Hosts: Wheat (Triticum aestivum), Barley (Hordeum vulgare), Oats (Avena sativa)
Pathogen: Gaeumannomyces graminis var. tritici (Ggt) causes
disease in wheat and barley, G. graminis var. avenae causes
disease in oats, and G. graminis var. graminis causes disease in
Symptoms and Signs
The first symptoms of take-all on wheat or barley occur on young seedlings (Fig. 1). Infected root and stem tissue darkens to a nearly black color and the lower leaves typically are chlorotic. If plants are not killed at this stage of growth, they tiller poorly or not at all, and black lesions develop on roots and extend up into the crown tissue as the disease progresses (Fig. 2). Dark runner hyphae are evident growing along root tissue (Fig. 3). Where soils remain ideally moist for this disease, e.g., in high rainfall areas or in irrigated fields, the disease may develop in patches, and eventually plants in patches (Fig. 4) develop white heads and die prematurely (Fig. 5). Plants at this stage of disease development pull easily from the soil because the roots are severely rotted (Figs. 6, 7) and usually have quite black crown tissue. If the wet conditions prevail beyond the period of crop maturity, perithecia may be produced at the base of infected culms (Figs. 8, 9).
Where soils are moist during stand establishment and tillering but then
become dry while the plants remain alive on deeper soil moisture, disease
symptoms typically do not progress up the culms nor does the disease show up as
patches, but infected plants may still produce white heads. The infected crowns
of plants with take-all under dry conditions are evident as grey to black tissue
where the tillers join the main stem, a symptom revealed only by cutting the
crown open with a sharp knife.
Ggt attacks species of Bromus, Hordeum, and Triticum
severely, and species of Dactylis, Festuca, Lolium, and Poa
moderately (18). As cited herein, specific isolates of Ggt can also attack Avena
sativa in Australia (27). Thus, the host range of Ggt is wide within members
of the Poaceae. Nilsson (17) lists over 402 grass species that are hosts to Ggt,
G. graminis var. avenae or G.graminis var. graminis.
Ggt is found throughout the world where wheat is grown under temperate
climates and also occurs in the tropics at high elevation (6). Recently,
take-all has been found in arid wheat producing areas of the world where
irrigation has been used, including Montana, Texas, North Dakota, Idaho,
Washington, and Oregon in the USA, and in Israel (1). Most likely these
outbreaks are examples of "dryland" take-all developing into the more
classical "wet land" take-all. This is indicated by i) the classical
version of the disease often appearing in the 2ndor 3rd
irrigated crop, and ii) it occurs uniformly within the field.
Isolation from host tissue. Because Ggt is normally a slow-growing fungus, isolation of Ggt from host tissue can be frustrating and difficult if proper precautions and techniques are not used. It is best to first wash the tissue well in running water for an hour or more. Then it should be surface disinfested in 1% silver nitrate for 30 sec, followed by 2-3 rinses in sterile distilled water (11). Treatment with NaOCl may be inadequate for eliminating many of the contaminants on root tissue. After blotting the excess water using sterile paper towels, small pieces of infected tissue should then be placed on the Ggt semi-selective medium (SM-GGT3) (11) or as recently modified (5). This medium contains PDA amended with L-DOPA, which turns black in close proximity to growing hyphae of Ggt (Fig. 10), plus antibiotics and antifungal agents to restrict growth of competing organisms. In my laboratory, SM-GGT3 has worked well except in the rare cases where dichloran-resistant strains of Rhizopus spp. are encountered. Although isolations normally are made from tissue that has been infected for only 2-3 wk, pure cultures can be obtained using SM-GGT3 from mature infected host tissues.
Isolations can also be made by spreading the mass of extruded asci and ascospores, which appears at the tip of the perithecium, over the surface of PDA containing streptomycin and vancomycin (3, 12). Ascospore cultures should be incubated at 20-25°C.
Before selective media were developed, baiting was a common isolation technique, and can still be used effectively. This technique is accomplished by embedding the suspect tissue in sterile vermiculite in close proximity to a germinating wheat seed. Seedlings are allowed to develop in the light for 2-3 wk at 15-20°C, after which isolations are made from the newly infected roots of the developing seedlings. Standard surface-disinfestation of 2-3 min in 0.5 % NaOCl is followed by placing tissue on a medium such as PDA or SM-GGT3.
