into a sterile test tube with 2.5 ml sterile distilled water. 4. Macerate the fungal mycelium block and pour about 1.5 ml on a PDA plate lined with sterile filter paper. See to it that the filter paper is covered with the suspension then incubate for 5 days at 28 °C until the filter paper is covered by the growth of the fungal mycelium. Or just pick 3-6 sclerotia and place them beside the filter paper and allow it to grow for 5 days until the whole filter paper is covered with mycelia. 5. Remove or lift the colonized filter paper carefully and transfer it to a dry sterile plate lined with filter paper. 6.Stack the plates in a box and dry them for two weeks under continuous light (fluorescent lamp) in a customized cabinet chamber. 7.When the filter paper is completely dried, cut aseptically into small paper disks at 3-5 mm . 8.Transfer the small filter papers in sterilized 2 ml microcentrifuge tubes using alcohol-flamed sterilized forceps. Make at least two tubes for each isolate one is for working (used for reviving) and the other one for storage only. 9.Tubes must be labeled properly (isolate name and date of isolation). 10.Keep the microcentrifuge tubes in a plastic DNA box and store them at -20°C or -80°C freezer. IV. Medium Preparation Yeast extract, 2g Sucrose, 5g Agar, 30g Distilled water, 1000ml Water Agar 1.Weigh and put all components and add into a two-liter Erlenmeyer flask and add one liter of distilled water. 2.Autoclaved at 121°C for 20 minutes. 3.Allow to cool down the medium and dispense thinly (about 25ml in a plastic disposable plate) just enough to cover the surface. 4.Allow the plates to dry completely before using. Procedure Water agar components and procedure were modified from Hayashi et al., 2009. Potato, 200g Agar, 20g Dextorose, 20g Distilled water, 1000 ml Potato Dextrose Agar Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 40 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 41 1.Boil 200 grams of sliced unpeeled potatoes in 500 ml of distilled water for 30 minutes. Collect the potato extract by filtering through nylon mesh. 2.Mix the dextrose, agar, and potato extract in a two-liter flask and make the volume to 1000 ml. 3.Allow to cool down the medium and dispense thinly (about 25ml in a plastic disposable plate) just enough to cover the surface. 4.Allow the plates to dry completely before using 5.Sterilize media by autoclaving at 121ºC for 15 minutes. Procedure *Autoclaved at 121°C for 20 minutes. V. Preparation of Plant Materials 1.Acclimatize the seeds by oven-drying them at 50°C for five days. Sow the acclimatized seeds directly in pots with a diameter of 200 mm. Place all pots in wooden trays lined with linoleum sheets, and fill the trays with water to saturate the soil in the pots. 2.Each pot should contain three hills, and sow one seed per hill for each genotype. Thin out two seedlings before inoculation to leave one per hill. 3.For the detached tiller test, flood the pots two days before tiller sampling to facilitate uprooting. Wash the roots of the tillers with tap water and place the tillers in large test tubes filled with 30 ml of distilled or RO water. 1.Use prepared Potato Dextrose Agar (PDA) plates to transfer R. solani culture for this experiment. PDA preparation requires 200 g of potato, 20 g of dextrose, and 20 g of agar is needed. Follow the instructions below: a.Boil 200 grams of sliced unpeeled potatoes in 500 ml of distilled water for 30 minutes. b.Collect the potato extract by filtering through cheesecloth. c.Mix the dextrose, agar, and potato extract in a 2-L flask and make the volume to 1,000 ml. d.Sterilize media by autoclaving at 121 ºC for 15 minutes. 2.Select a sclerotium with a diameter of 1.5 ± 0.3 mm from the seven-day-old culture as the standard inoculum. 3. Incubate the plates at 28°C for seven days. 4.Choose sclerotia from the peripheral edge of the plates. Identify the second sheath collar of the tiller from the topmost part as the inoculation site (mark this with a pen) (refer to figure 18A). 