Isolation, Identification, Storage, Pathogenicity Tests ... · 2O; 0.3 g strepto-mycin; 0.1 g...

PLANT HEALTH PROGRESS Vol. 16, No. 3, 2015 36

Diagnostic Guide

Isolation, Identification, Storage, Pathogenicity Tests, Hosts, and Geographic Range of Fusarium solani f. sp. pisi Causing

Fusarium Root Rot of Pea

Lyndon D. Porter, Grain Legume Genetics Physiology Research Unit, USDA-ARS, Prosser, WA 99350; Julie S. Pasche, Department of Plant Pathology, North Dakota State University, Fargo 58108; Weidong Chen, Grain Legume Genetics Physiology Research Unit, USDA-ARS, Pullman, WA 99164; and Robert M. Harveson, Panhandle Research and Extension Center, University of Nebraska, Scottsbluff 69361

Accepted for publication 8 September 2015. Published 15 September 2015.

Porter, L. D., Pasche, J. S., Chen, W., and Harveson, R. M. 2015. Isolation, identification, storage, pathogenicity tests, hosts, and geographic range of Fusarium solani f. sp. pisi causing Fusarium root rot of pea. Plant Health Progress doi:10.1094/PHP-DG-15-0013.

HOST: Pea (Pisum sativum L.)

DISEASE: Fusarium root rot

PATHOGEN Fusarium solani (Mart.) Sacc. f. sp. pisi (F. R. Jones) W. C.

Snyder & H. N. Hansen. Known by the previous name of Fusarium martii var. pisi F. R. Jones 1923 (Farr and Rossman 2015). The teleomorph is Haemanectria haematococca (Berkeley & Broome) Samuels & Nirenberg and Nectria haematococca Berk. and Br. is a common synonym for the sexual stage (Leslie and Summerell 2006).

TAXONOMY Kingdom Fungi; Phylum Ascomycota; Class Sordariomycetes;

Order Hypocreales; Family Nectriaceae; Genus Fusarium; Species solani. Special form: pisi. Current taxonomic identifi-cation is described at www.cabi.org/dfb and http://www.uniprot.org/taxonomy/?query=Fusarium+solani+f.+sp.+pisi&sort=score. Information regarding the general taxonomy of Fusarium solani can be found in several resources (Farr and Rossman 2015; Nelson et al. 1983; Toussoun and Nelson 1976).

SYMPTOMS AND SIGNS Above-ground symptoms of plants infected by F. solani f. sp. pisi include leaves that begin to turn yellow starting at the base of the plant and progressing to the top of the plant (Fig. 1). Below-ground stems of symptomatic plants contain black to brown lesions commonly found beginning at the point of seed attach-ment that can advance upwards to, or slightly above, the soil line and also spread downward to all parts of the root system (Figs. 2, 3, and 4). During early infection, small individual lesions can be seen on below-ground stem tissue (Fig. 5), but as the disease progresses, the lesions can coalesce into a single, continuous, black lesion that may reduce the number of lateral roots, depend-

ing on disease severity (Fig. 6). Below-ground stem lesions tend to be confined to the outer cortex and do not penetrate deep into the vascular tissue. This can be observed by scraping off the outer cortex to reveal healthy, white or light green vascular tissue under the black/brown lesion (Fig. 7). However, there can be severe below-ground stem rot, especially following flowering as the plant puts its resources into pod development and soil moisture becomes less available, as is observed in some dryland farming environments. F. solani f. sp. pisi can cause seedling damping off of highly susceptible pea cultivars (Fig. 8); however, damping off is not common in most cultivars. Wilting or death of emerged plants is not commonly associated with Fusarium root rot, but the presence of the disease can severely stunt the growth of infected plants and cause early maturation. Fusarium root rot often can be associated with a root rot complex that can include Aphanomyces euteiches Drechs. f. sp. pisi W. F. Pfender & D. J. Hagedorn causing Aphanomyces root rot; Fusarium oxysporum Schl. f. sp. pisi (J. C. Hall) W. C. Snyder & H. N. Hans. races 1 and 2 causing Fusarium wilt and near wilt, respectively; Thielaviopsis basicola (Berk & Broome) Ferraris causing black root rot; Rhizoctonia solani Kühn (AG-4) causing Rhizoctonia seedling rot; and Pythium spp. causing Pythium root rot and root tip necrosis (Davis and Shehata 1985). Nematodes, such as root lesion nematodes (Pratylenchus penetrans) (Riga et al. 2008), can also play a role in the root rot complex by causing root wounding which allows fungal pathogens to infect root systems (Fig. 9). Diagnosis of root rot pathogens based solely on symptom expression is difficult, especially when multiple pathogens are involved.

Soil compaction (Kraft and Boge 2001; Kraft and Giles 1979; Kraft and Wilkins 1989) and warmer (18 to 24°C) rather than cooler (13°C) soil temperatures (Kraft and Roberts 1969) favor the development of Fusarium root rot throughout the growing season. Currently, there are no effective seed or foliar fungicides that work to manage root infection by soilborne F. solani f. sp. pisi (Kraft and Papavizas 1983), and cultivars vary in levels of partial genetic resistance (Gretenkort and Helsper 1993;Grünwald et al. 2003; Hwang et al. 1995; Porter 2010). Pea cultivars with pigmented flowers tend to have greater partial resistance to F. solani f. sp. pisi than white-flowered cultivars (Grunwald et al. 2003; Kraft 1975). The pathogen can commonly infest the surface of seed via dust during harvest (Kraft 2001) and can be isolated

Corresponding author: Lyndon D. Porter. Email: [email protected]

doi:10.1094 / PHP-DG-15-0013 © 2015 The American Phytopathological Society

PLANT HEALTH PROGRESS Vol. 16, No. 3, 2015 Page 137

FIGURE 2 Below-ground stem and root lesions caused by Fusarium solani f. sp. pisi on the pea cultivar Dark Skin Perfection at a late vegetative growth stage.

