Diseases of Forage Grasses in Humid Temperate Zones

Foreword

Continuing conversion of agricultural lands to non­ agricultural use dictates the need for continued im­provement in the quality and production of forage grasses in grassland agriculture. The identification of forage grass diseases as they occur in the field is of paramount importance prior to the recommendation of adequate control measures or to the development of resistant cultivars through forage grass breeding programs. It is also important to develop improved techniques for expediting disease resistance and to facilitate the incorporation of disease resistant germplasm into forage grasses. The value of these improvement programs cannot be overestimated.

This bulletin recognizes the fiftieth anniversary of the U.S. Regional Pasture Research Laboratory. The Laboratory is located on the campus of The Pennsyl­vania State University, University Park. The Pasture Laboratory staff works in close cooperation with Pennsylvania Agricultural Experiment Station work­ers on forage problems of mutual interest.

Charles R. Krueger
Associate Dean for Research and Associate Director of the Pennsylvania Agricultural Experiment Station.

Overview

Forage grasses are utilized as pasture and hay and are a basic foundation of United States agriculture. These grasses support the production of beef and dairy cat­tle, each an annual multibillion dollar enterprise. The 1979 U.S. Department of Agriculture's Agricultural Statistics (1979), indicated that there were 110.9 mil­lion head of cattle worth $44.7 billion feeding on over 100 million acres of pastureland in the United States. Recent developments in grassland agriculture have focused on improved management practices which include planting warm-season forage grass species, increasing yields, quality and nutrient content of the grasses, disease resistance, and other mechanisms of disease control.

The warm-season grasses include big bluestem (Andropogon gerardi Vitman), little bluestem (Schizachrium scoparium (Michx.) Nash), indiangrass (Sorghastrum nutans (L.) Nash) and switchgrass (Pani­cum virgatum L.). These major native grasses of North America are distributed throughout the central and, to a lesser degree, the eastern United States. They are characterized by drought tolerance and relatively short growing season (mid-May to mid-September). Cool-season grasses grow from early April to Novem­ber. The peak-production period for warm-season grasses is the hot, often dry, months of July and Au­gust, a time when cool-season grasses produce little forage. For this reason, they are important supple­ments to the cool-season species for grazing and hay production on many farms.

Forage grasses are subject to a multitude of leaf, stem, floral, and root diseases. The leaf diseases in­clude rusts, smuts, and leafspots, and cause the ma­jor reductions in yields, as harvested forage consists primarily of leaf material. Grass diseases may limit ef­fective utilization of the grass species and affect herb­age quality, digestibility, tillering, and root growth. Perennial grasses are usually prone to one or more diseases that, over an extended period, may weaken the plant. A weakened plant is less likely to survive winter-kill.

Resistance in certain cultivars offers an efficient ap­proach to control of several of the more serious fun­gal, bacterial, and viral-caused diseases. Maintaining resistance in a cultivar may require continuous effort because disease resistance often loses effectiveness. Loss of effective resistance may be due to the acquisi­tion of new virulent genes in the pathogen. It be­ comes imperative to seek new and better sources of disease resistance from as broad a germplasm base as possible. Resistance to many of the diseases of the Gramineae has been reviewed by Braverman (1967, 1986).

This bulletin describes control measures for com­mon disease incitants on selected forage-, range-, and pasture-type grasses. The majority of the pathogens discussed are common in the United States. How­ever, a few bacteria and viruses not known to occur in the United States are included because of their de­structive potential, should the causal agents become established. Host susceptibility, favorable tempera­ture and moisture, and suitable vectors for virus transmission are the major determinants of inci­dence, rapidity, and severity of disease development. Resistant cultivars offer a prime means of control, and are most effective when used with good farm management and agricultural practices. Tillage, sani­tation, crop rotation, fertilization, and proper time of harvesting all influence development of diseases on farm crops.

Incitants discussed here are divided into the biotic agents - bacterial, fungal pathogens, and viral agents, and then selected abiotic agents, including environ­mental factors and nutritional disorders. Publications referenced for each disease are listed with the discussion; a complete bibliography is presented at the end.

This publication has been written for growers, teachers, and researchers concerned with grassland production. The biotic and abiotic diseases which af­fect temperate forage grasses are described. Control measures are listed, if known. A glossary defines se­lected technical terminology.

Nematodes damage fine turf. However, to the au­thors' knowledge there are no reports of such dam­age to range and pasture grasses.

Initially, the common name for each host species, followed by the Latin binomial, is used. Thereafter, only the common name is listed. Binomials not ac­companied by an appropriate common name indi­cates none was identified.

Diseases are listed according to hosts at end.

Literature

Braverman, S. W. 1967. Disease resistance in cool season forage, range and turf grasses. Bot. Rev. 33:329-378.

_________. 1986. Disease resistance in cool season forage, range and turf grasses II. Bot. Rev. 52:1-112.

United States Department of Agriculture. 1979. Agri­cultural Statistics, 1979.

Diseases incited by Bacteria

Symptoms caused by bacteria may be similar to those incited by fungi or viruses. However, symptoms characteristic of bacterial infections include water­ soaked lesions on foliage in the early stage of disease development. When observed in transmitted light, such lesions appear to be greasy. Under suitable con­ditions, bacterial exudates may occur on the diseased plant parts (Figure 1).

Figure 1. Cells of Xanthomonas campestris pv. graminis exuding from the vascular system of annual ryegrass (Lolium multiflorum Lam.)

Plant pathogenic bacteria are unicellular rods from 1 to 3 µm in length. They do not have well-defined nuclei or nuclear membranes. They do not form spores, but are covered with a slime which aids in survival under adverse conditions. Bacterial patho­gens are most commonly spread by plant debris, water, and insects. Bacteria can persist on seeds, in­fected plants, plant debris, and soil. Free moisture is usually required for infection, and penetration of host tissue is through wounds or natural openings. Bacterial pathogens invade the vascular system or in­tercellular spaces in host tissue. Necrosis is com­monly due to the effect of toxins or enzymes pro­duced by the pathogen.

At present, the control measures on field crops are sanitation and use of pathogen-free seed. Little is known about resistance in varieties of forage grasses. Bactericidal sprays have been used effectively in con­trol of some vegetable and ornamental crop diseases, but are too expensive for forage crop disease control.

Bacterial taxonomy, always in a state of flux, has undergone a large change in recent years. The prolif­eration of new techniques for determining genetic and phenotypic relationships along with the estab­lishment of regional repositories for type cultures makes it convenient for scientists to compare their cultures with the original type culture. Such compari­sons have shown that a large number of old species are phenotypically and genetically related, but have different host ranges. Thus, it is possible to group a number of closely related species under one species and create a new designation, "pathovar," as a sub­ group based on host range. Advantages of this new system become apparent upon examination of the re­cent literature on pseudomonads and xantho­monads. In the former, the species "syringae" includes all phytopathogenic pseudomonads that are oxidase negative and produce green fluorescent pig­ment. In the genus Xanthomonas, the species "campestris" consists of 123 pathovars grouped to­gether because of their characteristic growth on a spe­cific medium.

Study of this group of pathogens is not sufficiently complete to determine with certainty the existence of pathovars within the bacterial pathogens of forage grasses. Where the literature is clear that such a pathovar exists, we have used that designation. We have relied heavily on the work of others for descrip­tions of symptoms of these diseases on the small grains discussed herein.

Yellow Slime Disease of Orchardgrass

(Dactylis glomerata L.) caused by Corynebacterium rathayi (Smith 1913) Dowson 942.

Figure 2. Yellow slime, Corynebacterium rathayi, on orchardgrass.

Yellow slime disease, or bacteriosis, of orchardgrass occurs sporadically in Britain where it is introduced on imported seed. In Denmark and northern Germany, the disease is well established and can cause serious losses to seed growers. The bacteria also occur in the United States.

Symptoms

A yellow slime consisting of bacterial cells appears on the surface of the upper plant parts, especially the inflorescences. A dwarfing and distor­tion, through incomplete elongation of the upper in­ternodes, is also evident. Because of the sticky slime, the stalks often form knee-shaped bendings due to elongation of the under part of the stuck stem, and the inflorescence may push out laterally. Infected parts dry prematurely. Vessels and parenchyma are invaded by bacteria (Bradbury, 1973a).

Very similar diseases have been reported on other grasses as caused by other species of Corynebacterium.

Etiology

The etiology is not fully known. Apparently, wet periods in May and June favor develop­ment of yellow slime disease. The nematode Anguina tritici (Steinbuch 1799) Chitwood 1935, associated with spread of Corynebacterium tritici (Hutchison) Burkholder 1948 on wheat, may serve as a vector of C. rathayi to wheat (Triticum aestivum L.). But as this nematode does not go to Dactylis, Sabet (1954) sug­gested that an unknown nematode may serve as a vector. Bradbury (1973a) stated that direct plant to plant transmission seems very unlikely.

Host Range

The organism has been reported on or­ chardgrass, bermudagrass (Cynodon dactylon (L.) Pers.), and rye (Secale cereale L.) (Bradbury, 1973a).

Literature

Bradbury, J. F. 1973a. Corynebacterium rathayi. CMI Descriptions of Pathogenic Fungi and Bacteria. No. 376. Kew: Commonwealth Mycological Institute.

Sabet, K. A. 1954. On the host range and systematic position of the bacteria responsible for the yellow slime diseases of wheat (Triticum vulgare Vill.) and cocksfoot grass (Dactylis glomerata L.). Ann. Appl. Biol. 41:606-611.

Bacterial Stripe of Sorghum

(Sorghum bicolor (L.) Moench = S. vulgare Pers.), caused by Pseudomonas andropogonis (Smith 1911) Stapp 1928 (Pseudomonas stizolobii (Wolf 1920) Stapp 1935).

Figure 3. Bacterial stripe, Pseudomonas andropogonis, on sudangrass.

This bacterial species has a wide host range that in­cludes gramineous, leguminous, and ornamental species. In 1981, P. andropogonis reportedly caused a bacterial dark spot of coffee in Brazil (Hayward, 1983). There is some evidence of pathogenic speciali­zation, suggesting that listing of pathovars may be­ come possible.

Symptoms

The bacteria cause red streaks and blotches on the leaves and sheaths. Lesions at first appear to be water-soaked dark green specks. Single lesions are only a few millimeters in diameter but may extend to appear later as long red streaks be­ tween the veins. When lesions coalesce, they may cover a large part of a leaf blade. The red coloring is not marginal but is continuous throughout the lesion. Bacterial exudate occurs on the undersurface of the leaves; when dry, the exudate appears as light red scales. Lesions are similar in form on all varieties, but may vary from deep reddish-brown or purple to brick red. Lesions on a few varieties are tan to brown, but show no reddening, suggesting that difference in color is due to the host.

Etiology

The bacteria enter through the stomata. Spread is probably by wind-driven rain.

Host Range

Agropyron intermedium (Host.) Beauv. is resistant to certain strains but susceptible to others (Tominaga, 1968a, 1971).

Literature

Hayward, A.C. 1983. The Non-Fluorescent Pseudo­monads. In "Plant Bacterial Disease, A Diagnostic Guide." p. 107, (Ed. P.C. Fahy and G.J. Persley). Academic Press.

Tominaga, T. 1968a. Brown stripe of bromegrass and wheatgrass caused by Pseudomonas setariae (Okabe) Savulescu. Jap. J. Bact. 23:176-183.

_________. 1971. Studies on the diseases of forage crops in Japan. Bull. Nat. Inst. Agric. Sci. Gap.) Ser. C. No. 25. p. 205-306.

Bacterial Brown Stripe of Yellow Foxtail and Halo Blight of Oats

Bacterial Brown Stripe (Alopecurus spp.), Halo Blight of Oats (Avena sativa L.) and other grasses caused by Pseudomonas avenae Manns 1909 (Pseudomonas alboprecipitans Rosen 1922).

Figure 4. Halo blight, Pseudomonas avenae, on oats.

Figure 5. Brown stripe, Pseudomonas avenae, on mountain bromegrass.