5.Select the second collar below the tip of the culm as the inoculation site. Insert the inoculum 1 cm below the leaf collar, on the inner side of the sheath. Place the inoculated tillers in tubes (25 x 200 mm). VI. Inoculation A. Detached tiller assay (in a plant growth chamber) Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 42 Conduct inoculation during the maximum tillering stage (40-45 days after transplanting) of the rice plants. Place the inoculated tillers in tubes (25 x 200 mm) immediately after inoculation (refer to figure 18B). All inoculated tillers are incubated in a Conviron growth chamber with 25°C temperature and 12- hour photoperiod at a light intensity of 200 μmol setting. This can also be done in pots and inoculated plants are kept at normal GH house conditions with misting three to four times a day. Keep the relative humidity between 85-90%, and provide moisture by setting the misting system to spray every three hours for one to two minutes (refer to figure 18C). Place the pots and inoculated detached tillers inside the Conviron growth chambers immediately after inoculation. Set the temperature to 25°C and maintain a 12-hour photoperiod at 200 µmol. Keep the relative humidity between 85-90%, and provide moisture by setting the misting system to spray every three hours for one to two minutes (refer to figure 18C). 6. 7. 8. 9. Figure 20. Phenotypic evaluation under controlled conditions using detached tiller assay (DTA). (A) Rhizoctonia Solani seven-day-old culture; (B) DTA inoculation where sclerotium was inserted in the sheath collar of the tiller; (C) Plants placed inside the Conviron growth chamber with 25°C temperature and 12-hour photoperiod at a light intensity of 200 μmol setting. 43 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases Conduct the disease assessment seven to ten days after inoculation. For each tiller, record the following data (refer to Table 9): 1.Development Stage 2.Length of culm (mm) 3.Number of lesions (count only distinct lesions) 4.Maximum lesion Length (mm) 5.Vertical sheath colonization (mm) Disease Assessment Table 9. Sample datasheet used for Detached Tiller Assay (DTA) B. Rice Grain - Rice Hull (RGRH) Mixture 1.Sterilize bottles containing an RGRH mixture (refer to Figure 21A) in an autoclave at 121 °C and 15 psi for 1 hour; repeat this process twice. 2.After cooling, seed the inoculum in the bottles by placing a quarter of a Petri plate with a 5-day-old LR1 isolate inside each bottle. 3.Place the seeded bottles in an incubator at room temperature (28 or 29 °C). 4.The approximate age of the inoculum for field inoculation should be 7 to 10 days old (refer to Figure 21B). 5.Before field inoculation, place 5 g of the RHGM inoculum on aluminum foil for ease of application. 6.During inoculation, place the inoculum between the tillers of the central hill in each microplot (refer to Figure 21C). 7.The microfield layout is shown in Figure 22. Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 44 The experimental layout in Figure 22 considers the epidemiology of the disease, the vertical spread (disease intensification), and the horizontal spread (disease extensification). Vertical spread of the pathogen or the disease spread within the plant is assessed at 12 days after inoculation (DAI). This quantifies the level of physiological resistance using disease complementary variables. Whereas the disease spread between-plant or the horizontal spread of the pathogen quantifies both physiological resistance and disease escape. It considers the role of morphology, i.e., plant height and tiller number. Horizontal spread is observed twice at 22 and 32 DAI. See Table 10 for the variables recorded in the micro field experiment and Table 11 for sample datasheet. 8. Note Source or inoculated hill: Instead of using Swarna as the source hill, the test genotype itself will be inoculated and will serve as the source hill. There will be three assessments in this experiment at 12, 22, and 32 DAI when the following measurements will be recorded: plant height (cm), tiller incidence, leaf incidence, lesion number, and lesion height (cm). 9. Leaf incidence is an important parameter for this experiment especially during the 2nd and 3rd assessments (on the neighboring hills) since the pathogen requires tissue contacts (leaf to leaf or leaf to sheath) for it to spread horizontally. In getting the absolute value for the leaf incidence, we use: Conditional leaf incidence value = (Actual number of infected leaves from three diseased tillers/total number of leaves from three tillers)/3 Absolute leaf incidence value = Conditional value (decimal form) x tiller incidence (decimal form) A. B. C. Figure 21. Micro-field experiment. (A) Experimental set-up of selected genotypes replicated three times laid in randomized complete block design at CS04 screenhouse with misting set-up every 10 min at intervals of 3–4h; (B) Rice grain rice hull mixture (RGRH) for inoculation containing 1part rice grain:3 parts rice hull and a quarter of a week old Rhizoctonia solani culture in potato dextrose agar; and (C) Plants at maximum tillering stage inoculated by inserting 5 g of RGRH at the base of the hill Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 45 Figure 22. Microfield layout 46 Table 10. List of parameters assessed in detached tiller