FIGURE 1 Yellowing of foliar pea tissue beginning at the base of the plant and progressing upwards, caused by root lesions of Fusarium solani f. sp. pisi on the pea cultivar Genie at the first bloom growth stage.

FIGURE 3 Pea root tissue infected by Fusarium solani f. sp. pisi next to healthy tissue of an unknown cultivar at a late vegetative growth stage.

FIGURE 4 Infected pea seed and root tissue next to healthy tissue infected by Fusarium solani f. sp. pisi of the cultivar Dark Skin Perfection at a late vegetative growth stage.


FIGURE 5 Small to large black/brown lesions observed on white epicotyl tissue caused by Fusarium solani f. sp. pisi of an unknown pea cultivar at a late vegetative growth stage.

FIGURE 6 Root lesions of Fusarium solani f. sp. pisi reducing the number of lateral roots with increasing disease severity on the pea cultivar Dark Skin Perfection at a late vegetative growth stage.

FIGURE 7 Below-ground stem lesions caused by Fusarium solani f. sp. pisi on the pea cultivar Banner at an early pod fill reproductive growth stage. The infected cortex tissue on the far right and left was removed revealing healthy vascular tissue inside. The root in the center demonstrates the appearance of the infected outer cortex prior to removing the cortical tissue.

FIGURE 8 Pea plants infected by Fusarium solani f. sp. pisi causing seed rot, seedling damping off and stunting of multiple dry pea cultivars at pre-emergence, emergence and early vegetative growth stages.


from seed after surface sterilization (Begum et al. 2004), indicat-ing subsurface infection. Standard commercial seed treatments are effective in managing surface-infested or subsurface-infected seed (Begum et al. 2004) and can improve seed germination and early growth; however, seed treatments that move systemically to protect newly developing root tissue from F. solani f. sp. pisi are needed.

HOST RANGE F. solani f. sp. pisi is reported to infect the following hosts: alfalfa (Medicago sativa L.) (Smith et al. 2009); chickpea (Cicer arietinum L.) (Kraft 1969;VanEtten 1978; Westerlund et al. 1974); common bean (Phaseolus vulgaris L.) (Clarkson 1978; Miller et al. 1980; Smith et al. 2009); cottonwood (Populus deltoids Marsh.) (VanEtten 1978); gingseng (Panax gingseng C. A. Meyer) (VanEtten 1978); mulberry (Morus alba L.) (Matuo

and Snyder 1972; VanEtten 1978); lentil (Lens culinaris L.) (Lin and Cook 1977); red clover (Trifolium pretense L.) (Smith et al. 2009); sainfoin (Onobrychis viciifolia Scop.) (VanEtten 1978); and tuliptree (Liriodendron tulipifera L.) (VanEtten 1978). For more detail regarding the host range of F. solani f. sp. pisi, visit the Fungus-Host Distribution section of the Fungal Databases available at http://nt.ars-grin.gov/fungaldatabases/fungushost/fungushost.cfm (Farr and Rossman 2015).

GEOGRAPHIC DISTRIBUTION F. solani f. sp. pisi is cosmopolitan, found in pea-growing

regions worldwide, including Canada (Feng et al. 2010; McLaren et al. 2010), the Czech Republic (Ondřej et al. 2008), India (Hamid et al. 2012), southern Scandinavia (Persson et al. 1997), the United Kingdom (Etebu and Osborn 2010), New Zealand (Hawthorne et al. 1992), and the USA (Hampton and Ford 1965; VanEtten 1978).

PATHOGEN ISOLATION Isolates of F. solani f. sp. pisi can be obtained from surface-

sterilized, infected plant tissue or from infested soil using modified Nash-Snyder agar (MNSA) (Nash and Snyder 1962; Toussoun and Nelson 1976) (Fig. 10), a Fusarium-selective medium. A one-liter recipe of MNSA includes: 20.0 g Bacto-agar; 15.0 g peptone; 1.0 g KH2PO4; 0.5 g MgSO4•7H2O; 0.3 g strepto-mycin; 0.1 g neomycin; and 750 mg 1,2,3,4,5-pentachloro-6-nitrobenzene (PCNB). In addition, 0.1 g chlortetracycline hydro-chloride can be added to improve selectivity. The pH is adjusted to 5.5 to 6.5. The medium is autoclaved at 121°C for 20 min and streptomycin, chlortetracycline hydrochloride, neomycin, and PCNB are added to 5 ml of sdH2O and added to the medium when cooled to 50°C. Plates should be stored in the dark prior to use. For pathogen isolation, infected plant tissue is rinsed in

FIGURE 9 Pea roots of unknown cultivar infested and damaged (top photo) by the root lesion nematode (Pratylenchus penetrans, bottom photo) allowing other root rot fungi such as Fusarium solani f. sp. pisi to infect the root system at an early vegetative growth stage.

FIGURE 10 Colonies of Fusarium solani f. sp. pisi growing on modified Nash-Snyder agar, a selective medium for isolating Fusarium species. These colonies come from spores stored in sterile soil used to maintain spore viability for 20 or more years.


running tap water to remove any soil or debris and then soaked in a 10% sodium hypochlorite solution or 70% ethanol solution for 1.5 min, rinsed with sterile distilled water (sdH2O), and blotted dry with sterile paper. The tissue is placed on MNSA and incubated under 24-h, continuous, cool, white light at 23 to 25°C.