Pseudomonas avenae is the cause of bacterial leaf blight of oats (Avena saliva L.), bacterial leaf blight, stalk rot of maize (Zea mays L.) and teosinte [Euchlaena mexicana Schrad. ( =Zea mays spp. mexicana (Schrad.) Iltis)], and bacterial brown stripe of foxtail and other grasses (Bradbury, 1970b). The causal agent was named Pseudomonas alboprecipitans until Schaad et al. (1975) demonstrated that the criterion used to estab­lish this species was not valid, and recommended the designation Pseudomonas avenae. Investigations by Tominaga (1971) and a summary of available data by Bradbury (1973b) suggest that P. alboprecipitans (P. avenae) is closely related to P. setariae (Okabe) Burkholder 1939; Tominaga (1971) considers them synonymous. In addition to their identical culture tests, Tominaga (1971) reported that two isolates of P. alboprecipitans were serologically identical to one of P. setariae and closely related to a second. Goto (1964) suggested that P. setariae was synonymous with P. panici and P. panici-miliacei (lkata) Yaumati 1947.

In conclusion, under current rules and findings, P. avenae is the only valid name for bacteria still referred to in the literature as P. alboprecipitans, P. panici, P. panici-miliacei, and P. setariae.

Symptoms

These bacteria induce light brown spots and streaks of no particular size or shape (Bradbury, 1973b). The spots occur mostly on the blades and sheath, but the lesions may occur on any above­ ground part of the plant. Depending on the host, these lesions may vary from indefinite light yellow areas to greyish-green withered spots. On oats, there is often a reddish tinge. On broomcorn millet (Pani­cum miliaceum L.), this organism causes narrow, brown, water-soaked streaks on leaves, sheaths, and culms. When the streaks coalesce, the tissue becomes brown and translucent. Abundant exudate dries, forming shiny thin scales along the streaks. Similar lesions occur on the peduncles and pedicles of the panicle. In severe infections, the entire upper part of the plant is killed, and new shoots emerge at the base.

Etiology

The presence of bacterial exudates on the lesions suggests transmission by wind and rain (Bradbury, 1973a). The bacteria enter the plants through stomata and hydathodes (Bradbury, 1973a).

Host Range

These bacteria have been recorded as occurring on, or causing lesions when inoculated on, rice (Oryza sativa L.), foxtail millet (Setaria italica (L.) P. Beauv.), broomcom millet, barley (Hordeum vulgare L.), Agropyron intermedium, A. trichophorum (Link) Richt., rescuegrass [Bromus unioloides H.B.K. ( = B. catharticus Vahl)], smooth bromegrass (B. inermis Leyss.), B. marginatus Nees ex Steud., yellow bristle­ grass (Setaria lutescens Weigel), teosinte, meadow fox­ tail (Alopecurus pratensis L.), tall oatgrass (Arrhenatherum elatius (L.) Beauv. ex. J. & C. Presl.), oats, Japanese millet (Echinochloa frumentacea (Roxb.) Link), meadow fescue (festuca pratensis Huds.), vel­vetgrass (Holcus lanatus L.), annual ryegrass (Lolium multiflorum L.), sorghum, and wheat.

Literature

Bradbury, J.F. 1970b. Pseudomonas setariae. CMI De­scriptions of Pathogenic Fungi and Bacteria. No. 237. Kew: Commonwealth Mycological Institute.

__________. 1973a. Corynebaderium rathayi. CMI Descriptions of Pathogenic Fungi and Bacteria. No. 376. Kew: Commonwealth Mycological Institute.

__________. 1973b. Pseudomonas alboprecipitans. CMI De­scriptions of Pathogenic Fungi and Bacteria. No. 371. Kew: Commonwealth Mycological Institute.

Goto, M. 1964. Nomenclature of the bacteria causing bacterial leaf streak and bacterial stripe of rice. Rep. Fac. Agric. Shizwoka Univ. 14:3-10.

Schaad, N.W., C.I. Kado, and D.R. Sumner. 1975. Synonymy of Pseudomonas avenae Manns 1905 and Pseudomonas alboprecipitans Rosen 1922. Int. J. Syst. Bact. 25:133-137.

Tominago, T. 1971. Studies on the disease of forage crops in Japan. Bull. Nat. Inst. Agric. Sci. (Jap.) Ser. C. No. 25. p. 205-306.

Bacterial Leafblight of Foxtail

(Setaria lutescens) caused by Pseudomonas syringae van Hall 1902.

This pathogen is quite cosmopolitan in its host range, producing symptoms that vary somewhat on differ­ent hosts.

Symptoms

Symptoms vary, depending on the hosts. Lesions on all hosts are usually round-oblong, linear to irregular, and of various sizes. The lesions are not usually limited by the vascular bundles. On foxtail, the spots are small and dark brown, surrounded by a narrow light green halo. On sorghum, sudangrass (S. sudanense (Piper) Stapf), and Johnson grass (S. halepense (L.) Pers.) (Kendrick, 1926), the spots are red, or light-centered with a red margin, depending on the variety. Some varieties have spots with dark brown borders. Lesions on pearl millet (Pennisetum typhoides (Burm.) Stapf & C.E. Hubb) are dark brown and have a slight halo.

Etiology

Infection is through stomata or wounds. The organism is at first intercellular, but soon causes a collapse of the tissues. It then becomes intracellular. The presence of a halo often reflects reaction to a toxin. The causal agent may overwinter in seed, on seed, in stubble, or in the soil.

Host Range

In inoculation trials, the following species were resistant to P. syringae: Triticum vulgare Vill., oats, smooth bromegrass, tall oatgrass, orchardgrass, perennial ryegrass (Lolium perenne L.), Phalaris sp., Agrostis spp., timothy (Phleum pratense L.), meadow fescue (Festuca pratensis), and the following varieties of Setaria lutescens: Japanese, Siberian, common, Hungarian, and broomcorn.

Literature

Kendrick, J.B. 1926. Hokus bacterial spot of Zea mays and Holcus species. Iowa Agric. Exp. Res. Bull. 100.

Basal Glume Rot

caused by Pseudomonas syringae pv. atrofaciens (McCulloch 1920) Young, Dye & Wilkie 1978.

Pseudomonas syringae pv. atrofaciens occurs on wheat, barley, rye, and triticale (Dowson, 1957, Zillinsky, 1983). At this time, it has not been reported on forage grasses.

Symptoms

The pathogen attacks the leaves and in­florescences of the plant. The young lesions on inoculated leaves are small, dark water-soaked spots. These enlarge and elongate, turn yellow and finally tum light brown as the tissues dry. Unlike those pro­duced by the black chaff pathogen, Xanthomonas cam­pestris pv. translucens (Jones, Johnson & Reddy 1917) Dye 1978, the spots are not translucent nor do the bacteria exude in droplets to form encrustations. In­fected glumes exhibit dull brownish black areas at the base (basal glume rot), sometimes extending over nearly the entire surface of the glume. The dark stain­ing is more pronounced on the inner sides of glumes and lemmae, and the staining may extend to the ra­chis and kernels.

Etiology

Little is known about the disease cycle of this bacterium.

Host Range

Oats, barley, and wheat are hosts, but occurrence of the pathogen on forage grasses has not been reported.

Literature

Dowson, W.J. 1957. Plant Diseases Due to Bacteria. Cambridge Press. 232 p.

Zillinsky, F.J. 1983. Common Diseases of Small Grain Cereals. A Guide to Identification. CIMMT. Mexico. 141p.

Brown Spot/Halo Blight of Ryegrasses and Fescues and Blackish-brown Stripe on Bromegrasses

caused by Pseudomonas syringae pv. coronafaciens (Elliott) Stevens (Dye et al. 1980). Syn. Pseudomonas striafaciens (Elliott 1927) Starr & Burkholder 1942. Figures 6, 7, 8

Figure 6. Halo blight, Pseudomonas syringae pv. coronafaciens, on mountain bromegrass.

 

Figure 7. Halo blight, Pseudomonas syringae pv. coronafaciens, on annual ryegrass (early spring symptoms).

 

Figure 8. Halo blight, Pseudomonas syringae pv. coronafaciens, on annual ryegrass (late summer symptoms).

Schaad and Cunfer (1979) examined the physiolog­ical, biochemical, serological, and pathological prop­erties of several strains of Pseudomonas coronafaciens
(Elliott 1920), P. coronafaciens pv. zeae (Riberio, Durbin, Arny & Uchytill 1977), P. coronafaciens subsp. atrapurpurea (Stapp 1928), and Pseudomonas stria­faciens. They concluded that the divisions of strains of P. coronafaciens and the separation of P. coronafaciens from P. striafaciens (Elliott 1927) was not tenable be­ cause of the minor differences in pathogenicity and symptomology. We follow their recommendations and list P. striafaciens as a synonym of P. syringae pv. coronafaciens.

Symptoms

P. syringae pv. coronafaciens causes a spot disease of the foliage, sheaths, and glumes of several Gramineae. On leaves and glumes, the spots are cir­cular to elliptical water-soaked light olive-green spots with brown centers. Later they become linear and dark chocolate or purplish-brown to black. De­pending on the bacterial strain involved, the spots may or may not be accompanied by yellow halos. The centers become slightly depressed. Lesions may coa­lesce and destroy the entire leaf. Upper nodes are sometimes killed, and panicles may wither and die (Bradbury, 1970a,c).

Etiology

Infection takes place through wounds and stomata and is at first intercellular and later intracel­lular in the parenchyma. The pathogen may be trans­mitted on seed and may overwinter in lesions on dead bromegrass (Bradbury, 1970a).

Host Range

The pathogen has been reported (Tominaga, 1968b) to be pathogenic naturally or when inoculated onto the following grass species: quackgrass (Agropyron repens L.), red oats (Avena byzantina K. Koch), rescuegrass, smooth bromegrass, Japanese chess (B. japonicus Thunb. ex Murr.), soft chess (B. secalinus L.), cheatgrass (B. tectorum L.), mountain brome (B. hookerianus Thunb. ( = B. carinatus Hoof & Arn.)), meadow fescue, Hordeum bulbosum L., H. stenostachys Godron, barley, annual ryegrass and perennial ryegrass, timothy, rye, wheat, and maize.

The following grass species have been reported to have resistance to certain strains of the pathogen: big quakinggrass (Briza maxima L.), rescuegrass, Bromus hordeaceus L., smooth bromegrass, foxtail chess (B. rubens L.), orchardgrass, tall fescue (Festuca arun­dinacea Schreb.), meadow fescue, annual and peren­nial ryegrasses, and reed canarygrass (Phalaris arundinacea L.), timothy, and Kentucky bluegrass (Poa pratensis L.) (Tominaga, 1968b).

Literature

Bradbury, J.F. 1970a. Pseudomonas coronafaciens. CMI Descriptions of Pathogenic Fungi and Bacteria. No. 235. Kew: Commonwealth Mycological Institute.

__________. 1970c. Pseudomonas striafaciens. CMI De­scriptions of Pathogenic Fungi and Bacteria. No. 238. Kew:Commonwealth Mycological Institute.

Schaad, N.W., and B.W. Cunfer. 1979. Synonymy of Pseudomonas coronafaciens, Pseudomonas coronafaciens pathovar zeae, Pseudomonas coronafaciens subsp. atropurpurea and Pseudomonas striafaciens. Int. J. Syst. Bact. 29:213-221.

Tominaga, T. 1968b. Halo blight of ryegrass, brome­ grasses and fescues. Ann. Phytopathol. Soc. Jap. 34:242-249.

Bacterial Wilt of Orchardgrass, Annual Ryegrass, and Meadow Fescue

caused by Xanthomonas campestris pv. graminis (Egli, Goto, & Schmidt 1975) Dye 1978.

Figure 9. Bacterial wilt, Xanthomonas campestris pv. graminis, on annual ryegrass.

Bacterial wilt of forage grasses is a relatively new dis­ease, first described by Egli et al. (1975). They ob­ served the bacterium at several locations in Switzer­land, France, and Germany and discussed its disease-inciting potential. Later, Egli and Schmidt (1982) reported its occurrence in the United King­dom, Belgium, and New Zealand, and that its host spectrum is relatively large. They listed species of Lo­lium, Festuca, Trisetum, Dactylis, Phleum, Poa, and Arrhenatherum as hosts.