assay (DTA) and Microfield (MF) experiments Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases Table 11. Sample datasheets used for detached tiller assay (DTA) and Microfield (MF) experiments C. Corn Meal Sand (CMS) Mixture Used in high-throughput phenotyping method Mix the crushed corn seeds or corn grits in a flask or bottle with fine sand in a 1:3 ratio, three parts of corn grits and one part of fine sand (ex. 50 grams of sand and 150 grams of corn grits). Soak the mixture in distilled water with 400g sucrose for 30 minutes. Dispense 200g of CMS mixture in each bottle. Autoclave the CMS mixture at 121°C (15 psi) for 30 minutes. Repeat the sterilization twice at 24 hours alternate. After cooling, seeding of inoculum in the CMS mixture will be done by placing half of the Petri plate of five-day-old LR1 isolate inside each bottle. Incubate the seeded CMS bottles at 28-29°C for 10 days and observe for hyphal growth and spread (figure 21A). Before field inoculation, 40g of CMS inoculum will be placed in an aluminum foil for ease of application. The inoculum will be applied through broadcasting in a 1m area in a microplot (figure 21B).2 Figure 23. Corn meal sand (CMS) mixture used in high-throughput method. (A) CMS mixture colonized with R. Solani, 10 days after incubation, and (B) field inoculation through broadcast application. (Note: 1m = 40 grams; 1m = 25 plants (if P.D. is 20 cm x 20 cm) 2 2 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 47 Lore JS, Hunjan MS, Singh P, Willocquet L, Sri, S, Savary S. 2013. Phenotyping of partial physiological resistance to rice sheath blight. J. Phytopathol. 161:224−229. Willocquet L, Lore JS, Srinivasachary S, Savary S. 2011. Quantification of the components of resistance to rice sheath blight using a detached tiller test under controlled conditions. Plant Dis. 95:1507−1515. Srinivasachary, Beligan G, Willocquet L, Savary S. 2013. A strategy to identify sources of quantitative resistance in pathosystems involving disease escape and physiological resistance: The case study of rice sheath blight. Plant Pathol. 62:888−899. References We acknowledge previous members of the Plant Pathology and Host Plant Resistance Group for their contributions to the development and optimization of this sheath blight inoculation protocol: Eula Gems Oreiro, Laetitia Willocquet, Francisco Elazegui, Ellen Silab. Acknowledgements Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 48 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 49 1.5 Brown Spot (BS) Brown spot fungal disease may occur on any part of the rice plant at any stage of the crop, although the disease is commonly observed in mature plants. The leaves after infection show elliptical spots, which gradually increase in size and eventually reach the grains. The spots are brown, which may reach 1 cm or more in length in susceptible cultivars. Pin-point size brownish spots are observed in resistant cultivars BS is caused by the fungus, Bipolaris oryzae (formerly known as Helminthosporium oryzae). The causal agent can be easily identified by looking at the conidia or spores under a dissecting microscope. The conidia of B. oryzae are very small, club-shaped to cylindrical, generally curved, light brown to golden brown, and with 6 to 14 transverse cell walls. This fungus is seedborne and can also survive on infected rice straw and stubble. It spreads from plant to plant in the field by airborne spores. High relative humidity (86−100%) and temperatures between 20 and 26 °C favor disease development. For infection to occur, leaves require continuous wetness for from 8 to 24 hours. Isolating the fungus that causes the specific symptoms/disease requires suitable techniques to obtain a pure culture of the pathogen and thereby observe its pathogenic ability on plants. Isolation using direct-tissue planting is a common method used with plant pathogenic fungi. However, in the case of BS, single-spore isolation is best because the fungus could easily be overrun by other fast-growing fungi or saprophytes. This method will allow a single spore to germinate and further produce infective structures, making it a good source of pure inoculum for pathogenicity tests. Moreover, other fungal contaminants are separated. Authors: Mary Jeanie Telebanco-Yanoria, Genelou Atienza-Grande, Rafael Luis Natulan, Van Schepler-Luu Screening Protocol Follow the sampling protocol described for bacterial blight and blast. Briefly: 1.Before carrying out the collection, make sure that you obtain official documents that allow collecting biological materials from farmers’ fields. 2. Identify an infected field, talk to farmers to collect the passport information (Table 1). 