Isolation from soil. The direct
isolation of Ggt from soil has rarely been accomplished; probably because it is
a highly specialized root parasite that does not form or release spores into the
soil (3). Even the use of the semi-selective medium SM-GGT3 has not proven
successful, since it will not restrict the growth of all organisms except Ggt
(11). Some have been successful in isolating Ggt from soil if the hyphae are
handpicked from it, but in practice this has not usually been done. The best
method of isolating this fungus from soil is to bait it using a susceptible host
such as wheat seedlings (9, 10, 14). Basically, this procedure involves planting
wheat seeds in the suspect or test soil, incubating them under conducive
temperature (15-20°C), and moisture conditions (-10 kPa or wetter), removing
the seedlings after 3-4 wk, and isolating from them as described previously.
This technique will not detect isolates of zero to low virulence. However, the
use of this technique in conjunction with various soil dilutions allows one to
use the Most-Probable-Number (MPN) test for determining the number of propagules
in the soil (13).
If perithecia are not present, incubating infected stem bases in the light (daylight or fluorescent) under moist conditions, e.g. wrapped in moist paper towels at 15°C, often results in production of perithecia within 6-wk (9). Perithecia are not usually produced in culture on media such as potato dextrose agar (PDA), but can be induced to form on wheat leaf extract agar (WLA) (Boil 100 g of green wheat leaves in 1 l of distilled water for 10 min. Pour through two layers of cheesecloth and adjust to one liter with distilled water. Add 20 g of agar and autoclave) in 6-8 wk (M. Elliott, personal communication) or on an agar medium containing 1 % glucose and 0.2 % asparagine (24). Cultures should be incubated in light for optimal production of perithecia. Another technique to produce perithecia is to place surface-disinfested wheat seeds on an agar culture and allow the fungus to colonize the developing seedling (3). Similarly, sterilized wheat roots can be used as a substrate to induce perithecia formation. Perithecia also form readily from epiphytic growth on soybean pods (Glycine max) (20). However, not all isolates of Ggt will form perithecia. Although Ggt usually attacks wheat and not oats, an oat-attacking strain of Ggt has been described in Australia (27), which has longer ascospores (mean of 94 µm) than non-oat attacking strains (mean of 81 µm) (26).
To induce formation of conidia, Deacon (4) suggested flooding colonies with sterile distilled water. The large, phialidic conidia that develop will germinate in contrast to the micro-conidia that sometimes develop but have not been shown to germinate. However, conidia are not normally observed in young cultures of Ggt on media such as PDA. Conidia are found only in cultures of Ggt, never on naturally infected tissue so their role in take-all is unknown.
Walker (22) suggests growing a fungus suspected of being Ggt on PDA
supplemented with glucose at 10 g/L plus yeast extract at 1 g/L. One
characteristic aspect of the hyphal growth is the curling back of the hyphae on
the edge of the colony. Hyphopodia can develop on wheat coleoptiles when a
surface-disinfested grain is placed on established agar cultures.
Because Ggt normally exists in culture as mycelia, most researchers store it
in this form on slants of PDA, or 1/5 strength PDA (23) with transfer every 6 mo
to fresh media. However, many cultures will lose virulence over time whether
transferred repeatedly or not (16). Thus, it is recommended that isolates be
'passed' through a susceptible host with subsequent reisolation from infected
tissue (16). To avoid the problem of loss of virulence, storage of the fungus at
-70°C or lower is desirable. This process can be done by placing small
(8-mm-diameter) plugs of mycelium grown on PDA in 15% glycerol, which is placed
at -70°C. To recover the fungus, the glycerol plus mycelium is thawed, and the
mycelium is placed on PDA.
Inoculum production. A variety of methods are used to produce inoculum of Ggt for use in the greenhouse or field. No one method seems to be accepted universally so the various techniques will be described and the potential shortcomings or limitations listed.
For greenhouse tests, mycelium grown on PDA is effective as inoculum. Plugs cut from an agar culture can be placed within a growing medium such as sand or vermiculite so the roots must penetrate downward through the mycelium. We often use a small plastic 'Conetainer' (Ray Leach, Inc., Canby, OR) 25-cm tall, 2.5 cm-diameter and open at the bottom for drainage. A 20-mm-diameter inoculum plug fits nicely inside the tube so that the roots cannot escape contact with the inoculum. Hollins et al. (8) used a sand-cornmeal-water medium (500 g dry sand, 15 g cornmeal, 65 ml water) or a vermiculite-cornmeal-water medium (50 g vermiculite adjusted to pH 7.5, 30 g cornmeal, 180 ml water) for growth of Ggt. The fungus is cultured on these media for 4 wk with weekly shaking. They used the sand-cornmeal medium in the greenhouse and the vermiculite-cornmeal medium in the field. Halloran (7) used another formula for the sand-cornmeal-water (100: 13:13, by weight respectively) medium in 500 ml flasks autoclaved for 1 hr, inoculated with an 8-mm plug of mycelium, and then incubated for 1 mo.