Symptoms

The disease becomes most evident when the plants begin to head. The young leaves curl and wither, and shoots become stunted or may die. Less severely infected plants continue to form shoots, but their emerging inflorescences are small and distorted. The older leaves often become chlorotic and later necrotic along the margin of the blades. Chlorotic and necrotic zones form along the vascular bundles and usually extend over the length of the leaves into the sheaths. During the colder seasons, young leaves may exhibit such discoloration.

Formation of bacterial droplets or slime on surfaces of diseased leaves has not been reported. If diseased stalks are cut off, yellow bacterial ooze may be ob­served hanging on the inside walls of the lumen. Low-power magnification of the edge of a cut in­fected leaf in water should show masses of bacteria streaming out of the vascular system. Bacteria have been observed in root and stalk sections.

Etiology

Although the etiology has not been re­ported, studies with artificial inoculation have demonstrated that the bacteria enter the plant through wounds, and may be spread by cutting knives. Me­chanical harvesting is a likely mechanism for spread of the pathogen. The bacteria may overwinter in the roots, and seed transmission may also be a possibility.

Host Range

Egli et al. (1975) proposed naming the causal agent of bacterial wilt Xanthomonas graminis be­cause of its specific host range. Dye (1978) and Dye et al. (1980), using the standards for naming pathovars of phytopathogenic bacteria, placed this group of bacteria under X. campestris as a pathovar, i.e., X. c. pv. graminis. Additional research revealed that the agents causing bacterial wilt of forage grasses can be assigned to one of at least four intrasubspecific subdi­visions, distinguished by their host-specific patho­genicity. Egli and Schmidt (1982) identified these four pathovars as: X. campestris pv. graminis, pv. phlei, pv. poae, and pv. arrhenatheri from Phleum, Poa, and Arrhenatherum, respectively. The host range of pv. graminis is relatively wide.

DeCleene et al. (1981) determined that 10 of 11 for­age grass cultivars cultivated in Belgium were suscep­tible to X. campestris pv. graminis.

Literature

DeCleene, M., F. Leyns, M. Van Den Mooter, I. Swings, and J. DeVay. 1981. Reaction of grass vari­eties grown in Belgium to Xanthomonas campestris pv. graminis. Parasitica 37:29-34.

Dye, D.W. 1978. Genus IX Xanthomonas Dowson 1939. In J.M. Young, O.W. Dye, J.F. Bradbury, C.G. Panagopoulos, and C.F. Robbs. A proposed nomenclature and classification for plant patho­genic bacteria. N.Z. J. Agric. Res. 21:153-177.

_________, J.F. Bradbury, M. Goto, A.C. Hayward, R.A. Lelliott, and M.N. Schroth. 1980. Interna­tional standards for naming pathovars of phyto­-pathogenic bacteria and a list of pathovar names and pathotype strains. Rev. Plant Pathol. 59:153-168.

Egli, T., M. Goto, and D. Schmidt. 1975. Bacterial wilt, a new forage grass disease. Phytopathol. Z. 82:111-121.

_________, and D. Schmidt. 1982. Pathogenic variation among the causal agents of bacterial wilt of forage grasses. Phytopathol. Z. 104:138-150.

Translucent Leaf Stripe of Grasses and Cereals

caused by Xanthomonas campestris pv. translucens and related pathovars.

This disease, also known as bacterial stripe and black chaff, is common on cereal crops and occurs in all cereal-growing regions of the world (Zillinsky, 1983). All above-ground parts of the plant may be affected, but the disease occurs most commonly on the leaves and glumes.

Symptoms

Leaf lesions first appear as small water­ soaked areas which enlarge longitudinally, forming irregular translucent stripes which may extend the full length of the blade and sheath. The water-soaked lesions remain translucent for a long period before drying to a yellowish to brownish color. Blotch-like lesions may also occur and may cause portions of the leaf to shrivel and become light brown. Under humid conditions, bacterial exudate appears as small turbid drops along the lesions and hardens into yellowish resinous granules or into a yellow crust. If the flag leaf is infected before the head emerges, the bacterial exudate may prevent the sheath from opening, thereby impeding emergence of the head. If they emerge, the heads may be bent or distorted and part of the seed blighted.

If the glumes and lemmae are affected, the disease is called black chaff, easily recognized by the dark, linear, water-soaked streaks. As the disease prog­resses, the lesions merge, producing a dark staining of the glumes, lemmae, and peduncles. Under suita­ble conditions for the pathogen, the kernels may be­ come stained and shriveled.

Symptoms of black chaff may be confused with the brown necrosis caused by physiological reactions to certain environmental conditions and bacterial stripe caused by Pseudomonas syringae pv. coronafaciens. Therefore, isolation and growth on certain media are necessary for positive identification.

Etiology

X. campestris pv. translucens invades through the stomata, and spreads through the intercellular spaces of the parenchyma. The pathogen survives in its host, and is seed-borne, but will not survive in the soil. During the growing season, local spread is by wind­blown rain and contact.

Host Range

While the symptoms and pathogenic characteristics of the four X. campestris (Pammel) Dowson 1939 pathovars are similar, their host ranges are different. The host range for each of the four pathovars is presented here. The listings, except for that of X. campestris pv. translucens, are from Bradbury (1984). Publications of Tominaga (1967, 1971) have been summarized for the host range of the latter:

• X. campestris pv. cerealis (Hagborg 1942) Dye 1978. Hosts: Agropyron spp., Avena spp., Bromus spp., Hor­deum spp., rye, and Triticum spp.
• X. campestris pv. hordei (Hagborg 1942) Dye 1978. Hosts: smooth bromegrass, Hordeum spp.
• X. campestris pv. phleipratensis (Wallin & Reddy 1945) Dye 1978. Host: timothy
• X. campestris pv. translucens. Hosts: Japanese millet, orchardgrass, smooth bromegrass and quackgrass. Investigations completed by Tominaga (1967, 1971) showed that the following pasture grasses were resistant to the strains of the pathogen tested: red top (Agrostis alba L. = (A. gigantea Roth)), tall oatgrass, smooth bromegrass, Bromus marginatus, soft chess (B. mollis L.), tall fescue, meadow fescue, annual and perennial ryegrasses, and Kentucky bluegrass.
• X. campestris pv. vasculorum (Cobb 1893) Dye 1978. Hosts: signal grass (Brachiaria mutica (Forsk.) Stapf), Guinea grass (Panicum maximum Jacq.), and elephant grass (Pennisetum purpureum (Schum.).

Literature

Bradbury, J.F. 1984. Genus IL Xanthomonas Dowson 1939, 187. In "Bergey's Manual of Systematic Bacte­riology." Vol. 1, p. 199-210, (Ed. N.R. Krieg and J.G. Holt) Williams and Wilkins. Baltimore, Maryland.

Tominaga, T. 1967. Bacterial blight of orchard grass caused by Xanthomonas translucens f. sp. hordei. Hagbord. Jap. J. Bact. 22:628-633.

_________. 1971. Studies on the diseases of forage crops in Japan. Bull. Nat. Inst. Agric. Sci. (Jap.) Ser. C. No. 25. p. 205-306.

Zillinsky, F.J. 1983. Common Diseases of Small Grain Cereals. A Guide to Identification. CIMMT. Mexico. 141 p.

Suggested reading

Bradbury, J. F. 1984. Genus II. Xanthomonas. Dowson 1939, 187. In "Bergey's Manual of System­atic Bacteriology." Vol. 1, p. 199-210, (Ed. N. R. Krieg and J. G. Holt) Williams and Wilkins. Balti­more, Maryland.

Fahy, P. C., and G. J. Persley (eds.), 1983. Plant Bac­terial Diseases. A Diagnostic Guide. Academic Press. 393 p.

Palleroni, N. P. 1984. Family I. Pseudomonadaceae Winslow, Broadhurst, Buchanan, Krumwiede, Rogers and Smith 1917, 555. In "Bergey's Manual of Systematic Bacteriology." Vol. 1, p. 141-199, (Ed. N. R. Krieg and J. G. Holt) Williams and Wilkins. Baltimore, Maryland.

Diseases incited by Fungi

The fungi, an extremely heterogeneous group of lower plants, are devoid of chlorophyll. They exist ei­ther as saprophytes or parasites, with reproductive structures that usually bear well-defined spores. Veg­etative growth is attained primarily by threadlike mycelia that are specialized hyphae. Reproduction is generally by means of spores that are specialized uni­ or multicellular propagative bodies. Fungus spores may be formed asexually or by the sexual process. Spores are microscopic in size, ranging from about 5 to 100 micrometers in length, depending on the spe­cies. They are dispersed by wind, splashing or flow­ing water, animal vectors, and machinery. Although infections of plants are often initiated from germinat­ing spores, in some species infection may be initiated from mycelia from specialized vegetative resting structures such as small seed-like sclerotia.

Diseases incited by Fungi: Rusts

caused by Puccinia spp. and Uromyces dactylidis Otth.

The rust fungi, worldwide in distribution, are highly specialized obligate parasites that cause diseases of high economic significance. They produce from one to five distinct spore forms. Autoecious rusts produce their spores on a single host or on two closely related hosts, and the heteroecious rusts produce spores on two unrelated hosts. Physiologic forms (formae spe­ciales) of rust fungi are differentiated according to their ability to attack different host species.

Crown Rust

caused by Puccinia coronata Cda.

Figure l0. Crown rust, Puccinia coronata, on perennial ryegrass.

Crown rust is worldwide in distribution; specialized forms occur on many cultivated and wild grasses. Many physiological races also occur within formae spe­ciales. In the northeastern United States, P. coronata is destructive on meadow fescue. The disease incitant causes considerable loss of foliage and a reduction in forage quality. Leaves of infected plants usually die. Buckthorns (Rhamnus cathartica L. and R. frangula L.) are alternate hosts of this heteroecious rust. Crown rust is also the most important grass rust in Great Britain, where it causes severe damage in perennial and annual ryegrasses and meadow fescue.

Symptoms

Scattered bright orange-yellow pustules (uredia) develop on the upper and lower leaf surfaces of grass hosts, but are found mainly on the upper side of the leaf. Uredia may also occur on leaf sheaths. In severe epiphytotics, the leaves tum pale yellow and wither completely. A powdery mass of urediospores forms in the uredia. During winter in milder climates, urediospores continue to develop alongside the black, linear telial stage which pro­ duces teliospores. Telia often form in rings around the uredia. In a heavy infestation, infected plants wither, tum yellow, and die from excessive loss of moisture.

Etiology

The fungus overwinters as teliospores. These germinate in the spring, producing basidiospores that infect both buckthorn species. Aecio­spores are subsequently produced on the alternate host and are disseminated by wind and rain to a sus­ceptible grass host. Uredia develop from the aeciospore infection in the host and the life cycle is complete with the subsequent production of telio­spores. In milder climates, the fungus may over­ winter as mycelium in the host or as uredial pustules that resume sporulation in the spring.

Host Range

The host range includes species of Agropyron, Agrostis, Arrhenatherum, Bromus, Dactylis, Elymus, Festuca, Lolium, Phalaris, Phleum, and Poa (Cummins, 1971).

Control

Resistant cultivars offer an appropriate means to control crown rust in economically important Gramineae. Braverman (1967, 1986) recorded many cultivars, comprising several grass genera and species, developed specifically for crown rust resistance.

Literature

Braverman, S.W. 1967. Disease resistance in cool sea­son forage, range and turf grasses. Bot. Rev. 33:329-378.

____________. 1986. Disease resistance in cool season for­age, range and turf grasses II. Bot. Rev. 52:1-112.

Cummins, G.B. 1971. The Rust Fungi of Cereals, Grasses and Bamboo. Springer-Verlag. New York. 570 p.

Stem Rust

caused by Puccinia graminis Pers.

Figure 11. Stem rust, Puccinia graminis, on perennial ryegrass.

Stem rust (black stem rust), worldwide in distribu­tion, is highly destructive on cereals and affects nu­merous grasses of economic importance by reducing yield, quality of forage, and seed production. The fungus is heteroecious, alternating from grass to bar­berry (Berberis vulgaris L.) or Mahonia spp. The disease is most damaging in moderately humid areas.