3.To determine the size where the survey will be conducted, use stratified sampling techniques and hierarchical scheme within the selected hotspot target areas. This includes four levels of sampling, to divide the target sites into logical categories (refer to Figure 4 and 5). a.Level 1 (Field): From an identified symptomatic field, determine the sampling units using the W pattern. The sampling unit is defined as a hill. Five hills are sampled from fields with an area of less than one hectare and eight hills are sampled from larger fields. b.Level 2 (Village): In cases where infections are spread in neighboring fields, identify 3-4 nearby fields that are 1 km apart from each other. Prioritize the diversity of rice varieties grown and select 3-4 different rice varieties. c.Level 3 (District): Search for other villages nearby that have infected fields. Follow the village collection scheme. d.Level 4 (Division): Search for 1-3 districts nearby that have infected fields. Follow the district collection scheme. 4.From each sampled hill, collect five leaves with typical brown symptoms with circular brown lesions. 5.Place the leaves from one hill in separate envelopes to avoid mixing them with samples from other hills. 6.Do not sample hills near the border of a field to avoid bias caused by interplot interference. This occurs when the pathogen or disease intensity is affected by adjacent plots or external factors. 7.Properly label each envelope with the sample ID (refer to Figure 6) 8.Place the coin envelopes inside a plastic bag and seal with tape. 9. Immediately place the plastic bags (with coin envelopes) inside a chest filled with ice while in the field. 10.In the lab, aerate or air-dry all samples collected for 4-8 hours and kept in -20°C. I. Sampling Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 50 II. Single spore isolation 1.Cut 3.5 cm from the infected leaf/panicle samples with a typical brown spot lesion. 2.Place two to three cut sections in a sterile glass slide in a sterile petri dish with moistened filter paper. 3. Incubate the plates in a plastic box for about 18 to 24 hours at room temperature (25 to 28°C). 4.Examine the brown lesions under a dissecting microscope to observe the sporulating lesion. 5.Use a 9-inch Pasteur pipette with a very fine tip (tip melted by heating with a gas burner, pulled and cooled). Pick lightly the top of the conidial masses by touching it with the tip of the Pasteur pipette and then spread it gently on a water agar plate; distances between spores should allow you to pick up a single spore. Pick a single spore and transfer it into a marked spot on the water agar plate. Incubate the plates at 28°C for three to four days in order to see the growth of macroscopic mycelia (refer to Figure 24). Cut a fungal mycelium and transfer it to a rabbit food agar plate/slant. Incubate for five to seven days prior to stock culture for long-term storage. 6. 7. 8. 9. 10. Figure 24. Macroscopic mycelium 5-6 days after single spore isolation III. Preparation of stock culture for long-term storage (-20°C) 1.Place two pieces of pre-cut sterile Whatman no. 1 filter paper (equally cut into 4 pcs) on the surface of rabbit food agar plate. 2.Cut a fungal mycelium block 5x15 cm of a 6-7 d old culture of B. oryzae using a transfer needle/spatula. 3.Transfer it into a sterile test tube with 5 ml sterile distilled water. 4.Macerate the fungal mycelium block and pour about 1.5 ml in rabbit food agar lined with sterile filter paper. See to it that the filter paper is covered with the suspension then incubate for 3-4 days at 28 °C until the filter paper is covered by the growth of the fungal mycelium. 5.Remove or lift the colonized filter paper carefully and transfer it to a dry sterile plate lined with filter paper. Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 51 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 52 IV. Medium Preparation Yeast extract, 2 g Sucrose, 5 g Agar, 30 g Distilled water, 1000 ml Water Agar Stack the plates in a box and dried for two weeks under continuous light (fluorescent lamp) in a customized cabinet chamber. When the filter paper is completely dried, cut aseptically into small paper disks at 3-5 mm2. Transfer the small filter papers in sterilized 2 ml microcentrifuge tubes using alcohol-flamed sterilized forceps. Make at least two tubes for each isolate, one is for working (used for reviving) and the other one for storage only. Tubes must be labeled properly (isolate name and date of isolation). Keep the microcentrifuge tubes in a plastic DNA box and store at -20°C or -80°C freezer. 6. 7. 8. 9. 10. 1.Weigh and put all components and add into a two liter Erlenmeyer flask then add 1 liter of distilled water. 