Further isolations are made from the fungal growth emanating from the tissue three to five days after plating. Spores obtained from the plates can be diluted with sdH2O and re-plated on MNSA to obtain single-spore isolates of the fungus.

PATHOGEN IDENTIFICATION F. solani f. sp. pisi is characterized by a cream-colored to white

colony with a yellow to tan center when grown on MNSA and on potato dextrose agar (PDA) (Fig. 11). The fungus produces large, canoe-shaped macroconidia, usually consisting of three septae,

FIGURE 11 Growth of Fusarium solani f. sp. pisi on modified Nash-Snyder agar (left) and potato dextrose agar (PDA) (right), in all pictures. The center of each culture on both media has a yellow to tan color surrounded by white to cream-colored mycelium (top photo). When observing the cultures through the bottom of the Petri plate, there are no real distinct color differences in the growth between the media (middle photo). However, as the cultures dry down on the media, the culture growing on PDA may develop a green color (bottom photo).

FIGURE 12 Macroconidia (canoe-shaped spores) with one to three septae and microconidia (smaller oval-shaped spores with zero to 1 septum) of Fusarium solani f. sp. pisi. Pictures taken using red and brown filters to enhance spore morphology.


and smaller, oval-shaped microconidia (Figs. 12 and 13) produced at the end of conidiophores (Fig. 14). Fusarium species are very difficult to identify based solely on morphology; therefore, molecular identification of Fusarium species can be found using Fusarium I.D. (Geiser et al. 2004) (http://isolate.fusariumdb.org). This database enables the identification of Fusarium species based on DNA sequence information. Comparisons between Fusarium species are primarily made based on sequences of the translation elongation factor 1-alpha, nuclear large subunit 28S rDNA, and the nuclear ribosomal internal transcribed spacer region (Geiser et al. 2004; O’Donnell 2000). In addition, quant-itative PCR using pathogenicity genes (PDA, PEP1, PEP3, and PEP5) have been used to determine variance in virulence and to identify and quantify the presence of F. solani f. sp. pisi in agricultural soils (Etebu and Osborn 2009, 2010). Also, isolates of certain fungal species can be divided into vegetative compatibility groups (VCG) based on the ability of different individuals within a population to form heterokaryons (Leslie 1993). VCG of F. solani populations causing dry rot on potato (Sharifi et al. 2008) and root rot on tobacco (Jusoh et al. 2013) have been character-

ized using nitrate non-utilizing mutants. This same technique could be used to evaluate VCG in F. solani f. sp. pisi, but this research has not been conducted.

PATHOGEN STORAGE Continuous transfer of F. solani f. sp. pisi cultures on PDA

maintained at room temperature is not recommended since cultures stored in this fashion are prone to mutations resulting in weak cultures with poor virulence or loss of pathogenicity (Toussoun and Nelson 1976). Half-strength PDA with strepto-mycin has been used to reduce the likelihood of these mutations happening during isolation and continuous transfers. Of the storage methods listed in this section, the sterile soil method is the authors’ preferred storage method, owing to its simplicity and successful means of storage of spores for extended periods of time while maintaining pathogenicity.

Sterile soil method. An isolate of F. solani f. sp. pisi is grown on MNSA under continuous, fluorescent, cool, white light for six days at 23 to 25°C. Using aseptic techniques under sterile conditions, six milliliters of sdH2O is added to the agar surface, and a sterile glass rod is used to make a slurry of the culture. The slurry is then strained through a single layer of cheesecloth to remove mycelia, and retain micro- and macroconidia and two milliliters is added to 10 grams of soil (3 parts silt loam soil, 1 part finely ground peat moss, and 1 part finely crushed perlite) contained in a sterile Petri dish. The peat moss is sifted through a 2,000-micron sieve, and the soil medium is autoclaved for 45 min on two consecutive days prior to use. The soil is thoroughly mixed to homogenize the consistency of the spores throughout the medium. The infested soil is allowed to dry overnight in a hood and placed in a sterile, plain-end glass test tube (16 mm × 150 mm) and capped with a disposable closure that allows for some gas exchange. Gas exchange is considered to promote longevity

FIGURE 13 Small oval-shaped clear microconidia of Fusarium solani f. sp. pisi produced from mycelium growing on the surface of modified Nash-Snyder agar (top and bottom photos). In the top photo, a conidiophore with branches containing microconidia can be seen growing from an infested grain of sand.

FIGURE 14 A single conidiophore of Fusarium solani f. sp. pisi. Conidia are not often observed attached to conidiophores when viewing specimens mounted on microscope slides. Picture taken using an orange filter to enhance morphology .


of spores but this has not been scientifically evaluated to deter-mine its benefit in increased survival time. Spores consisting of micro- and/or macroconidia are stored at 4°C. Isolates of F. solani f. sp. pisi have been successfully stored in this medium for more than 20 years. To recover spores from the soil, plate a small amount of soil on MNSA and incubate under the previously described light and temperature conditions for 5 to 7 days. If too much soil is placed on the agar, individual colonies are difficult to select.

Agar slant method (VanEtten 1978). Agar slants are made consisting of V8-juice agar (Toussoun and Nelson 1976) contained in small screw-cap test tubes (1 × 10 cm). A Fusarium isolate is transferred to the middle of the slant and maintained at 24 to 27°C under continuous, fluorescent, cool, white light for 1 to 2 weeks. Cultures are stored at 6°C and transferred every three months to maintain viability. This method has an increased risk of mutations compared to spore storage methods owing to the repeated transfer of mycelium.