Symptoms

The red rust, or uredial, stage is promi­ nent on leaves and culms of grasses during the grow­ ing season. As infection progresses, the epidermis is ruptured by uredial pustules containing orange-red spore masses (urediospores). These spores may be­ come wind-blown and spread to other plants, initiating infection. As the diseased plant matures, brown-black, oblong to elongated telia (black rust stage) develop in the uredia or in new sori on sheaths and culms. The telia give rise to black teliospores.

Etiology

The fungus overwinters as teliospores. In more mild climates, overwintering occurs as urediospores or teliospores on the host plant or as dormant mycelium. The teliospores germinate in the spring to produce basidiospores that become wind­ blown and infect leaves of barberry or Mahonia spp. Pycniospores and aeciospores develop on the alter­nate host, and the latter spores infect the grass host. Subsequently, uredia are formed and the life cycle is completed.

Host Range

The host range includes species of Agropyron, Agrostis, Andropogon, Dactylis, Festuca, Lolium, Phleum, and Poa (Cummins, 1971).

Control

Braverman (1967, 1986) listed numerous cultivars of several grass genera and species devel­oped for stem rust resistance.

Literature

Braverman, S.W. 1967. Disease resistance in cool sea­son forage, range and turf grasses. Bot. Rev. 33:329-378.

___________. 1986. Disease resistance in cool season for­ age, range and turf grasses II. Bot. Rev. 52:1-112.

Cummins, G.B. 1971. The Rust Fungi of Cereals, Grasses and Bamboo. Springer-Verlag. New York. 570 p.

Stripe Rust

caused by Puccinia striiformis West.

Stripe rust (yellow rust) occurs on many economically important grasses, particularly in the Northern hemi­sphere. In Ireland, a heavy attack will render forage unpalatable and reduce seed production. In England, P. striiformis is very destructive on orchardgrass (Carr, 1971).

Symptoms

Uredia are lemon-yellow and amphige­nous and become more numerous toward the leaf tips. Telia develop, usually on the lower leaf surface, during late summer and autumn. Teliospores are dark brown to black and are covered by the host's epidermis.

Etiology

The fungus overwinters as urediospores. No aecial stage has been reported; teliospores germi­nate to produce basidiospores which subsequently reinfect the same host and eventually form uredia of this autoecious rust.

Host Range

The host range includes species of Agropyron, Agrostis, Festuca, Lolium, Phleum, Poa, and D. glomerata (Cummins, 1971).

Control

Braverman (1967, 1986) has summarized those genera and species in the Gramineae with re­ported resistance to stripe rust.

Literature

Braverman, S.W. 1967. Disease resistance in cool sea­son forage, range and turf grasses. Bot. Rev. 33:329-378.

___________. 1986. Disease resistance in cool season for­ age, range and turf grasses II. Bot. Rev. 52:1-112.

Carr, A.J. 1971. Grasses. p. 286-295. In Diseases of Crop Plants. J.H. Western Ed. Macmillan. London. 404 p.

Cummins, G.B. 1971. The Rust Fungi of Cereals, Grasses and Bamboo. Springer-Verlag. New York. 570 p.

Yellow Leaf Rust

caused by Puccinia poae-nemoralis Otth.

Yellow leaf rust attacks several grass genera but does not infect cereals. P. poae-nemoralis has several spe­cialized forms, but only the form that parasitizes tall oatgrass has a pycnial and aecial stage on barberry.

Symptoms

The fungus produces round to oval, yellow-brown to red-brown uredial sori on the upper leaf surface and also on the leaf sheaths. The sori are covered by the epidermis and surrounded by chlo­rotic leaf tissue, and may be arranged in dense groups (Weibull, 1983). Telia, rarely found, are brown to black and covered by the epidermis. When present, they are found on the lower leaf surface.

Etiology

The fungus overwinters as urediospores unless teliospores are produced. New infections in the spring result from wind-blown urediospores pro­duced from overwintering mycelium.

Host Range

The host range includes species of Fes­tuca and Poa (Cummins, 1971).

Control

Braverman (1986) has listed a number of resistant P. pratensis cultivars.

Literature

Braverman, S.W. 1986. Disease resistance in cool sea­son forage, range and turf grasses II. Bot. Rev. 52:1-112.

Cummins, G.B. 1971. The Rust Fungi of Cereals, Grasses and Bamboo. Springer-Verlag. New York. 570 p.

Weibull, Peter. 1983. Descriptions of grass diseases No. 12. Weibull's Gras-tips. p. 22-25, December 1983.

Leaf Rust

caused by Puccinia poarum Niels.

Figure 12, Stem rust, Puccinia poarum, on Kentucky bluegrass.

This fungus is primarily a pathogen on Poa spp., with an alternate host of coltsfoot (Tussilago farfara L.), and is worldwide in distribution.

Symptoms

The rust produces oval to elongate orange-yellow uredia on the upper leaf surface and occasionally on the culms. The telia are black and abundant, and appear on both surfaces of the leaf.

Etiology

Aeciospores from coltsfoot are the main source of infection. Urediospores may be disseminated by wind to infect new plants. However, the uredial stage is brief, with telia appearing two to three weeks after an initial infection. The fungus overwinters as teliospores that germinate to form basidiospores in the spring, subsequently infecting coltsfoot.

A distinguishing feature of P. poarum is the produc­tion of two generations of aeciospores, especially in Ireland and Great Britain (O'Rourke, 1976). The first generation of aeciospores appears in late spring and the second in late summer. Urediospore infection oc­curs between each generation of aeciospores.

Host Range

The host range includes species of Agrostis, Festuca, Phleum, and Poa (Smiley, 1983).

Control

Braverman (1986) has listed numerous Kentucky bluegrass cultivars with resistance to P. poarum.

Literature

Braverman, S.W. 1986. Disease resistance in cool sea­son forage range and turf grasses II. Bot. Rev. 52:1-112.

O'Rourke, C.J. 1976. Disease of grasses and forage legumes in Ireland. An Foras Taluntais. Oak Park Res. Centre. Carlow. 115 p.

Smiley, R.W. 1983. Compendium of turfgrass dis­eases. American Phytopathological Society. 102 p.

Uromyces Leaf Rust

caused by Uromyces dactylidis Otth.

The principal leaf rust of orchardgrass, caused by U. dactylidis, occurs in mid to late summer. In the north­eastern United States, severe infection will reduce forage yields and quality (Kreitlow et al., 1953). The rust is heteroecious, with crowfoot (Ranunculus spp.) as alternate hosts.

Symptoms

Yellowish-brown powdery uredia occur, primarily on the upper leaf surface. Telia form on the lower leaf surface or on culms and produce yellow­ brown teliospores.

Etiology

The fungus overwinters as teliospores; these germinate to produce basidiospores. These spores are wind-blown onto crowfoot and develop into the pycnial and aecial stages. Aeciospores on crowfoot are disseminated by rain and wind to an ap­propriate host. Urediospores appear in midsummer.

Host Range

The host range includes Dactylis glomerata and species of Agrostis, Festuca, and Poa (Couch, 1973; Smiley, 1983).

Control

Braverman (1986) listed orchardgrass cultivars resistant to U. dactylidis.

Literature

Braverman, S.W. 1986. Disease resistance in cool sea­son forage, range and turf grasses II. Bot. Rev. 52:1-112.

Couch, H.B. 1973. Diseases of Turfgrasses. Reinhold Publishing Corp. New York. 348 p.

Kreitlow, K.W., J.H. Graham, and R.J. Garber. 1953. Diseases of forage grasses and legumes in the northeastern states. Pa. Agric. Exp. Stn. Bull. 573.

Smiley, R.W. 1983. Compendium of turfgrass dis­eases. American Phytopathological Society. 102 p.

Diseases incited by Fungi: Smuts

caused by Ustilago spp., Urocystis spp., Entyloma spp., and Tilletia spp.

The numerous smuts which occur on grasses infect the inflorescences, leaves, stems (culrns), and caryopses. The spore-bearing sori are specific to certain morphological portions of the grasses. Smuts are worldwide in distribution and can be of major economic importance. If foliage is attacked, forage yield and quality are reduced. If inflorescences are infected, seed yield is reduced.

Smuts are divided into leaf, culm, and head smuts. Leaf smuts of grasses appear on the leaf blades and leaf sheaths. These smuts are divided on the basis of symptoms: stripe smut, flag smut, and spot smut. Sori of these smuts are linear and form long or short stripes between the leaf veins. The leaf epidermis ruptures, discharging the spores and shredding the leaves. Chlamydospores are formed in the shredded tissue. Sori in leaf spot smuts are covered by the epidermis and are more permanent. In seedlings where crown tissue is invaded, infection is followed by a systemic infection of the primordia. Spore formation occurs in the leaves as they become fully developed. In perennial grasses, the smut mycelium may persist in dormant buds and crown tissue for several years. In some strains of stripe smut, infection of bud primordia occurs in established perennial plants. Seed infection occurs in some grasses.

Stripe Smut

caused by Ustilago striiformis (West.) Niessl.

Stripe smut is common on many grasses and world­wide in distribution. Physiologic races and varieties are distinguished on certain grasses.

Symptoms

Stripe smut is systemic, and the patho­gen is soil and seed-borne. Infection results in long, yellow-green narrow streaks which extend the entire length of the leaf blade. The epidermis eventually ruptures, exposing underlying black spore masses. As the spore masses mature, the leaf blades curl down, turn brown, shred, and eventually die. Fungal sori may also appear on the leaf sheath. In a heavy in­fection, growth is retarded and inflorescence development is inhibited.

Etiology

The pathogen overwinters as dormant my­celium in plant debris or as chlamydospores in the soil or on the seed. In the spring, infection hyphae develop from the chlamydospores and penetrate the coleoptiles of young seedlings or tillers in older grasses. The host is systemically colonized. Cool, moist weather favors disease development. Masses of chlamydospores develop in the colonized foliage, and these spores eventually rupture the epidermis and are dispersed by wind and rain.

Host Range

Stripe smut occurs on red top, creeping bentgrass (Agrostis alba Huds.), colonial bentgrass (A. tenuis Sibth.), orchardgrass, timothy, and Kentucky bluegrass. It is less widely distributed on species of Agropyron, Bromus, Elymus, Festuca, Lolium, and reed canarygrass (Fischer, 1953; Smiley, 1983).

Control

Braverman (1986) listed several cultivars of grass genera and species developed for resistance to stripe smut. Fungicide seed treatment and crop rota­tion are useful control practices.

Literature

Braverman, S.W. 1986. Disease resistance in cool sea­son forage, range and turf grasses II Bot. Rev. 52:112.

Fischer, G.W. 1953. Manual of the North American smut fungi. Ronald Press Co. New York. 343 p.

Smiley, R.W. 1983. Compendium of turfgrass diseases. American Phytopathological Society. 102 p.

Flag Smut

caused by Urocystis agropyri (Preuss)Schroet.

Flag smut infects a wide range of the Gramineae and is worldwide in distribution. The disease is usually not destructive.

Symptoms

Symptoms induced by U. agropyri are similar to those of stripe smut, but the disease is man­ifest especially in the upper leaves of infected hosts. According to O'Rourke (1976), diseased plants are stunted, and production of total dry matter and inflo­ rescence is markedly reduced. Sori, which develop beneath the epidermal cells in the leaf, rupture and release powdery masses of spore balls. Chlamydo­spores of flag smut may be distinguished from stripe smut spores with a microscope; flag smut produces spore balls in the smut sori. The spore balls consist of one to four dark, smooth, spores surrounded by smaller, hyaline to pale brown, sterile cells (Kreitlow et al., 1953). Leaf sheaths may also be infected.

Etiology

The etiology of flag smut is similar to that of stripe smut. Infection occurs in coleoptiles of young seedlings or in underground lateral buds of mature plants. The host is systemically colonized, and sori eventually erupt through the epidermis and disperse by wind and rain.

Host Range

Flag smut infects species of Agropyron, Agrostis, Bromus, Elymus, Phleum, Poa, orchardgrass and red fescue (Festuca rubra L.) (Fischer, 1953; Smiley, 1983).

Control

Some systemic fungicides provide an effec­tive control of flag smut in breeding stock nurseries (O'Rourke, 1976). Several Kentucky bluegrass cultivars are noted by Braverman (1986) as resistant.

Literature

Braverman, S.W. 1986. Disease resistance in cool sea­son forage, range and turf grasses II. Bot. Rev. 52:112.