A technique used by many researchers involves the growth of Ggt on sterilized oat kernels for 4-wk at 20°C, followed by air-drying. We have used the following procedure: 150 ml dry oats are soaked overnight in water, the liquid is then decanted, and the oats are autoclaved for 1 hr at 121°C. Some favor autoclaving the oats on three successive days to insure sterility. Simon et al. (21) have compared this oat kernel medium with one involving ryegrass seed or millet (Panicum miliaceum L.) seed prepared as just described. They favor use of ryegrass seed inoculum because it does not contain unrealistically large food reserves as compared to whole oat inoculum.
Inoculum size and application rate can affect infectivity. When infested oat kernels were fractured into various sizes, the maximum effect of inoculum was observed with the 0.71-1.0 mm fraction. Others (25) also noted that the most infectious fragments are 0.5 mm or larger in size. In greenhouse tests, inoculum rates of 0.1% to 5.0% by weight were used when mixed with soil or other growth mixes. In the field, inoculum can be spread over the soil surface and then rotovated into the soil, or can be applied directly into the drill row either with the seed or below the seed. Placement of inoculum below the seed is accomplished by one pass with drill openers set deep to "place" inoculum only followed by a second pass in the same rows but with openers shallow to plant seed only. We found that rates of 1-5 g of fractured oat kernels per 3.3 m of row applied simultaneously with the seed to the drill row provides a high degree of infestation. Rothrock (19) spread fractured oat kernels over the soil surface at rates of 1.8 or 3.6 g/m2 with disease incidence of nearly 100 % at either rate. MacNish et al. (15) described a medium of very low nutrient status which includes equal parts by volume of millet seed, silica sand, and water. These three components are mixed, placed in trays, and autoclaved for 1 hr at 121°C. Twenty-four hours later the components are again thoroughly mixed and autoclaved. The next day 1-L glass jars are half-filled with this mixture and autoclaved for 20 min. This procedure prevents a hard coagulation of the sand and millet in the bottom of the jar. The medium is inoculated with Ggt, incubated for 3 wk at 25°C and the contents then air-dried and split into colonized millet seed and surface-colonized sand by passing it over a 1.4-mm mesh sieve. Small pieces of millet seed are removed by winnowing in front of a fan. The colonized sand grains are used at rates up to 1300 grains per 100 cm3 of soil, with 800 - 1,000 grains per 100 cm3 providing maximum infection.
Pathogenicity determinations. Tests of pathogenicity Ggt isolates can be carried out in the greenhouse or the field. The degree of pathogenicity or virulence is assessed in a variety of ways including disease rating or scoring of infected roots, reduction in dry weight of foliage, production of white heads after anthesis, and reduction in thousand kernel weight and yield of grain. When pathogenicity towards seedlings is rated, many have used a disease scoring system where numbers are used to represent the percentage of roots infected. One such system (8) is as follows: 0=no roots with blackened tissue, 1=1-25% of roots with blackened tissue, 3=26-50% blackened, 5=51-75% blackened, 7=76-100% blackened. This same disease index has been used to rate field-grown plants for disease severity (8). Although other disease scales have been used, most of them are based on the degree of infection of the roots.
To facilitate the rating of roots for infection, some use a root-assessment
tray (2). This tray involves one compartment through which running water flows
to remove soil and plant debris, with a second compartment filled with calm
water for viewing the roots with a dissecting microscope (Fig. 11). Most
researchers favor examination of infected roots under water with a white
The most current key and taxonomic reference to Gaeumannomyces is that
of Walker (22).
1. Cohen, R., Krikun, J., and Amiv, J. 1982. Appearance of take-all of wheat in Israel. Hassadeh 72:1726-1727 (In Hebrew).
2. Cotterill, P. J., and Sivasithamparam, K. 1988. Use of a root assessment tray for the detection of take-all on lateral roots of wheat seedlings. Plant and Soil 110:140-142.
3. Cunningham, P. C. 1981. Isolation and culture. Pages 103-123 in: Biology and Control of Take-all. M.J.C. Asher and P.J. Shipton, eds. Academic Press, New York.
4. Deacon, J. W. 1973. Phialophora radicicola and Gaeumannomyces graminis on roots of grasses and cereals. Trans. Br. Mycol. Soc. 61:471-485.