Lyophilization method (Fisher et al. 1982). Carnation leaf agar (CLA) is made by taking carnation leaf segments measuring 5 × 5 mm that have been dried in an oven at 45 to 55°C for two hours. Subsequently, they are placed in an aluminum canister and sterilized using gamma irradiation. Several leaf segments are added to the surface of water agar containing 1.5 to 2% agar and autoclaved and cooled to 45°C. An isolate of F. solani f. sp. pisi is added to the cooled and solidified medium and allowed to grow for 7 to 10 days. Several colonized carnation leaves are added to 5 ml vials, followed by 0.5 ml of sterile skim milk. Vials are loosely capped with a split rubber stopper to allow for air evacuation, placed on a tray, and quick-frozen with liquid nitrogen. An acrylic plate slightly larger than the tray is placed on top of the vials. A drying chamber (Model 10-MR-SA, The VirTis Co., Gardiner, NY) maintained at –35°C in a refrigerated freeze-dryer is used to lyophilize the tissue. Following 10 min of refrigeration, the vacuum pressure is released and maintained at 10 μm Hg on a McLeod gauge. The chamber is heated to 15°C and tissue samples allowed to dry for 16 to 20 h. The vials are sealed under vacuum pressure by inflating a rubber diaphragm in the chamber over the tray, forcing the rubber stoppers in the vials to seal. This storage method has resulted in 100% preservation of viable cultures after a 4-year period.

Freezing method (Leslie and Summerell 2006). Cultures are grown on agar slants in 10 × 75 mm test tubes containing 1.25 ml of PDA. A spore suspension is made with sterile 15% glycerol (15:85 glycerol:distilled water). A Pasteur pipette, with a full pipette barrel is used to provide approximately 2 ml of the 15% glycerol solution to the surface of the culture, and the agar surface is agitated with the pipette tip to free spores and mycelium. The suspension is removed from the tube and transferred immediately to a 2-ml cryovial. The cryovial may be left on the bench for 30 min to an hour, if necessary, before being placed into a box in the ultra-low temperature freezer at –70°C. If the vial sits for more than an hour, it should be inverted several times or vortexed before freezing to insure the spores and hyphal fragments have not settled to the bottom. Cultures are recovered by scraping some of the ice from the frozen suspension. Cultures that have thawed completely should be remade, but if no significant thawing has occurred, the contents of the vial may be refrozen and reused. Cultures stored by freezing remain viable for a minimum of two years as long as the contents remain sterile.

Silica gel method (Windels et al. 1988). Cultures are grown on PDA or CLA under fluorescent light (four General Electric or Sylvania 40-watt tubes), supplemented with black light (one Sylvania 40-watt tube, BLB series) for a 12-h photoperiod.

Suspensions of conidia are made from actively sporulating cultures by scraping the colony surface and placing conidia into screw-cap culture tubes (100 × 13 mm) containing 2 ml of sterile skim milk (previously autoclaved for 15 min). When cultures are grown on CLA, two or three leaves containing spores are trans-ferred to tubes of skim milk. Screw-cap culture tubes (100 × 13 mm) are filled with 3 cm3 silica gel (specification Mil-D-3716 or Davison Chemical Corporation, commercial grade H and 05, mesh 6-16, code 05-08-08-237). Silica gel (non-indicating, no dye) is dry-heat sterilized at 180°C for 1.5 h and stored at 5°C until used. Tubes are placed in an ice bath for at least 30 min before use. A pre-cooled tube is held horizontally to distribute the particles along the side of the tube and 0.3 ml of the conidial suspension is evenly and aseptically applied to the silica gel. The suspension is added in ample volume to moisten the particles, if too much moisture is added, the silica gel will fuse. The contents of the culture tube are immediately mixed by vortexing to enhance the distribution of conidia on the silica gel, and the tube is placed in an ice bath until cool (some heat is released when moisture contacts silica gel). Before storage, each isolate is checked for viability by shaking a few particles onto PDA. A single layer of Parafilm is stretched around the screw cap and the culture is stored at 4.5 ± 0.5°C.

Cellulose filter paper method (Egel and Hoke 2009). Isolates to be stored are grown on PDA or CLA under continuous, fluorescent, cool, white lights for six days. Whatman #1 filter papers discs (GE Healthcare, Little Chalfont, UK) are placed in a glass Petri plate and autoclaved for 15 min at 121°C. Discs are aseptically placed on the surface of ¼-strength PDA media in Petri plates (9-cm diameter) and incubated at room temperature (25°C) under a 12-h photoperiod of fluorescent, cool, white light. F. solani f. sp. pisi isolates are transferred to the medium and allowed to grow for 7 to 10 days until the mycelial growth completely covers the filter paper discs. The filter paper with the fungal growth is removed from the agar, and placed on a sterile surface and allowed to dry for one day. Once dry, it is cut into small pieces (5-mm diameter) using aseptic techniques under sterile conditions and placed in cryovials and stored at 5°C until used.

GREENHOUSE/GROWTH CHAMBER PATHOGENICITY TESTS

Of the pathogenicity tests described in this section, the authors prefer the seed-soak method, owing to its simplicity in set up, consistent results, and based on unpublished data indicating this method correlates well with field screening results from Fusarium root rot disease nurseries.