Fischer, G.W. 1953. Manual of the North American smut fungi. Ronald Press Co. New York. 343 p.

Kreitlow, K.W., J.H. Graham, and R.J. Garber. 1953. Diseases of forage grasses and legumes in the northeastern states. Pa. Agric. Exp. Stn. Bull. 573.

O'Rourke, C.J. 1976. Diseases of forage grasses and legumes in Ireland. An Foras Taluntais. Oak Park Res. Centre. Carlow. 115 p.

Smiley, R.W. 1983. Compendium of turfgrass dis­eases. American Phytopathological Society. 102 p.

Leaf Spot Smuts (Blister Smuts)

caused by Entyloma spp.

Leaf spot smuts are the third category of the leaf smuts. They are caused by Entyloma spp. and are common on many grasses. A representative smut of this group, E. dactylis (Pass.) Cif., is worldwide in dis­tribution. These smuts are differentiated from Tilletia and Urocystis in that sori of Entyloma in the leaf forms discolored spots that are frequently light in color, hence the common name "white smuts." Aerial co­nidia may be formed on the surface of the spots, giv­ing the spots an appearance similar to that of pow­dery mildew.

Symptoms

Sori are formed on the leaves and less fre­quently in the floral bracts. Fruiting structures resem­ble tar-like angular to oblong spots. The epidermis over the developing masses of chlamydospores re­mains intact. This gives a blister-like appearance.

Etiology

Entyloma blister smuts are not systemic in the host. According to Smiley (1983), most of the in­fection results from sporidia spreading on the leaves. Sporidia are disseminated by rain, leaf-to-leaf con­tact, and by movement of equipment and animals.

Host Range

The fungus occurs on species of Agrostis, Festuca, Phleum, and Poa (Smiley, 1983).

Literature

Smiley, R.W. 1983. Compendium of turfgrass dis­eases. American Phytopathological society. 102 p.

Culm Smuts

caused by Ustilago spp.

These smuts, also called sheath smuts, are found in western North America, South America, Europe, North Africa, and Asia. Dickson (1956) reports four species: U. spegazzinii (Hirschh.) Fischer ( = U. hypodytes Aucht.) appears to be the most ubiquitous.

Symptoms

Smut sori are superficial on the inter­nodes of the culms and occasionally on aborted inflorescences. Naked linear dark brown to black sori are covered by leaf sheaths. Infections also occur in the crown-bud primordia, and sori appear for two to three seasons following infection. Mycelium will also persist in stolons and crowns of perennial grasses.

Control

Some selections of crested wheatgrass (Agro­pyron cristatum (L.) Beauv.) apparently are resistant (Dickson, 1956).

Literature

Dickson, J.G. 1956. Diseases of Field Crops. McGraw Hill. New York. 517 p.

Head Smuts

caused by species of Ustilago, Sorosporium, and Sphacelotheca.

These smuts, most abundant in western North America, form sori in grass inflorescences. Head smuts (loose smuts) produce spores in the total inflo­rescence, or just in the floral bracts and ovaries. They are differentiated from kernel smuts, in which sori form only in the ovaries.

These three genera of the Ustilaginaceae are differ­entiated by these characters:

• Ustilago--sori naked or without an enclosing membrane and forming black, dusty masses at maturity;
• Sorosporium--sori generally dusty and without an enclosing membrane;
• Sphacelotheca--sori generally in the inflorescence, replacing the kernels; and with a false external membrane.

Ustilago bullata Berk. causes common head or ear smut in a variety of grasses in the western United States and in drier climates; it has also been found in the central and eastern United States. Sori form in the spikelets, involving all or part of the floral bracts, and are enclosed in the epidermal membranes of the floral structure. The loose to semi-covered spore mass is dark brown to purple-black. The powdery mass of spores that replaces the kernels is usually covered by the glume; glumes are rarely destroyed completely. Seedling infection occurs in perennial grasses, and the disease may survive in the host for years.

Host Range

Species of Agropyron, Bromus, Elymus, and Festuca are common hosts (Fischer, 1953).

Control

Several rescuegrass cultivars resistant to head smut have been developed; mountain bromegrass 'Bromar' is also resistant.

Ustilago avenae (Pers.) Rostr. causes a loose smut of tall oatgrass, generally wherever the grass is grown. The seeds are replaced by black, compact sori which eventually become powdery. These sori replace the floral parts.

Literature

Fischer, G.W. 1953. Manual of the North American smut fungi. Ronald Press Co. New York. 343 p.

Kernel (Covered) Smuts

caused by Tilletia spp.

Some kernel smuts caused by species of Tilletia infect only the ovary of the respective host. These smuts are common in western North America, the inter­mountain states, and in similar climatic areas in other countries. The smut sori form in the ovaries and as­sume the general shape of the caryopsis.

Symptoms

This smut affects the floral parts of the host. Diseased seed remains green longer than healthy seed, and when mature is somewhat darker brown than healthy seed. Eventually, only the peri­carp of diseased seeds remain, with their interiors completely replaced with brown, powdery spore masses.

Etiology

The smut fungus survives as chlamydo­spores on the seed and overwinters in this stage. In the spring, chlamydospores germinate concurrently with the seed. Infection hyphae eventually form and penetrate a seedling. The fungus grows within the host until floral formation, at which stage the flower tissue is colonized, and subsequently the entire peri­ carp of the seed is replaced by smut spores.

Host Range

Agrostis spp. are hosts (Fischer, 1953).

Literature

Fischer, G.W. 1953. Manual of the North American smut fungi. Ronald Press Co. New York. 343 p.

Powdery Mildew

caused by Erysiphe graminis DC. ex Merat.

Figure 13. Powdery Mildew, Erysiphe graminis, on orchardgrass.

Powdery mildew occurs on most Gramineae, wher­ever they may be growing. The disease is economic­ally important in grass nurseries, grass breeding stocks, and expansion plantings, but not so in pas­tures. In Great Britain, Davies et al. (1970) have shown that powdery mildew reduces the yield and quality of 'Lior' annual ryegrass.

E. graminis, an obligate parasite on grasses, in­cludes many specialized varieties and races. Few are restricted to a single grass species. Several races may attack a single plant species, and thus provide an ex­cellent opportunity to form new races of the fungus.

Symptoms

The fungus attacks the aerial parts of the plant, but generally only the foliage is injured. The mildew first appears as oblong, irregular, white, powdery blotches on the upper surface of the leaf. Orange-yellow areas develop beneath the white blotches. The powdery mycelial colonies enlarge, and may coalesce until the total leaf surface is covered with grayish-white mycelium. Eventually, the leaf yellows and turns brown. The powdery tufts of my­celium support numerous conidiophores from which conidia are formed in long chains. Symptoms are less evident on grasses grazed or cut closely. Cleistothecia may develop and appear in the powdery mycelial tufts as tiny, round brown-black fruiting structures.

Etiology

The severity of this powdery mildew de­pends primarily upon climatic conditions. Powdery mildew appears during cool and somewhat cloudy conditions. Couch (1973) has shown that in the United States four conditions are essential for maxi­mum mildew development:

1. reduced air circula­tion;
2. high atmospheric humidity;
3. low light in­ tensity,
4. air temperature about 18^oC.

The pathogen overwinters as cleistothecia and mycelia on living host tissue; the cleistothecia are probably of secondary importance (Kreitlow et al., 1953; Weibull, 1978b). New infections are caused by conidia from overwintering host plants. Additional infections may occur, when cleistothecia containing ascospores may develop and serve to maintain the fungus in hot weather or under conditions not favor­able for production of mycelia and/or conidia. Co­nidia will germinate over a wide temperature range. Free moisture is a deterrent to the germination of conidia.

Host Range

Most species of the Gramineae are mil­dew susceptible. The disease is of economic significance in Poa, Festuca, Dactylis, and Bromus. Suscepti­ble species were tabulated by Sprague (1950).

Control

Sulfur dusts, cycloheximide, and systemic fungicides will control this fungus in seed crops and in the greenhouse (O'Rourke, 1976). Chemical con­trol is impractical on grazing lands, and resistant cul­tivars offer the best means of control in these plant­ings. Braverman (1986) has listed numerous grass species reported to be powdery mildew resistant, along with cultivars developed specifically for mil­dew resistance.

Literature

Braverman, S.W. 1986. Disease resistance in cool sea­son forage, range and turf grasses II. Bot. Rev. 52:1-112.

Couch, H.B. 1973. Disease of Turfgrasses. Reinhold Publishing Corp. New York. 348 p.

Davies, H., A.E. Williams, and W.A. Morgan. 1970. The effect of mildew and leaf blotch on yield of cv. Lior Italian ryegrass. Plant Pathol. 19:135-138.

Kreitlow, K.W., J.H. Graham, and R.J. Garber. 1953. Diseases of forage grasses and legumes in the northeastern states. Pa. Agric. Exp. Stn. Bull. 573.

O'Rourke, C.J. 1976. Diseases of forage grasses and legumes in Ireland. An Foras Taluntais. Oak Park Res. Centre. Carlow. 115 p.

Sprague, R. 1950. Diseases of Cereals and Grasses in North America. Ronald Press. New York. 538 p.

Weibull, Peter. 1978b. Descriptions of grass diseases No. 2. Weibull's Gras-tips 21:5-6.

Ergot

caused by Claviceps purpurea Tul.

Figure 14. Ergot, Claviceps purpurea, on orchardgrass.

Ergot affects worldwide the inflorescence of numer­ous cereals and grasses of economic importance in temperate zones. Specialized races occur, particularly in Japan. Many Claviceps species have been described on the basis of host specialization, rather than distinct morphological characteristics. The economic im­portance of C. purpurea is two-fold: affected plants produce ergots (sclerotia) instead of seed, thereby re­ducing yield, and the alkaloidal substances in the er­gots are toxic to livestock or humans feeding on the infected grains. According to O'Rourke (1976), ergot­ infested grain when ingested may cause two distinct syndromes-gangrene of the extremities and stimu­lation of the nervous system, causing convulsions. Ergot-infested grain will cause abortion in cows and ewes (Carr, 1971).

Symptoms

The disease first appears as a sticky "honeydew" ooze on the young ovary. The sugary ooze attracts insects and also forms a substrate for other microorganisms (Cunfer, 1976). This primary infection and subsequent development of a sclero­tium preempts the ovary. The most noticeable phase of the disease is the appearance of horny, curved, dark olive to olive-to-purple sclerotia that project from the inflorescence. A sclerotium is generally two to three times the length of the seed and may be cor­rugated longitudinally. Transverse or longitudinal cracks with reddish margins may occur, exposing the light-colored interior. Sclerotia produce a conspicu­ous odor (Walker, 1970). The sclerotia remain at­tached to the plant until it is mature. Sclerotia in­ fecting large-seeded grass genera such as Arrhenatherum, Bromus, Festuca, and Lolium may be as long as 20 mm. Narrow, smaller sclerotia (up to 10 mm long) are confined to Dactylis, Phleum, and Phalaris, which are grasses with smaller seeds. Unfortunately, sclerotia are usually harvested with the seed.

Etiology

Ergot sclerotia fall to the soil when the host plant matures, and overwinter in soil or in stored grain (Kreitlow et al., 1953). In the spring, the sclero­tia germinate to form minute mushroom-like stalks with rounded heads in which perithecia develop. Ar­ rival of the wind-blown ascospores must occur at full floral development of the host plant. It is believed that insects transmit the ascospores during pollina­tion. Spores infect the young ovaries of the grass flowers by direct penetration, which eventually pro­duces the "honeydew" ooze from this primary infection. The ooze contains millions of tiny conidia of Sphacelia segetum Lev., which is the asexual stage of the fungus. The inoculum is transmitted by insects and water to healthy flowers. The sclerotia eventually develop below the tiny mass of Sphacelia spores, ma­ture, and complete the cycle.

Host Range

Ergot infects a multitude of genera in the Gramineae. Most of the known hosts have been reported by Sprague (1950).

Control

The use of ergot-free seed provides disease control, for it eliminates a potential inoculum source. Ergots can be separated from seed by floating them off in brine. Crop rotations of two to three years be­ tween susceptible crops and legumes or non­-susceptible hosts, mowing of roadside grasses, and cleaning borders and headlands prior to formation of the "honeydew" stage will reduce the secondary inoculum.