5. Elliott, M. L. 1989. An improved selective medium for isolation of Gaeumannomyces-like fungi. Phytopathology 79:1177.
6. Garrett, S. D. 1981. Introduction. Pages 1-11 in: Biology and Control of Take-all. M.I.C. Asher and P.J. Shipton, eds. Academic Press, New York.
7. Halloran, G. M. 1974. Ophiobolus graminis resistance in the genera Agropyron and Secale and its possible significance to wheat breeding. Euphytica 23:225-235.
8. Hollins, T. W., Scott, P. R., and Gregory, R. S. 1986. The relative resistance of wheat, rye, and triticale to take-all caused by Gaeumannomyces graminis. Plant Pathol. 35:93-100.
9. Hornby, D. 1969. Methods of investigating populations of the take-all fungus (Ophiobolus graminis) in soil. Ann. Appl. Biol. 64:503- 513.
10. Hornby, D. 1981. Inoculum. Pages 271-293 in: Biology and Control of Take-all. M. J. C. Asher and P. J. Shipton, eds. Academic Press, New York.
11. Juhnke, M. E., Mathre, D. E., and Sands, D. C. 1984. A selective medium for Gaeumannomyces graminis var. tritici. Plant Dis. 68:233-236.
12. Latham, A. J., and Linn, M. B. 1961. A non-fungistatic bacteriostatic medium containing streptomycin and vancomycin. Plant Dis. Rep. 45:866-867.
13. Maloy, O. C., and Alexander, M. 1958. The "most probable number" method for estimating populations of plant pathogenic organisms in the soil. Phytopathology 48:126-128.
14. MacNish, G. C., Dodman, R. L., and Flentje, N. T. 1973. Bioassay of undisturbed soil cores for the presence of Gaeumannomyces graminis var. tritici. Aust. J. Biol. Sci. 26:1267-1276.
15. MacNish, G. C., Liddle, J. M., and Powelson, R. L. 1986. Studies on the use of high- and low-nutrient inoculum for infection of wheat by Gaeumannomyces graminis var. tritici. Phytopathology 76:815-819.
16. Naiki, T., and Cook, R. J. 1983. Factors in loss of pathogenicity in Gaeumannomyces graminis var. tritici. Phytopathology 73:1652- 1656.
17. Nilsson, H. E. 1969. Studies of root and foot rot diseases of cereals and grasses. 1. On resistance to Ophiobolus graminis Sacc. Lantb - Hogsk. Annlr. 35:275-807.
18. Nilsson, H. E., and Smith, J.D. 1981. Take-all of grasses. Pages 433 - 448 in: Biology and Control of Take-all. M. J. C. Asher and P. J. Shipton, eds. Academic Press, New York.
19. Rothrock, C. S. 1988. Relative susceptibility of small grains to take-all. Plant Dis. 72:883-886.
20. Roy, K. W., Abney, T. S., Huber, D. M., and Keeler, R. 1982. Isolation of Gaeumannomyces graminis var. graminis from soybean in the Midwest. Plant Dis. 66:822-825.
21. Simon, A., Rovira, A. D., and Foster, R. C. 1987. Inocula of Gaeumannomyces graminis var. tritici for field and glasshouse studies. Soil Biol. Biochem. 19:363-370.
22. Walker, J. 1981. Taxonomy of take-all fungi and related genera and species. Pages 15-74 in: Biology and Control of Take-all. M. J. C. Asher and P. J. Shipton, eds. Academic Press, New York.
23.Weller, D. M., and Cook, R. ]. 1983. Suppression of take-all of wheat by seed treatments with fluorescent Pseudomonads. Phytopathology 73:463-469.
24. Weste, G., and Thrower, L. B. 1963. Production of perithecia and microconidia in culture by Ophiobolus graminis. Phytopathology 53:354.
25. Wilkinson, H. T., Cook, R.J., and Alldredge, J. R. 1985. Relation of inoculum size and concentration to infection of wheat roots by Gaeumannomyces graminis var. tritici. Phytopathology 75:98- 103.
26. Yeates, J. S. 1986. Ascospore length of Australian isolates of Gaeumannomyces graminis. Trans. Br. Mycol. Soc. 86:131-136.
27. Yeates, J. W., Fang, C. S., and Parker, C.A. 1986. Distribution and
importance of oat-attacking isolates of Gaeumannomyces graminis var. tritici
in Western Australia. Tr. Br. Mycol. Soc. 86:145- 152.