2.Autoclaved at 121°C for 20 minutes. 3.Allow to cool down the medium and dispense thinly (about 25ml in a plastic disposable plate) just enough to cover the surface. 4.Allow the plates to dry completely before using. Procedure Components and procedures were modified from Hayashi et al., (2009). Rabbit Food, 20 g Agar, 18 g Distilled water, 1000 ml Rabbit Food Agar Procedure: 1.Soak 20 g of rabbit food in 500 ml distilled water for five minutes. 2.Mix and stir the mixture and add 500 ml of distilled water. 3.Get the extract by filtering it with plastic nylon mesh. Volume it up to 1000 m and add 20g of agar. 4.Plug the flask with cotton and then autoclave at 121°C for 20 minutes at 15 psi. 5.Before plating add a pinch of Streptomycin for every liter of media then dispense the melted. media in a plate (approximately 20 plates/liter or about 50 ml/plate) and in a slant (9 ml). *Autoclaved at 121°C for 20 minutes. V. Plant Preparation and Maintenance 1.Soak seeds three days before sowing. Sow seeds in sterilized soil with basal fertilizer and grow for 18 days. If directly seeded, allow 21 days for growing seedlings (at 4-5 leaf stages). 2.Two days before inoculation (i.e. Day 17), apply nitrogen to promote vegetative growth. Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 53 Table 12. Sample inoculation schedule of Brown Spot Trial 1 Trial 2 Plant Preparation Soaking 17-Apr 04-May Seeding (2-3 days after soaking) 19-Apr 06-May Fertilizer Application (1-2 days before inoculation) 08-May 25-May Inoculation/Inoculum prep (18 days after seeding) 07-May 24-May Evaluation (6-7 days after inoculation) 13-May 30-May No. of Trays No. of isolates SM2 SM2 No. of plates/isolate 20 20 No. of flask to prepare (liters) 1 1 Inoculum needed (ml) 300 300 Culture Preparation Media Prep (4 days before culture prep) 15-Apr 02-May Pouring/Planting media 15-Apr 02-May Culture multiplication (subculture) (15 days before inoculation) 19-Apr 06-May Inoculum Preparation 07-May 24-May Revive isolates from filter (10 days before subculture) 09-Apr 26-Apr VII. Inoculation by Spraying Method 1.Spray the spore suspension using the airbrush attached to a compressor machine. Clean the airbrush using 70% Ethanol then rinse with sterile distilled water. 2.Make sure to cover the leaves evenly with a fine mist to mimic natural brown spot infection (200 plants will need about 30ml inoculum. 3.Place inoculated plants in a large plastic box and cover with the box lid. 4. Incubate the plants inside the moist jute sack chamber for 24 hours at ~25 °C. 5.After 24 hours, take out the plants and place them inside the mist room (~23-24°C) until disease assessment is covered with plastic film for five days. VI. Fungal Culture Preparation 1.Revive fungal isolate from filter paper stock for seven days in rabbit food Agar plates (This should be done concurrently with the seed soaking). The plated medium should be completely dry before use to avoid too much moisture during the incubation time. 2.After seven days, subculture the revived isolate into fresh rabbit food agar plates and allow fungal growth for 10 to 15 days under 12-hour dark and 12-hour near UV lighting. 3.Prepare the inoculum by dispensing about 15 ml of sterile distilled water with Tween 20 (500 ul Tween 20: 1L of sterile distilled water) and scrape the surface of the plates using a sterile glass slide. 4.Filter the spore suspension in a single layer of sterile miracloth. 5.Adjust the spore concentration to 15,000 spores/ml. Place the suspension in ice to prevent spore germination. VIII. Disease Assessment 1.Evaluate the plants at five days post-inoculation (dpi) following the IRRI SES for brown spots (refer to Table 13 and see Figure 25) and data sheet (see Table 14). Table 13. Rating Scale from IRRI SES with modifications Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 54 Figure 25. Diagrammatic rating scale guide by Lenz et al. (2010) with six levels of severity: 1.6, 3.2, 6.4, 12.6, 23.1, and 38.6% (top to bottom) Table 14. Sample data sheet for brown spot Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 55 Figure 26. Sample Leaf Sampling Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 56 Take a representative leaf sample from each line and mount it in a double- sided tape (see sample picture below). 2. 57 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases II. Measuring the total leaf area 1.Using the polygon tool, draw a selection around the leaf that you want to assess (see Figure 28). 2.Go to the “Edit” tab, press “Clear Outside”, then “SAVE” as a separate TIF file. Step-by-step protocol for quantification of diseased leaf areas using ImageJ software 1.Open ImageJ. 