Literature

Carr, A.J. 1971. Grasses. p. 286-295. In Diseases of Crop Plants. J.H. Western Ed. Macmillan. London, 404 p.

Cunfer, B.M. 1976. Water potential of ergot honey­dew and its influence upon colonization by micro­organisms. Phytopathology 66:449-452.

Kreitlow, K.W., J.H. Graham, and R.J. Garber. 1953. Diseases of forage grasses and legumes in the northeastern states. Pa. Agric. Exp. Stn. Bull. 573.

O'Rourke, C.J. 1976. Diseases of grasses and forage legumes in Ireland. An Foras Taluntais. Oak Park Res. Centre. Carlow. 115 p.

Sprague, R. 1950. Diseases of Cereals and Grasses in North America. Ronald Press. New York. 538 p.

Walker, J.C. 1970. Plant Pathology. McGraw-Hill Book Co. New York. 819 p.

Blind Seed

caused by Gloeotinia temulenta (Prill & Del.) Wilson, Nobel & Gray (Syn. Phialea temulenta Prill & Del.).

Blind-seed disease may cause considerable losses in seed production. Perennial ryegrass is the most com­mon host, but annual ryegrass and some other grass genera of economic importance are also susceptible. The disease is endemic in nearly all places where rye­ grass is grown for seed-Europe, Oregon (United States), Australia, and New Zealand especially true in regions where the summers are cool, moist and early. An early infection destroys the embryo, while a later infection results in the typical blind seed which fails to germinate. According to O'Rourke (1976), there is conflicting evidence on the possible effects of blind seed ingested by grazing animals. Feeding infected seeds to sheep in New Zealand apparently caused no ill effects. In France, however, ill effects on animals were attributed to the fungus.

Symptoms

The fungus, prevalent in cool, wet grow­ing seasons, infects the developing embryo, which then becomes a partially shrunken rusty-brown cary­opsis rather than a healthy seed. Infected seeds are indistinguishable from healthy seeds but can be de­tected when glumes are removed.

Etiology

The fungus overwinters in infected seeds in the soil and produces above-ground apothecia coinci­dent with the developing ryegrass florets. Cool, damp soil favors an increase in apothecial develop­ment by G. temulenta, and also increases the length of time which inflorescences remain open. Ascospores are wind-blown and gain entry to an open floret just below the stigma, completing the primary infection. Within two weeks of ascospore germination, abun­dant macroconidia are produced in a pinkish, slimy matrix on the seed. The conidia are cylindrical, slightly lunate, and hyaline. Macroconidia are dis­persed by rain, wind-driven water droplets, and by physical contact as inflorescences brush against each other. Macrospores, functioning in a manner similar to the ascospores, set up secondary infections. Microconidia also occur in pink sporodochia on the seeds. An early infection destroys the embryo, pre­venting seed formation and producing the typical "blind seed."

Host Range

The disease is prevalent on annual and perennial ryegrasses and species of Agrostis, Dactylis, Festuca, Phleum, Poa, and Secale.

Control

Pathogen-free seed appears to be a promis­ing approach to control of blind seed disease. Apparently, the fungus will not survive two years in stor­age. Hot-water treatment provides some control. Resistant cultivars also present a reasonable ap­proach to control. Tetraploid cultivars are more sus­ceptible than the early maturing selections. Wright and Breese (1966) and Wright (1967) reported that blind seed disease was under phytogenetic control and designed a backcross program aimed at introducing resistance into 'S.24.' Breeding for disease escape, rather than for direct resistance to the pathogen, has been investigated (Wright, 1967). Ac­cording to O'Rourke (1976), the very early and very late flowering types generally escape infection by the fungus.

Literature

O'Rourke, C.J. 1976. Diseases of grasses and forage legumes in Ireland. An Foras Taluntais. Oak Park Res. Centre. Carlow, 115 p.

Wright, C.E. 1967. Blind seed disease of ryegrass. Euphytica 16:122-130.

________, and E.L. Breese. 1966. The genetic control of blind seed disease resistance in Lolium perenne in relation to breeding practice. Proc. 10th Int. Grassl. Congr. p. 737-741.

Leaf Streak

caused by Scolecotrichum graminis Fckl.

Leaf streak, also known as brown stripe, brown streak, or brown leaf blight, is widely distributed on more than 150 grasses in the temperate zones of the United States, Europe, and South America (Braverman, 1958). The pathogen also attacks cereals and is frequently reported on rye. Brown stripe is present throughout the growing season, but is partic­ularly evident during mid-summer and autumn when leaves and culms are maturing. Severe foliar at­tacks before maturity cause withering and dying of leaves and reduction in the quality of forage (Kreitlow et al., 1953). Morphological variants of the fungus occur. Braverman (1958) and Graham et al. (1963) have shown that infection by conidia from or­chardgrass and tall oatgrass is restricted to the origi­nal host species, whereas the causal agent from timo­thy is pathogenic to both orchardgrass and timothy.

Symptoms

Initial foliar lesions appear as tiny, ellip­tical, purplish-brown water-soaked spots with dark brown borders visible on both leaf surfaces. Subse­quent necrosis of adjacent tissue forms elongated brown streaks. Shape, color, and size of the devel­oping lesions depend upon the host species and the age of the plant. Centers of the older lesions usually are ashy gray to almost white. In a moist atmosphere, dense parallel tufts of conidiophores and conidia develop in older lesions. At the close of the growing season, small subepidermal black stromata form in a parallel arrangement. This is the distinguishing fea­ture of leaf streak. Symptoms are similar on all hosts.

Etiology

Scolecotrichum graminis overwinters as stro­mata in debris and as mycelium in leaf tissue. In spring, the stromata rupture and release conidio­phores and conidia, sources of primary inoculum on new spring growth. The conidia are disseminated primarily by rain.

Host Range

Scolecotrichum graminis attacks a multi­tude of grasses. Smooth bromegrass is one of the few economic grasses resistant to the fungus. A complete list of hosts has been recorded by Sprague (1950).

Control

Disease resistant cultivars offer a practical means for controlling the disease. Several workers have studied the resistance of orchardgrass to S. graminis, and numerous cultivars with varying de­grees of resistance have been developed (Braverman, 1986).

Literature

Braverman, S.W. 1958. Leaf streak of orchardgrass, timothy, and tall oatgrass incited by Scolecotrichum graminis. Phytopathology 48:141-143.

_________. 1986. Disease resistance in cool season for­age, range and turf grasses II. Bot. Rev. 52:1-112.

Graham, J.H., K.E. Zeiders, and S.W. Braverman. 1963. Sporulation and pathogenicity of Scoleco­trichum graminis from orchardgrass and tall oatgrass. Plant Dis. Rep. 47:255-256.

Kreitlow, K.W., J.H. Graham, and R.J. Garber. 1953. Diseases of forage grasses and legumes in the northeastern states. Pa. Agric. Exp. Stn. Bull. 573.

Sprague, R. 1950. Diseases of Cereals and Grasses in North America. Ronald Press Co. New York. 538 p.

Leaf Scald

caused by Rhynchosporium secalis (Oud.) Davis and by R. orthosporum Caldwell.

Both Rhynchosporium secalis and R. orthosporum occur on many cereal and grass hosts; the diseases which they incite are also referred to as leaf blotch and leaf streak. Both species are widely distributed in North America, Europe, and Asia. In the United States, Elliott (1962) recorded the destructive nature of R. or­thosporum on orchardgrass with disease peaks in April-May and July-August. Latch (1966) showed that R. orthosporum-infected perennial ryegrass was unpalatable to sheep in New Zealand, while in the United States Gross et al. (1975) noted that R. secalis reduced the digestibility of smooth bromegrass in grazing animals. In the Nordic countries, R. orthosporum is quite common on orchardgrass, meadow fescue, red fescue, timothy, and Kentucky bluegrass, while R. secalis is common on wheat­ grasses and to a lesser extent on bromegrasses and annual and perennial ryegrasses (Weibull, 1978c).

Symptoms

Leaf scald (R. secalis) appears as dark, bluish-gray water-soaked blotches which become light gray with darker brown margins. The lesions may extend up to 30 mm and constrict and shred the leaf blades. The centers of the lesions are gray and be­come covered with conidia. While R. secalis forms ir­regularly shaped lesions on the upper leaf surface and sheaths, lesions on the under surface frequently show pointed terminals, relating to an overall diamond-shape appearance (O'Rourke, 1976).

Lesions produced by R. orthosporum on grass leaves and sheaths are similar to those caused by R. secalis. O'Rourke (1976) has noted that the scald-like blotches produced by R. orthosporum are usually more diffuse than the lesions produced by R. secalis, but not as diamond-shaped on the under-leaf surface. Lesions on the lower leaf surface are chocolate-brown at the margin and pale in the center. Sprague (1950) de­scribed R. orthosporum on orchardgrass as "white stripe," since mature lesions become elongated and off-white. Lesions caused by either Rhynchosporium species may coalesce, causing death of the leaves and reduction in quality of the forage. Conidia of the two species differ in shape; those of R. secalis have a char­acteristic beak, while conidia of R. orthosporum are slightly narrower and symmetrical.

Etiology

Both species overwinter as dormant myce­lium on dead leaves and crop residue. New lesions may form throughout the winter in milder climates. Conidia develop during cool, moist spring weather and are readily disseminated by wind and rain to healthy leaves. New spores will be produced on older leaves as long as conditions are favorable. Cool weather favors development of the disease, although conidia may germinate in temperatures up to 28^oC. Seed transmission occurs to a limited extent in R. secalis.

Host Range

Sprague (1950) has recorded the range of hosts.

Control

Eliminating crop residues, rotating crops, and seeding resistant cultivars will help to control leaf scald. Use of resistant cultivars was reported by Braverman (1986).

Literature

Braverman, S.W. 1986. Disease resistance in cool sea­son forage, range and turf grasses II. Bot. Rev. 52:1-112.

Elliott, E.S. 1962. Disease damage in forage grasses. Phytopathology 52:448-451.

Gross, D.F., C.J. Martin, and J.G. Ross. 1975. Effect of diseases on in vitro digestibility of smooth bromegrass. Crop Sci. 15:273-275.

Latch, G.C.M. 1966. Fungus diseases of ryegrass in New Zealand. I. Foliar diseases. N.Z. J. Agric. Res. 9:394-409.

O'Rourke, C.J. 1976. Diseases of grasses and forage legumes in Ireland. An Foras Taluntais. Oak Park Res. Centre. Carlow. 115 p.

Sprague, R. 1950. Diseases of Cereals and Grasses in North America. Ronald Press Co. New York. 538 p. Weibull, P. 1978c. Descriptions of grass diseases No. 3. Weibull's Gras-tips 21:7-8.

Mastigosporium Leaf Fleck

caused by Mastigosporium spp.

Figure 15. Mastigosporium leaf fleck, Mastigosporium spp., on orchardgrass.

Species of Mastigosporium infect several valuable for­age grasses, including timothy, perennial ryegrass, and orchardgrass in Europe, New Zealand, and North America. M. rubricosum (Dearn. & Barth.) Sprague is common on orchardgrass in Ireland throughout the year (O'Rourke, 1976). The disease has been reported to reduce orchardgrass yields and forage quality. O'Rourke (1976) indicates that a 10 percent level of infection reduces orchardgrass yield by 30 percent and its water-soluble carbohydrate con­tent by 50 percent.

Symptoms

The disease first appears as elliptical water-soaked spots on both surfaces of the leaf. These darken to purplish-brown flecks with bright orange-tan margins. As the flecks enlarge, the centers become pale and support the growth of many co­nidia. Marginal lesions may cause localized constric­tions of the leaf blade (O'Rourke, 1976). Lesions may also coalesce to form irregular blotches on as much as one-half of the leaf surface.

Etiology

The disease is favored by cool, damp weather, and its effects are most prominent in early spring and autumn. When climate so favors, conidia are borne on short conidiophores that continuously bear new conidia and are dispersed by water. The pathogen persists in leaf tissue during summer and winter and produces conidia when conditions are fa­vorable. Makela (1970) in Finland reported a stro­matic resting stage of M. rubricosum and other Mastigosporium species. Such a stage is probably nec­essary to insure the pathogen's survival during long cold winters.