Seed soak method. This method was modified from Kraft and Kaiser (1993). An isolate of F. solani f. sp. pisi is grown on MNSA in a Petri dish under continuous, fluorescent, cool, white light for six days, after which three agar plugs (4 mm in size) containing mycelium of F. solani f. sp. pisi are transferred from the leading edge of a colony to a flask containing Kerr’s medium (Kerr 1963). A one-liter recipe of Kerr’s medium includes: 2.0 g NaNO3; 1.0 g KH2PO4; 0.5 g KCl; 0.5 g MgSO4•7H2O; 0.01 g FeSO4•7H2O; 0.5 g yeast extract; 30.0 g sucrose; and 1,000 ml distilled water. The medium is autoclaved for 20 min and cooled to room temperature prior to adding the agar plugs. The culture is incubated on a shaker at 120 rpm under continuous, fluorescent, cool, white light for six days at 23 to 25°C, and strained through a single layer of cheesecloth to remove mycelia. The medium containing the spores is centrifuged at 2,500 rpm (rotor diameter 30 cm) for 5 min. The Kerr’s medium is removed and spores are re-suspended in sdH2O. This step can be repeated to ensure spores


are washed free of medium. Spore concentrations are determined using a hemocytometer and diluted to 1 × 106 spores/ml water. All seeds are surface sterilized in 10% sodium hypochlorite, rinsed in sdH2O, and allowed to dry at 25°C. Seed is soaked for 16 h in the spore suspension or with sdH2O for non-inoculated controls. Seeds of each genotype are placed in 100-ml beakers and a standard volume of inoculum is stirred to form a homogenous mixture and poured over the seed in ample volume to submerge. Seeds are planted individually in pasteurized, coarse-grade perlite in plastic planter cones (0.25-liter Conetainer, Stuewe and Sons Inc., Tangent, OR) beginning with the non-inoculated controls to avoid contamination. It is best to keep the perlite wet to avoid dust issues and to facilitate filling cones. The cones are arranged in a split-plot design, the main plot being the inoculation factor and the genotypes being the subplots. Seeds are planted at a depth of 2.5 cm. Water should not be applied until two to three hours after planting, after which plants are irrigated as needed, generally every 24 to 36 h, and the perlite is watered to saturation. Place inoculated and non-inoculated plants at a sufficient distance from each other to avoid splash contamination, and always water the non-inoculated controls first. Each genotype should have inocu-lated and non-inoculated controls to assess the levels of natural contamination or infestation on or within the seed being eval-uated. A 14-h photoperiod is maintained using 400-watt metal halide lamps as needed for supplemental lighting.

Quantitative evaluations such as root disease severity (RDS), plant height, and dry root and shoot weights are recorded 21 days after planting. RDS is accessed using a visual scale from 0 to 6 adapted from Grünwald et al. (2003), where: 0 = no diseases symptoms; 1 = slight hypocotyl lesions; 2 = lesions coalescing around epicotyls and hypocotyls; 3 = lesions starting to spread into the root system with root tips starting to be infected; 4 = epicotyl, hypocotyl, and root system almost completely infected and only a slight amount of white, uninfected tissue visible; 5 = completely infected root; and 6 = plant failed to emerge.

Modified sand cornmeal layering method. This method was modified from Bilgi et al. (2008). An isolate of F. solani f. sp. pisi is placed on PDA and incubated at room temperature (23 to 25°C) under continuous, fluorescent, cool, white light with alternating 12-h light and dark cycles for 5 days. Sand-cornmeal inoculum is prepared at a 9:1:2 ratio by weight of sand, cornmeal, and sdH2O. The mixture is stirred in a flask, covered with foil, autoclaved for 45 min, and cooled to room temperature. Eight agar plugs (5 mm), colonized with mycelia of F. solani f. sp. pisi, are added to the mixture and the flask is covered again with foil. The flask is swirled by hand at least once daily and maintained at room temperature for up to 10 days. The mixture is stirred with a sterile spatula to establish a uniform consistency prior to layering. Plastic drinking cups (266 ml) with three to six equally spaced, small drainage holes in the bottom (created using a hot needle) are filled with 15 g of pasteurized coarse-grade vermiculite. Fifteen grams of sand-cornmeal inoculum is placed over the vermiculite followed by 8 g of additional vermiculite. One pea seed, surface sterilized using the protocol described in the seed-soak method, is placed on this layer. Seed is covered with 8 g of additional vermiculite. Water (80 ml) is added slowly to each cup to allow the vermiculite to absorb it without runoff. Cups are placed on plastic trays with openings to allow for water drainage and placed in a growth chamber or greenhouse with a 14-h light cycle at 21°C and a 10-h dark cycle at 18°C for 10 days. Plants are watered daily until vermiculite is saturated. Plants are removed from cups and roots rinsed in water to remove excess vermiculite. Root rot severity is rated as previously described.

The non-inoculated controls are established following the same protocol but using non-colonized agar plugs.

Pipette method. This method was modified from Lockwood (1962). The same protocol previously described in the seed-soak method is used to surface sterilize seed, produce inoculum of F. solani f. sp. pisi at a concentration of 1 × 106, and rate plants for disease severity 21 days after planting. Pea seed is sown to a depth of 2.5 cm in either pasteurized sand or coarse-grade perlite wetted with a standardized amount of water prior to use to facilitate handling and dust issues. The medium should not be dry but should not be saturated, to allow inoculum to move through the soil profile. Inoculations can be conducted using one of three methods: seeds are inoculated by applying 1 ml of inoculum directly over the top of each seed at planting; higher levels of inoculum (10 to 50 ml) also can be applied to the soil surface and inoculum watered down to the seed; or plants can also be inocu-lated with 1 to 2 ml of inoculum placed at the base of emerged plants 10 days after planting. Plants are watered daily to prevent plant medium from drying.