2.Open the desired image file (jpeg). 3.Zoom in on the ruler attached to the assessment sheet. 4.Use the line tool to measure 1 cm of the ruler (see Figure 27). I. Calibrating the Scale Figure 27. Setting the scale using the ruler attached to the assessment sheet Go to the “Analyze” tab and then press “Set Scale”. Change the known distance to “1” and the unit of length to centimeters. 5. 6. Figure 28. Leaf selected using the polygon tool (top) and saved as a TIF file (bottom). Open the TIF file. Go to “Image” tab then click on “Adjust” then select “Color Threshold”. Adjust the “Hue”, “Saturation”, and “Brightness” until the whole leaf is selected. Press “Select” to select the highlighted area (see Figure 29). Go to the “Analyze” tab then press “Measure” (or Ctrl + M). 3. 4. 5. 6. Figure 29. Appearance of the highlighted leaf in the software III. Measuring the diseased leaf area 1.Adjust the “Hue”, “Saturation”, and “Brightness” until the diseased areas are selected (see Figure 30). 2.Press “Select” to select the highlighted area. 3.Measure the diseased leaf area. Figure 30. Appearance of the highlighted brown spots in the software 58 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases IV. Computing for the %DLA V. Assigning index values 1.After computing the %DLA, assign the corresponding severity value as stipulated in the SES for BS disease. 2.You may also independently assign the severity value using the diagrammatic scale generated by Lenz et al. (2010). Step-by-step protocol for quantification of diseased leaf areas using ImageJ software and Ilastik software 1.Open ImageJ. 2.Open the desired image file (jpeg). 3.Zoom in on the ruler attached to the assessment sheet. 4.Use the line tool to measure 1 cm of the ruler (see Figure 27). I. Calibrating the Scale (as described above) II. Measuring the total leaf area (as described above) III. Measuring the diseased leaf area using Ilastik software Image Annotation in Ilastik 1.Open ilastik and select “Pixel Classification” under the “Create New Project” heading. 2. In the “Input data” tab, click Add New… � Add separate Image(s)… 3.Select images that will be used to train the software (i.e., representative brown- spot infected leaves) 4.Proceed to the “Feature Selection” tab and clock Select" “Features” (see Figure 31). 5.Put a check on the box where Color/Intensity and Sigma 0.70 intersect. Figure 31. Interface of the “Selected Features” 59 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases %DLA = x 100 ( )Diseased leaf area total leaf area Proceed to the “Training” tab, make the necessary labels, then annotate (see Figure 32). 6. Figure 32. An example of how the labels are used to make annotations Click “Live Update” then make more annotations until the desired outcome is achieved. Once done with the training, proceed to the “Prediction Export” tab and change the source to “Simple Segmentation” (see Figure 33). 7. 8. Figure 33. An example of how the segmentations should look after training Click “Choose Export Image Settings” � “Transformations” � tick “Convert to Data Type” � choose unsigned 8-bit from the dropdown menu � “Output File Info” � “Change Format” to TIFF. 9. 60 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases Batch processing 1.Proceed to the “Batch Processing” tab. 2.Press “Select Raw Data Files” then select the individual leaves TIF files from ImageJ. 3.Click “Process all Files”. Once done, files with “Segmentation” in their names will appear alongside the processed files. Put the segmentations in one folder. 4.Open ImageJ � File � Import � Image Sequence (see Figure 34). Figure 34. Import “Image Sequence” interface Click “Browse” then choose the folder with segmentations � “Change Type” to “RGB” � Check Sort names numerically and use virtual stack. Go to “Image” then click on “Adjust” then select “Color Threshold”. Set the values of the components similar to Figure 35. Press “Select’ then” Measure”. 5. 6. 7. 61 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases Figure 35. Threshold color values for the selection of the “brown spot” segmentation. Press the arrow in the image sequence to proceed to the next image (see Figure 36). Repeat Step 7 until finished. 