Host Range

In addition to the grasses already men­tioned, M. rubricosum has been reported on Agrostis spp. (Sprague, 1950).

Control

Several cultivars have been developed for resistance to Mastigosporium leaf fleck (Braverman, 1986). Additional control measures include cutting or grazing the foliage and prompt removal of infected leaves (O'Rourke, 1976).

Literature

Braverman, S.W. 1986. Disease resistance in cool sea­son forage, range and turf grasses II. Bot. Rev. 52:1-112.

Makela, K. 1970. The genus Mastigosporium Reiss. in Finland. Karstenia 11:5-22.

O'Rourke, C.J. 1976. Diseases of grasses and forage legumes in Ireland. An Foras Taluntais. Oak Park Res. Centre. Carlow. 115 p.

Sprague, R. 1950. Diseases of Cereals and Grasses in North America. Ronald Press. New York. 538 p.

Spermospora Leafspot

caused by Spermospora spp.

Fungi of this genus cause a leafspot disease on vari­ous grasses in North America, Ireland, and presuma­bly Europe. Several species have been described, of which S. lolii MacGarvie & O'Rourke may cause mo­derate damage in ryegrasses and fescues.

Symptoms

Elliptical or irregular foliar lesions (de­pending upon the host) may vary from 4 mm to 10 mm in length and are gray-brown or reddish-brown. Discoloration spreads beyond the necrotic tissues (O'Rourke, 1976). Necrotic spots are amphigenous and may be concentrated at the leaf margins, pres­enting a scald effect. Mature lesions on annual and perennial ryegrasses and fescues may become bleached in their centers, giving an eyespot appear­ance. The centers bear whitish masses of conidia.

Etiology

Presumably, S. lolii conidia overwinter on diseased tissue and provide the primary source of in­oculum in the spring. Spread of the pathogen by wind-blown or water-disseminated conidia continues throughout the growing season. In milder climates, S. lolii may produce conidia throughout the year.

Host Range

Sprague (1950) has listed several grasses as hosts to Spermospora spp.

Control

According to O'Rourke (1976), frequent cut­ting restricts build-up of the disease by removing in­fected leaf material.

Literature

O'Rourke, C.J. 1976. Diseases of grasses and forage legumes in Ireland. An Foras Taluntais. Oak Park Res. Centre. Carlow. 115 p.

Sprague, R. 1950. Diseases of Cereals and Grasses in North America. Ronald Press. New York. 538p.

Diseases incited by Fungi: Leafspots

caused by Drechslera spp. and Bipolaris spp. (Syn. Helminthosporium spp.).

Many graminicolous species of Drechslera, formerly classified in the genus Helminthosporium Link, cause leafspots, net blotches, leaf blotches, and occasion­ally foot rots and turf decay on a wide range of Gra­mineae in the temperate areas of North America, Europe, and Japan. Separation into two genera is based primarily upon the manner of spore germina­tion (Shoemaker, 1959). Those species formerly in Helminthosporium whose conidia germinate from all cells were placed in Drechslera (Shoemaker, 1959, 1962), and those whose conidia germinate primarily from end cells were placed in Bipolaris. Species of these two genera exhibit a wide range of host speci­ficity, from D. poae (Baudys) Shoem., that incites melting out of bluegrass, to H. sativum Pam., King & Bakke ( = Bipolaris sorokinianum (Sacc. ex. Sorok.) Shoem.), that causes leafspot/blotch on many grass species.

The perfect (sexual) stages of D. dactylidis Shoem. (Pleospora phaeocomes (Rab.) Wint.), D. bromi (Died.) Shoem., (Pyrenophora bromi (Died.) Drechs.), D. siccans (Drechs.) Shoem. (P. lolii Dovaston), and H. sativum (Cochiobolus sativus (Ito & Kurib.) Dastur.) have been described. Conidia of these and other spe­cies (for which a perfect stage has not yet been reported) serve as primary sources of inoculum. With the exception of D. catenaria (Drechs.) Ito, there is similarity in the etiology of diseases caused by Drechslera spp. and H. sativum. This characteristic is recognized in the discussions that follow. Conidia form on leaves and are disseminated by wind and water droplets, infecting nearby plants. Early spring infections, particularly evident in D. catenaria, D. dictyoides, D. poae, D. siccans, and H. sativum, may be caused by planting infected seed. Brown leafspot caused by D. bromi is most prominent during cool, wet weather of the spring and fall months.

Descriptions, host range, and recommended con­trol measures for seven Drechslera spp. and H. sativum follow.

Literature

Shoemaker, R.A. 1959. Nomenclature of Drechslera and Bipolaris, grass parasites segregated from Helminthosporium. Can. J. Bot. 37:880-886.

________. 1962. Drechslera Ito. Can. J. Bot. 40:809-843.

Brown Leafspot

caused by Drechslera bromi.

Figure 16. Brown leafspot, Pyrenophora bromi, on smooth bromegrass.

Smooth bromegrass is very susceptible to brown leafspot. The fungus usually attacks this host wher­ever smooth bromegrass is grown in the temperate areas of the United States and in Canada. The fungus apparently is restricted to species of Bromus.

Symptoms

The disease first appears as small, dark brown, oblong spots on the foliage which develops in the spring. As the growing season progresses, the spots become dark purplish-brown, elongate, and are surrounded by a yellow band or halo (Kreitlow et al., 1953). Lesions may coalesce, forming large yellow to brown areas. Lesions may envelop the entire leaf.

Host Range

The fungus is restricted to smooth bromegrass and several annual and perennial Bromus spp.

Control

Resistant cultivars offer partial means of controlling the disease (Braverman, 1986). Berg et al. (1983) studied inheritance of leafspot resistance in smooth bromegrass and demonstrated that lesion size is regulated by multiple genes and that suscepti­bility to the fungus may be dominant or epistatic to resistance. For these reasons, they concluded that it will be difficult to develop populations with high lev­els of resistance to the pathogen.

Literature

Berg, C.C., R.T. Sherwood, K E. Zeiders, and R.R. Hill, Jr. 1983. Inheritance of brown leaf spot resis­tance in smooth bromegrass. Crop Sci. 23:138-140.

Braverman, S.W. 1986. Disease resistance in cool sea­son forage, range and turf grasses IL Bot. Rev. 52:1-112.

Kreitlow, KW., J.H. Graham, and R.J. Garber. 1953. Diseases of forage grasses and legumes in the northeastern states. Pa. Agric. Exp. Stn. Bull. 573.

Net Blotch

caused by Drechslera dictyoides (Drechs.) Shoem.

Figure 17. Net blotch, Drechslera dictyoides, on annual ryegrass.

Net blotch, one of the most common foliar diseases of tall and meadow fescue, is prevalent in the eastern and southeastern United States and widespread in the Nordic countries.

Symptoms

Foliar lesions, present throughout the growing season, first appear as scattered, dark brown spots on the upper and lower leaf surfaces. When the disease is severe, symptoms will be seen on the leaf sheath. Lesions expand to about 15 mm long and up to 10 mm wide. The larger lesions often develop into extensive lateral and transverse brown threads, creating the net-like total withering of the leaf. Le­sions usually remain small when the host is growing rapidly. If grass growth is slow, lesions may spread across the leaf blade and cause an early senescence and yellowing of the leaf tip. This yellowing is char­acteristic of a heavy infection (Weibull, 1978a). The net blotch symptom is most evident from spring to autumn.

Host Range

The fungus is widespread on annual and perennial ryegrasses, meadow fescue, and to a lesser extent on tall fescue. D. dictyoides f. sp. dictyoides (Braverman & Graham) Shoem. occurs pri­marily on meadow and tall fescues. D. dictyoides f. sp. perenne (Braverman & Graham) Shoem. occurs mainly on annual and perennial ryegrass (Braverman and Graham, 1960).

Control

Braverman (1967, 1986) has summarized the reported resistance of meadow and tall fescues and annual and perennial ryegrasses to net blotch.

Literature

Braverman, S.W. 1967. Disease resistance in cool sea­son forage, range and turf grasses. Bot. Rev. 33:329-378.

_________. 1986. Disease resistance in cool season for­age, range and turf grasses II. Bot. Rev. 52:1-112.

_________, and J.H. Graham. 1960. Helminthosporium dictyoides and related species on forage grasses. Phytopathology 50:691-695.

Weibull, Peter. 1978a. Descriptions of grass diseases No. 1. Weibull's Gras-tips 21:3-4.

Brown blight

caused by Drechslera siccans.

Brown blight is a common foliar disease on annual and perennial ryegrasses in the eastern United States and in Europe. In Ireland, the disease is most preva­lent on ryegrasses and meadow fescue. D. siccans is fairly common in the Nordic countries.

Symptoms

Symptoms caused by D. siccans resemble those incited by D. dictyoides. However, characteristic net markings are either very faint or lacking on the leaves. The fungus produces numerous dark brown spots, of various shapes, which may coalesce to form large mottled discolored areas. More mature lesions show ash-gray centers and a yellowing of surround­ing tissue (O'Rourke, 1976). If the infection is severe, the leaves become yellow at the tip. The blade and sheath gradually wither and die (Kreitlow et al., 1953).

Host Range

Brown blight, common on annual and perennial ryegrasses, is less common on meadow fescue and orchardgrass.

Control

Several perennial ryegrass cultivars have been developed specifically for resistance to D. siccans (Braverman, 1986).

Literature

Braverman, S.W. 1986. Disease resistance in cool sea­son forage, range and turf grasses II. Bot. Rev. 52:1-112.

Kreitlow, K.W., J.H. Graham, and R.J. Garber. 1953. Diseases of forage grasses and legumes in the northeastern states. Pa. Agric. Exp. Stn. Bull. 573.

O'Rourke, C.J. 1976. Diseases of grasses and forage legumes in Ireland. An Foras Taluntais. Oak Park Res. Centre. Carlow. 115 p.

Leafspot

caused by Drechslera catenaria.

Drechslera leafspot is not as common as tawny blotch (Stagonospora foliicola (Bres.) Bubak), but it is destruc­tive and capable of severely damaging reed canarygrass stands.

Symptoms

The initial foliar symptoms appear as tiny, dark green spots surrounded by a chlorotic halo. Mature lesions are elongated and reddish­ brown, often with light-buff-colored centers. As lesions increase in size, they often coalesce to form large areas of necrotic tissue on susceptible clones (Zeiders, 1976).

Etiology

There is evidence suggesting that patho­genesis by D. catenaria is accomplished primarily by secretion of a toxic substance(s) which precedes my­celial colonization of the tissue. The fungus over­ winters as mycelium in tissue infected the previous year. D. catenarium may also be seed-borne.

Host Range

The fungus is common on reed canarygrass, woodreed (Cinna arundinacea L.), and to a lesser extent on orchardgrass and bentgrass.

Control

Reed canarygrass genotypes with good re­sistance to D. catenaria exist (Zeiders, 1976). These clones could be used in breeding programs designed to increase disease resistance.

Literature

Zeiders, K.E. 1976. A new disease of reed canary­grass caused by Helminthosporium catenarium. Plant Dis. Rep. 60:556-560.

Leafspot/Blotch

caused by Drechslera dactylidis.

This disease may occur on orchardgrass from May to October in the northeastern and north central United States, and sometimes causes serious damage.

Symptoms

In the field, the disease appears as mar­ginal, light brown, irregular leaf lesions up to 15 mm long and 5 mm wide. Scattered conidiophores de­velop on the necrotic tissue, and black perithecia of Pleospora phaeocomes form underneath the epidermis (Graham, 1955a). In artificial inoculations, young le­sions develop as yellow to orange, round or oblong areas 1 to 4 mm in diameter. When infection is heavy, mature lesions may coalesce and envelop the entire leaf. The causal fungus is unique in that it regularly produces a certain proportion of curved, branched, or three-pointed spores in culture and on infected tis­sue (Zeiders, 1980). This characteristic is important in the laboratory diagnosis of the disease.

Host Range

D. dactylidis has a very narrow host range among forage grasses. Orchardgrass is the most important host.

Control

Disease severity can be reduced by crop rotation, timely (early) cutting to prevent a build-up of inoculum, and development of resistant cultivars.

literature

Graham, J.H. 1955a. A disease of orchardgrass caused by Pleospora phaeocomes. Phytopathology 45:633-634.