Test tube method. This method was derived from Dyer and Ingram (1990). The same protocol previously described in the seed soak method is used to surface sterilize seed. Seeds are placed in the dark on sterile, moistened germination paper and sealed in Petri dishes using Parafilm to maintain 100% relative humidity. Germinating peas are individually placed into polyester plugs consisting of cylinders of polypropylene fibers measuring 2.5 × 3.0 cm. Plugs were previously autoclaved in half-strength Hoagland solution #1 (Table 1) (Hoagland and Arnon 1950). The plug containing a pea is placed in a sterile test tube (internal diameter 20 mm), filled with 10 ml of half-strength Hoagland solution #1 so the plug is suspended 2 cm above the solution, and the test tube is sealed with a cap. Peas are incubated at 25°C with a 16-h photoperiod (80 µmol/m2s) until the roots are 3 cm long. Peas are removed from the test tubes, and the 10 ml of half-strength Hoagland solution #1 is replaced with 10 ml of inoculum at 1 × 106 spores/ml water. Spores were produced from 10 to 14

TABLE 1 Contents of one liter of Hoagland Solution #1

Product Amount H2O 993.5 ml

1M KH2PO4a 0.5 ml

1M KNO3 2.5 ml

1M Ca(NO3)2•4H2O 2.5 ml

1M MgSO4•7H2O 1.0 ml

H3BO3b 2.86 g

MnCl2•4H2Ob 1.81 g

ZnSO4•7H2Ob 0.22 g

CuSO4•5H2Ob 0.08 g

H2MoO4•H2Ob 0.02 g

0.5% Iron tartrate solution 1.0 ml

0.1 NH2SO4b variable

a 1M = signifies a one molar solution of the compound.

b Products with a “b” superscript are dissolved together in 1 liter of water, and one milliliter of the solution is added to one liter of Hoagland Solution #1.

c Titration is used with 0.1 N H2SO4 until solution reaches a pH of 6.


day-old cultures of F. solani f. sp. pisi growing in 0.1% water agar. All spores were filtered through one layer of cheesecloth, centrifuged at 2,500 rpm (rotor diameter 30 cm) for 5 min, the water agar decanted, and the spores re-suspended in sdH2O. The solution is centrifuged again, and re-suspended in 0.1% water agar, which maintains the spores in suspension. Plants are re-inserted into the test tubes so the ends of the roots are immersed in the spore suspension, leaving a 2-cm air gap between the polyester plug and the agar layer. After 14 days, plants are removed from the tubes and plugs, and disease assessments are made using a key modified from Clarkson (1978): 0 = healthy, no symptoms; 1 = slight discoloration of root tips; 2 = root tips darkened, small brown fleck-like lesions on root; 3 = coalescence of flecks producing large lesions; 4 = general discoloration of root, not girdling root; 5 = deep discoloration of root, general foot rot girdling root; 6 = dead, complete collapse of tissues, plant wilted. See Dyer and Ingram (1990) for a picture of the test tube method.

FIELD PATHOGENICITY TESTS Spore inoculation method (Kraft and Berry 1972). An isolate

of F. solani f. sp. pisi is grown on MNSA for six days, after which three agar plugs containing the isolate are transferred to Kerr’s medium and incubated on a shaker rotating at 120 rpm under continuous, fluorescent, cool, white light for six days at 23 to 25°C. The inoculum is incubated for 1 week at 24°C, and the conidial suspension strained through a double layer of cheese-cloth and stored in a cold room at 5°C until further use.

The conidial suspension is diluted with water to a concentration of 6 × 106 to 8 × 106 spores/ml water as determined by a hemo-cytometer. The spore suspension is uniformly sprayed on the soil surface at a rate of 268 ml/m2 of soil using a backpack sprayer. The inoculum is incorporated to a depth of 15 cm with a rototiller. Non-infested control plots are roto-tilled first. Peas are planted in the inoculated and non-inoculated soil. Random soil samples are taken from artificially inoculated soil and non-inoculated soils prior to infesting the soil and one week after infestation to determine populations of F. solani f. sp. pisi. Soil samples consist of three homogenously mixed soil cores (2.5 × 15.2 cm). Rhizo-sphere soil samples can also be collected 14 days after plant emergence and at full bloom (Kraft et al. 1969). All soil samples are air dried and assayed by serial dilution onto plates of MNSA. Disease estimates are made on plants dug at full bloom using a 0-to-6 disease severity rating scale as previously described (Grünwald et al. 2003).

Infested grain inoculation method. Liquid cultures of F. solani f. sp. pisi are generated as previously described for the spore inoculation method. Grain is placed into an aluminum or tin tray and soaked in distilled water until completely imbibed. Caution should be taken to avoid prolonged soaking times and temperatures that facilitate fermentation of seed. Excess distilled water is decanted and the grain is autoclaved for 1 h on two consecutive days. The fungal solution is poured over the imbibed, sterilized, and cooled grain under aseptic conditions and grown at room temperature (25ºC) for 14 to 21 days until the grain is completely colonized by the fungus. The grain is spread onto a flat surface, dried for 1 to 2 weeks, and held in cool, dry condi-tions until planting. Approximately 1 g of infested grain per 30.5 cm of row is placed into the furrow prior to, or at, planting with the pea seeds. Disease severity is assessed as described above for the spore inoculation method.

ACKNOWLEDGMENTS Funding for this project was provided by the National Institute of Food

and Agriculture, U.S. Department of Agriculture under Agreement No. 2012-51120-20252 by the North Central IPM Center Working Group Grants Program.