8. Figure 36. Highlighted brown spot segmentation (left) and selected brown spot segmentation (right) 62 Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases Barnwal MK, Kotasthane A, Magculia N, Mukherjee, PK, Savary S, Sharma AK, Singh HB, Singh US, Sparks AH, Variar M, Zaidi N. 2013. A review on crop losses, epidemiology and disease management of rice brown spot to identify research priorities and knowledge gaps. Eur J Plant Pathol 136:443–457. DOI 10.1007/s10658-013-0195-6. Burgos MRG, Katimbang MLB, Dela Paz MAG, Beligan GA, Goodwin PH, Ona IP, Mauleon RP, Ardales EY, Vera Cruz CM. 2013. Genotypic variability and aggressiveness of Bipolaris oryzae in the Philippines. Eur J Plant Pathol. DOI 10.1007/s10658-013-0256-x. Dela Paz MG, Goodwin PH, Raymundo AK Ardales E, Vera Cruz CM. 2006. Phylogenetic analysis based on ITS sequences and conditions affecting the type of conidial germination of Bipolaris oryzae. Plant Pathology 55:756–765. Doi: 10.1111/j.1365-3059.2006.01439. Hayashi N, Kobayashi N, Vera Cruz CM, Fukuta Y. 2009. Protocols for the sampling of diseased specimens and evaluation of blast disease in rice. JIRCAS Working Report, 63:17–33. IRRI. 2013. Standard Evaluation for Rice. International Rice Research Institute. https://ricepedia.blogspot.com/2018/04/2013-irri-ses-standard-evaluation.html Lenz G, Balardin RS, Corte GD, Marques LN, Debonal D. 2010. Diagrammatic scale for assessment of rice brown spot severity. Ciencia Rural, 40(4), 752–758. https://doi.org/10.1590/ S0103-84782010005000061 Mew TW, Hibino H, Savary S, Vera Cruz CM, Opulencia R, Hettel GP, eds. 2017. Rice diseases: Biology and selected management practices. Los Baños (Philippines): International Rice Research Institute. PDF e-book. http://rice-diseases.irri.org . Raghavan C, Mauleon R, Lacorte V, , Zaw H, Bonifacio J, Singh RK, Huang BE, Leung H. 2017. Approaches in characterizing genetic structure and mapping in a rice multi-parental population. G3: Genes, Genomes, Genetics, 7(6):1721–1730. Schindelin J, Arganda-Carreras I, Frise E, Kaynig V, Longair M, Pietzsch T, Preibisch S, Rueden C, Saalfeld S, Schmid B, Tinevez J, White DJ, Hartenstein V, Eliceiri K, Tomancak P, Cardona A. 2012. Fiji: an open-source platform for biological-image analysis. Nat Methods 9:676–682. https://doi.org/10.1038/nmeth.2019 Schneider C, Rasband W Eliceiri K. 2012. NIH Image to ImageJ: 25 years of image analysis. Nat Methods 9:671–675. https://doi.org/10.1038/nmeth.2089 ilastik 0.5: Cell classification–2D workflow (https://www.youtube.com/watch?v=1EjGberIEZo ). Interactive image segmentation with ilastik (https://www.youtube.com/watch?v=F6KbJ487iiU ). Pixel classification (https://www.ilastik.org/documentation/pixelclassification/ pixelclassification). References We acknowledge previous members of the Plant Pathology and Host Plant Resistance Group for their contributions to the development and optimization of this brown spot protocol: Ruby Burgos, Ellen Silab, Casiana Vera Cruz, Vanica Lacorte- Apostol. Acknowledgements Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 63 https://ricepedia.blogspot.com/2018/04/2013-irri-ses-standard-evaluation.html http://rice-diseases.irri.org/ https://doi.org/10.1038/nmeth.2019 https://doi.org/10.1038/nmeth.2089 https://www.youtube.com/watch?v=1EjGberIEZo https://www.youtube.com/watch?v=F6KbJ487iiU Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 64 1.6 False smut (FS) False Smut (FS) causes chalkiness of grains which leads to reduction in grain weight. It also reduces seed germination. The disease can occur in areas with high relative humidity (>90%) and temperature ranging from 25−35 ºC. Rain, high humidity, and soils with high nitrogen content also favors disease development. Wind can spread the fungal spores from plant to plant. False smut is visible only after panicle exertion. It can infect the plant during flowering stage. The disease affects the early flowering stage of the rice crop when the ovary is destroyed. The second stage of infection occurs when the spikelet nearly reaches maturity. Plants infected with false smut have individual rice grain transformed into a mass of spore balls. These spore balls are initially orange and then turn into greenish black when these mature. In most cases, not all spikelets of a panicle are affected, but spikelets neighboring smut balls are often unfilled. The disease affects the early flowering stage of the rice crop when the ovary is destroyed. The second stage of infection occurs when the spikelet nearly reaches maturity. These causes chalkiness and can reduce 1,000-grain weight. It also causes a reduction in seed germination of up to 35%. In damp weather, the disease can be severe and losses can reach 25%. In India, a yield loss of 7−75% was observed. Authors: Mary Jeanie Telebanco-Yanoria, Casiana Vera Cruz, Van Schepler-Luu Screening Protocol Follow the sampling protocol described for bacterial blight and blast. Briefly: 1.Before carrying out the collection, make sure that you obtain official documents that allow collecting biological materials from farmers’ fields. 2. Identify an infected field, talk to farmers to collect the passport information (Table 1). 3.To determine the size where the survey will be conducted, use stratified sampling techniques and hierarchical scheme within the selected hotspot target areas. This includes four levels of sampling, to divide the target sites into logical categories (refer to Figure 4 and 5). a.Level 1 (Field): From an identified symptomatic field, determine the sampling units using the W pattern. The sampling unit is defined as a hill. Five hills are sampled from fields with an area of less than one hectare and eight hills are sampled from larger fields. b.Level 2 (Village): In cases where infections are spread in neighboring fields, identify 3-4 nearby fields that are 1 km apart from each other. Prioritize the diversity of rice varieties grown and select 3-4 different rice varieties. c.Level 3 (District): Search for other villages nearby that have infected fields. Follow the village collection scheme. d.Level 4 (Division): Search for 1-3 districts nearby that have infected fields. Follow the district collection scheme. 4.From each sampled hill, collect orange-colored smut balls from FS-infected rice plants by cutting the panicle 5.Place the samples from one hill in separate envelopes to avoid mixing them with samples from other hills. 6.Do not sample hills near the border of a field to avoid bias caused by interplot interference. This occurs when the pathogen or disease intensity is affected by adjacent plots or external factors. 7.Properly label each envelope with the sample ID (refer to Figure 6) 8.Place the coin envelopes inside a plastic bag and seal with tape. 9. Immediately place the plastic bags (with coin envelopes) inside a chest filled with ice while in the field. 10.In the lab, aerate or air-dry all samples collected for 4-8 hours and kept in -20°C. 11.Samples can be viable from 3 to 6 months depending on the stage of smut ball development. I. Sampling Module 1: Pathogen Collection, Isolation, Cultivation and Storage, Inoculation, and Evaluation for Bacterial and Fungal Diseases 65 1.Using sterile scissors, cut one smut ball from an infected panicle. 2.Place the smut ball into a sterile 2-ml Eppendorf tube using sterile forceps and label the tube properly (refer to Figure 37). This can be kept at -20 °C for 1 to 3 months prior to isolation. 3.To pick up the spores touch the smut ball very lightly using a 9-inch Pasteur pipette with a very fine tip (tip melted by heating with a gas burner/alcohol lamp, pulled and cooled). 4.Then touch it gently on the surface of the plated water agar medium. Usually, this area is marked with a triangle at the bottom of the plate. II. Single Spore Isolation 1. Place two pieces of pre-cut sterile Whatman no.1 filter paper (equally cut into four pieces) on the surface of the PSA plate (refer to Figure 37). 2. Cut a fungal mycelium block of 5 x 15 cm from a 10- to 15-day-old culture of U. virens using a transfer needle/spatula. 3. Transfer it into a sterile test tube with 5 ml of sterile distilled water. 4. Macerate the fungal mycelium block and pour about 1.5 ml into a PSA plate lined with sterile filter paper. See to it that the filter paper is covered with the suspension then incubate it for 5 to 7 days at 28 °C until the filter paper is covered by the growth of the fungal mycelium. 5. Remove or lift the colonized filter paper carefully and transfer it to a dry sterile plate lined with filter paper. 6. Stack the plates in a tray and dry/expose them for 2 weeks in continuous light (fluorescent lamp) installed in a cabinet or in a desiccator. 7. When the filter paper is completely dried, cut aseptically into small paper disks at 3 to 5 mm2. 8. Transfer the cut filter papers to sterilized 2-ml microcentrifuge tubes using alcohol-flamed sterilized forceps. Make at least two tubes for each isolate; one is for working stock (used for reviving) and the other one for storage only. 9. Tubes must be labeled properly (isolate name and date of storing). 10. Keep the microcentrifuge tubes in a plastic DNA box and store them in a freezer at -20 or -80 °C for up to 2 to 3 years. III. Preparation of stock culture for long-term storage (-20°C) Spread it gently on a water agar plate; distances between spores should allow for picking up a single spore under the dissecting microscope and spreading the spore mass on the marked area. Pick a single spore and transfer it onto the marked spot on the water agar plate and allow it to germinate for 36 hours or incubate the plates in a plastic box at room temperature (25 to 28 °C). Or allow the conidia to germinate for 36 hours and transfer one germinated spore by cutting the aga