LITERATURE CITED

Begum, N., Alvi, K. Z., Haque, M. I., Raja, M. U., and Chohan, S. 2004. Evaluation of mycoflora associated with pea seeds and some control measures. Plant Pathol. J. 48:48-51.

Bilgi, V. N., Bradley, C. A., Khot, S. D., Grafton, K. F., and Rasmussen, J. B. 2008. Response of dry bean genotypes to Fusarium root rot, caused by Fusarium solani f. sp. phaseoli, under field and controlled conditions. Plant Dis. 92:1197-1200.

Clarkson, J. D. S. 1978. Pathogenicity of Fusarium spp. associated with foot-rots of peas and beans. Plant Pathol. 27:110-117.

Davis, D. W., and Shehata, M. A. 1985. Breeding for resistance to root-rot pathogens of peas. Pages 237-246 in: The Pea Crop: A basis for improvement. P. D. Hebblethwaite, M. C. Heath, and T. C. K. Dawkins, eds. Mid-County Press, London.

Dyer, P. S., and Ingram, D. S. 1990. An improved test for evaluating the pathogenicity of isolates of Fusarium solani f. sp. pisi on pea. Ann. Appl. Biol. 117:469-472.

Egel, D., and Hoke, S. 2009. Long term refrigerator storage of Didymella bryoniae and other fungi on filter paper. Nat. Plant Diag. Net. 4:5-6.

Etebu, E., and Osborn, A. M. 2009. Molecular assays reveal the presence and diversity of genes encoding pea footrot pathogenicity determinants in Nectria haemotococca and in agricultural soils. J. Appl. Microbiol. 106:1629-1639.

Etebu, E., and Osborn, A. M. 2010. Molecular quantification of the pea footrot disease pathogen (Nectria haematococca) in agricultural soils. Phytoparasitica 38:447-454.

Farr, D. F., and Rossman, A. Y. Fungal Databases, Systematic Mycology and Microbiology Laboratory, USDA-ARS. http://nt.ars-grin.gov/fungaldatabases/

Feng, J., Hwang, R., Chang, K. F., Hwang, S. F., Strelkov, S. E., Gossen, B. D., Conner, R. L., and Turnbull, G. D. 2010. Genetic variation in Fusarium avenaceum causing root rot on field pea. Plant Pathol. 59:845-852.

Fisher, N. L., Burgess, L. W., Toussoun, T. A., and Nelson, P. E. 1982. Carnation leaves as a substrate and for preserving cultures of Fusarium species. Phytopathology 72:151-153.

Geiser, D. A., Jimenez-Gasco, M. M., Kang, S., Makalowska, I., Veeraraghavan, N., Ward, T. J., Zhang, N., Kuldau, G. A., and O’Donnell, K. 2004. Fusarium-ID v. 1.0: A DNA sequence database for identifying Fusarium. Eur. J. Plant Pathol. 110:473-479.

Gretenkort, M. A., and Helsper, J. P. F. G. 1993. Disease assessment of pea lines with resistance to foot rot pathogens: Protocols for in vitro selection. Plant Pathol. 42:676-685.

Grünwald, N. J., Coffman, V. A., and Kraft, J. M. 2003. Sources of partial resistance to Fusarium root rot in the Pisum core collection. Plant Dis. 87:1197-1200.

Hamid, A., Bhat, N. A., Sofi, T. A., Bhat, K. A., and Asif, M. 2012. Management of root rot of pea (Pisum sativum L.) through bioagents. Afr. J. Microbiol. Res. 6:7156-7161.

Hampton, R. O., and Ford, R. E. 1965. Pea diseases in Washington and Oregon. Plant Dis. Rep. 49:235-238.

Hawthorne, B. T., Rees-George, J., and Broadhurst, P. G. 1992. Mating behavior and pathogenicity of New Zealand isolates of Nectria haematococca (Fusarium solani). N. Z. J. Crop Hort. Sci. 20:51-57.

Hoagland, D. R., and Arnon, D. I. 1950. The water-culture method of growing plants without soil. Agric. Exp. Stn. Circ. 347. Univ. of Calif., Berkeley.

Hwang, S. F., Howard, R. J., Chang, K. F., Park, B., Lopetinsky, K., and McAndrew, D.W. 1995. Screening of field pea cultivars for resistance to Fusarium root rot under field conditions in Alberta. Can. Plant Dis. Surv. 75:51-56.

Jusoh, M. N. b., Zin, N. b. M., and Nagao, H. 2013. Vegetative compatibility group of Fusarium solani pathogenic to tobacco plant in peninsular Malaysia. Songklanakarin J. Sci. Technol. 35:615-662.

Kerr, A. 1963. The root rot-Fusarium wilt complex of peas. Aust. J. Biol. Sci. 16:55-69.

Kraft, J. M. 1969. Chickpea, a new host of Fusarium solani f. sp. pisi. Plant Dis. Rep. 53:110-11.


Kraft, J. M. 1975. A rapid technique for evaluating pea lines for resistance to Fusarium root rot. Plant Dis. Rep. 59:1007-1011.

Kraft, J. M. 2001. Fusarium root rot. Pages 13-14 in: Compendium of Pea Diseases and Pests, 2nd Ed. J. M. Kraft and F. L. Pfleger, eds. APS Press, St. Paul, MN.

Kraft, J. M., and Berry, J. W., Jr. 1972. Artificial infestation of large field plots with Fusarium solani f. sp. pisi. Plant Dis. Rep. 56:398-400.

Kraft, J. M., and Boge, W. 2001. Root characteristics in pea in relation to compaction and Fusarium root rot. Plant Dis. 85:936-940.

Kraft, J. M., and Giles, R. A. 1979. Increasing green pea yields with root rot resistance and subsoiling. Pages 407-413 in: Soil-Borne Plant Pathogens. B. Schippers, ed. Academic Press, New York.

Kraft, J. M., and Kaiser, W. J. 1993. Screening for disease resistance in pea. Pages 123-144 in: Breeding for Stress Tolerance in Cool-Season Food Legumes. K. B. Singh and M. C. Saxena, eds. John Wiley and Sons, New York.

Kraft, J. M., and Papavizas, G. C. 1983. Use of host resistance, Trichoderma, and fungicides to control soilborne diseases and increase seed yields of peas. Plant Dis. 67:1234-1237.

Kraft, J. M., and Roberts, D. D. 1969. Influence of soil water and temperature on the pea root rot complex caused by Pythium ultimum and Fusarium solani f. sp. pisi. Phytopathology 59:149-152.

Kraft, J. M., and Wilkins, D. E. 1989. The effects of pathogen numbers and tillage on root disease severity, root length, and seed yields in green peas. Plant Dis. 73:884-887.

Kraft, J. M., Haglund, W. A., and Reiling, T. P. 1969. Effect of soil fumigants on control of pea root rot pathogens. Plant Dis. Rep. 53:776-780.

Leslie, J. F. 1993. Fungal vegetative compatibility. Annu. Rev. Phytopathol. 31:127-150.

Leslie, J. F., and Summerell, B. A. 2006. Fusarium solani (Martius) Appel & Wollenweber emend. Snyder & Hansen. Pages 250-254 in: The Fusarium Laboratory Manual. Blackwell Publishing Ltd, Oxford, UK.

Lin, Y., and Cook, R. J. 1977. Root rot of lentils caused by Fusarium roseum ‘Avenaceum’. Plant Dis. Rep. 61:752-755.

Lockwood, J. L. 1962. A seedling test for evaluating resistance of pea to Fusarium root rot. Phytopathology 52:557-559.

Matuo, T., and Synder W. C. 1972. Host virulence and the Hypomyces stage of Fusarium solani f. sp. pisi. Phytopathology 62:731-735.

McLaren, D. L., Conner, R. L., Hausermann, D. J., Henderson, T. L., Penner, W. C., and Kerley, T. J. 2010. Field pea diseases in Manitoba in 2009. Can. Plant Dis. Surv. 90:148-149.

Miller, D. E., Burke, D. W., and Kraft, J. M. 1980. Predisposition of bean roots to attack by the pea pathogen, Fusarium solani f. sp. pisi, due to temporary oxygen stress. Phytopathology 70:1221-1224.

Nash, S. M., and Snyder, W. C. 1962. Quantitative estimations by plate counts of propagules of the bean root rot Fusarium in field soils. Phytopathology 52:567-572.

Nelson, P. E., Toussoun, T. A., and Marasas, W. F. O. 1983. Descriptions and illustrations of well-documented Fusarium species, F. solani. Pages 146-150 in: Fusarium Species: An Illustrated Manual for Identification. Pennsylvania State Univ. Press, University Park, PA.

O’Donnell, K. 2000. Molecular phylogeny of the Nectria haematococca-Fusarium solani species complex. Mycologia 92:919-938.

Ondřej, M., Dostálová, R., and Trojan, R. 2008. Evaluation of virulence of Fusarium solani isolates on pea. Plant Protect. Sci. 44:9-18.

Persson, L., Bødker, L., and Larsson-Wikström, M. 1997. Prevalence and pathogenicity of foot and root rot pathogens of pea in southern Scandinavia. Plant Dis. 81:171-174.

Porter, L. D. 2010. Identification of tolerance to Fusarium root rot in wild pea germplasm with high levels of partial resistance. Pisum Genet. 42:1-6.

Riga, E., Porter, L. D., Mojtahedi, H., and Brocke, G. F. 2008. Pratylenchus neglectus, P. thornei, and Paratylenchus hamatus nematodes causing yield reduction to dryland peas and lentils in Idaho. Plant Dis. 92:979.

Sharifi, K., Zare, R., and Rees-George, J. 2008. Vegetative compatibility groups among Fusarium solani isolates causing potato dry rot. J. Biol. Sci. 8:374-379.

Smith, I. M., Dunez, J., Phillips, D. H., Lelliott, R. A., and Archer, S. A. 2009. European Handbook of Plant Diseases. Blackwell Scientific Publ., Oxford, London. doi: 10.1002/9781444314199

Toussoun, T. A., and Nelson, P. E. 1976. A Pictorial Guide to Identification of Fusarium Species. 2nd Ed. Pages 7-25. Pennsylvania State Univ. Press, University Park, PA.

VanEtten, H. D. 1978. Identification of additional habitats of Nectria haematococca mapping population VI. Phytopathology 68:1552-1556.

Westerlund, F. V., Jr., Campbell, R. N., and Kimble, K. A. 1974. Fungal root rots and wilts of chickpea in California. Phytopathology 64:432-436.

Windels, C. E., Burnes, P. M., and Kommedahl, T. 1988. Five-year preservation of Fusarium species on silica gel and soil. Phytopathology 78:107-109.

Isolation, Identification, Storage, Pathogenicity Tests ... · 2O; 0.3 g strepto-mycin; 0.1 g...

Documents

Transcript of Isolation, Identification, Storage, Pathogenicity Tests ... · 2O; 0.3 g strepto-mycin; 0.1 g...