Zeiders, K.E. 1980. A variable-spored isolate of Drechslera dactylidis pathogenic on orchardgrass and corn. Plant Dis. 64:211-213.

Drechslera leaf Streak

caused by Drechslera phlei (Graham) Shoem.

This leaf streak disease of timothy was first observed in central Pennsylvania and subsequently was re­ported to occur throughout the northeastern United States (Graham, 1955b).

Symptoms

The disease is characterized by irregular light brown, necrotic streaks with conspicuous chlorotic borders. The necrotic areas, which are 1 to 5 mm wide, are often marginal and may extend the entire length of the leaf blade (Graham, 1955b). Often, le­sions may coalesce, thereby causing a browning of most of the leaf. If the chlorotic symptom is lacking, the disease is difficult to distinguish from leaf streak. However, under moist conditions, the leaf streak dis­ease is characterized by parallel rows of dark tufts of conidiophores of the causal fungus on infected leaves.

Host Range

The fungus is limited to timothy and other Phleum spp.

Control

Resistant cultivars offer a means of reducing severity of Drechslera leaf streak of timothy (Braverman, 1986).

literature

Braverman, S.W. 1986. Disease resistance in cool sea­son forage, range and turf grasses II. Bot. Rev. 52:1-112.

Graham, J.H. 1955b. Helminthosporium leaf streak of timothy. Phytopathology 45:227-228.

Melting Out, leafspot, and Foot Rot

caused by Drechslera poae.

This fungus causes a leafspot and foot rot of Kentucky bluegrass, and according to Kreitlow et al. (1953), melting out is probably the most prevalent and serious disease of the host in the northeastern part of the United States. D. poae is also common on many Poa spp. in northern Europe and is considered to be the most important pathogen on Kentucky bluegrass. Damage from the disease is enhanced by frequent cuttings of the grass.

Symptoms

The disease appears as well defined purplish-black to reddish-brown oval leaf spots with a chocolate-brown to purple-brown margin and a light center (Weibull, 1979a). Lesions often form in rows along the margin of the leaf. Some lesions ex­tend the width of the leaf blade, causing it to break over or wither from the tip downward to the lesion. The symptoms frequently extend to the inflores­cence, and lesions may coalesce to form a large necrotic area extending across the entire leaf to the sheath and stem base. This results in weakening and death of the plant. Frequently, the immature inflores­cence in the boot is diseased, with the flower heads inside being attacked.

Host Range

D. poae is confined to species of Poa.

Control

High clipping or mowing (1 1/2 inches or more) reduces the incidence of melting out by main­taining the less susceptible older leaves without stimulating succulent new shoots from the crown (Kreitlow et al., 1953). Several cultivars have been de­veloped for resistance to D. poae, as reviewed by Braverman (1986).

Literature

Braverman, S.W. 1986. Disease resistance in cool sea­son forage, range and turf grasses II. Bot. Rev. 52:1-112.

Kreitlow, K.W., J.H. Graham, and R.J. Garber. 1953. Diseases of forage grasses and legumes in the northeastern states. Pa. Agric. Exp. Stn. Bull. 573.

Weibull, Peter. 1979a. Descriptions of grass diseases No. 7. Weibull's Gras-tips. 22:unpaginated.

Bipolaris Foot Rot, Leaf Blight, and Seedling Blight

caused by Bipolaris sorokinianum (sexual stage: Cochliobolus sativus).

The disease incitant causes a foot rot, leaf blight, and seedling blight on a wide range of grasses in North America. While the pathogen is of significant eco­nomic importance on wheat, barley, and oats, it is ca­pable of causing moderate to severe damage on cool-­ and warm-season grasses.

Symptoms

The predominant symptom on grasses is either a leafspot or leaf blight. Lesions caused by B. sorokinianum are elongated, dark to purple, and mea­ sure 2 to 5 mm. Older lesions may become light­ colored in the center. As disease severity increases with time, lesions often coalesce, causing foliage to become blighted. This reduces forage quality. The disease is most severe on switchgrass during the warmest part of the summer.

Host Range

The fungus attacks a multitude of the Gramineae as noted by Sprague (1950).

Control

Resistant cultivars offer the best means of re­ducing the severity of B. sorokinianum (Braverman, 1967, 1986).

Literature

Braverman, S.W. 1967. Disease resistance in cool sea­son forage, range and turf grasses. Bot. Rev. 33:329-378.

_________. 1986. Disease resistance in cool season for­ age, range and turf grasses II. Bot. Rev. 52:1-112.

Sprague, R. 1950. Diseases of Cereals and Grasses in North America. Ronald Press Co. New York. 538 p.

Spot Blotch of Switchgrass

caused by Bipolaris sorokinianum (Helminthosporium sativum).

Spot blotch, causing moderate to severe damage, was the most prevalent disease on leaves of switchgrass in Pennsylvania during a 5-year study (Zeiders, 1984).

Symptoms

Lesions caused by B. sorokinianum are elongated (1 to 1.5 by 3 to 5 mm), dark to purple. Older lesions may become light-colored in the center. As disease severity increases, lesions often coalesce and leaves become blighted, reducing quality of the forage. The early symptoms of spot blotch, tiny dark elongated spots, begin to appear in mid to late June. The disease often becomes severe from mid-July through September.

Etiology

Spores (conidia) of B. sorokinianum are pro­duced on dead infected leaf tissue and disseminated by air currents to new leaves, causing new infections. New disease lesions can be seen about 48 hours after inoculation. The severity of spot blotch is related to the duration of high relative humidity and leaf wetness.

Control

Resistant cultivars appear to be the best means of reducing the severity of spotblotch. Zeiders (1984) noted wide variability in reaction of 11 switchgrass varieties and strains to spot blotch. Dam­age ranged from mild to severe. The Ky-729 strain showed the most resistance, and three other strains were moderately resistant. These differences provide evidence of genetic variability for resistance to spot blotch.

Literature

Zeiders, K.E. 1984. Helminthosporium spot blotch of switchgrass in Pennsylvania. Plant Dis. 68:120-122.

Eyespot

caused by Heterosporium phlei Gregory.

Figure 18. Eyespot, Heterosporium phlei, on timothy.

Eyespot, a common foliar disease of timothy, was first described in New York State and since has been reported throughout North America, Europe, and Japan. The disease probably occurs wherever the crop is grown. Roberts et al. (1955) described the de­structive nature of eyespot to timothy in New York. Sakuma and Marita (1961) reported that eyespot re­duced leaf crude protein by 26 percent.

Symptoms

Heterosporium phlei occurs on both leaf surfaces as small, light-colored, oval lesions with pale grayish-tan centers surrounded by a narrow purple border, which eventually fades to brown. The disease may also be referred to as purple spot (O'Rourke, 1976). When lesions are abundant, the intervening tissue frequently becomes yellow; affected leaves turn brown and wither prematurely (Kreitlow et al., 1953).

Etiology

The fungus probably overwinters in dead infected leaf tissue. Although eyespot is more preva­lent during the summer months, it may be found on green tissue during the spring and early autumn months. Primary infection in the spring probably oc­curs from conidia disseminated by wind and rain. However, it is difficult to find spores on diseased leaves in the field (Kreitlow et al., 1953). Conidia ger­minate over a wide temperature range, which may account for the occurrence of H. phlei on leaf tissue over a long growing season.

Host Range

Eyespot is confined to Phleum species. In Europe, it is found on timothy and P. bertolonii DC.; in North America the host range includes alpine tim­othy (P. alpinum L. ( = P. pratense)), P. phleoides (L.) Karst., and P. nodosum L. as well.

Control

Resistant cultivars, noted by Smith (1970) and Hanson (1972), offer some control of H. phlei.

Literature

Hanson, A.A. 1972. Grass varieties in the United States. USDA Agric. Handb. No. 170. 124 p.

Kreitlow, K.W., J.H. Graham, and R.J. Garber. 1953. Diseases of forage grasses and legumes in the northeastern states. Pa. Agric. Exp. Stn. Bull. 573.

O'Rourke, C.J. 1976. Diseases of forage grasses and legumes in Ireland. An Foras Tulantis. Oak Park Res. Centre. Carlow. 115 p.

Roberts, D.A., R.T. Sherwood, K.D. Fezer, and C.S. Ramamurthi. 1955. Diseases of Forage Crops in New York, 1954. Plant Dis. Rep. 39:316-317.

Sakuma, T., and T. Marita. 1961. Heterosporium leafspot of timothy and its causal fungus, Heterosporium phlei Gregory. Bull. Hokkaido Agric. Exp. Stn. 7:77-90.

Smith, J.D. 1970. Resistance of timothy cultivars to Heterosporium phlei, Drechslera phlei and frost injury. Can. J. Plant Dis. Sur. 50:95-98.

Stagonospora Diseases of Forage Grasses

Stagonospora species cause diseases on a number of forage grasses. Important diseases are purple leaf­ spot of orchardgrass and tawny blotch of reed canarygrass. These two diseases and a less important disease of smooth bromegrass are described in some detail.

Purple Leafspot of Orchardgrass

caused by Stagonospora arenaria Sacc.

Purple leafspot of orchardgrass occurs throughout the eastern United States. Lesions appear on leaves as soon as new growth begins and continue to de­velop in the summer and fall except during periods of prolonged hot, dry weather. The disease reaches its peak either shortly before or at time of heading. Leaves wither and turn brown when heavily attacked by the fungus, reducing the nutritive value of the for­age. Infections progress rapidly in the fall and new le­sions can be found until late November.

Symptoms

The lesions usually appear as small, somewhat elongate, blackish-brown to deep purple spots. When lesions are abundant they coalesce, causing the leaf to turn brown and die. Frequently the browning develops at the tip of a leaf or along the margin in long brown streaks. Small golden-brown bodies, the pycnidia of the fungus, develop within the dead areas of a leaf.

Etiology

Infection occurs from spores that over­ winter in pycnidia in dead stems and leaves. Upon emerging from the pycnidia they are disseminated by spattering rain or in wind-blown fragments of dead plants. Pycnidia usually form in rows in the dead parts of leaves.

Host Range

A list of susceptible species has been compiled by Sprague (1950).

Control

Breeding for resistance is a practical means of reducing disease severity. Although immune plants have not been found, resistance to purple leaf­ spot can be effectively increased by inoculation and selection in the greenhouse (Zeiders et al., 1984).

Literature

Sprague, R. 1950. Diseases of Cereals and Grasses in North America. Ronald Press Co. New York. 538 p.

Zeiders, K.E., C.C. Berg, and R.T. Sherwood. 1984. Effect of recurrent phenotypic selection on resis­tance to purple leafspot in orchardgrass. Crop Sci. 24:182-185.

Leafspot of Smooth Bromegrass

caused by Stagonospora bromi A.L. Sm. & Ramsb.

Leafspot caused by S. bromi does not achieve eco­nomic importance in all years, but it is reported to be a destructive parasite of smooth bromegrass and other species.

Symptoms

Leafspots on smooth bromegrass are dark brown, elongated, and often pointed at the ends. The inner portions become light tan as the le­sion expands. Pycnidia develop within the infected tissue. Although similar in appearance to brown spot caused by Drechslera bromi, lesions are less angular in outline and may be distinguished by the light central area and the pycnidia in the infected tissue (Zeiders and Graham, 1962).

Etiology

The etiology of S. bromi is similar to that of S. arenaria.

Host Range

The reported hosts are smooth bromegrass and Bromus species (Sprague, 1950).

Control

In screening tests at the U.S. Regional Pas­ture Research Laboratory, selections of smooth bromegrass varied from very susceptible to resistant in reaction to S. bromi (Zeiders and Graham, 1962).

Literature

Sprague, R. 1950. Diseases of Cereals and Grasses in North America. Ronald Press Co. New York. 538 p.

Zeiders, K E., and J.H. Graham. 1962. Stagonospora bromi on Bromus inermis. Plant Dis. Rep. 46:729-731.

Tawny Blotch

caused by Stagonospora foliicola.

Tawny blotch is probably the most important disease of reed canarygrass in the temperate zones of the world. Reed canarygrass is often grown as a cover crop on areas that are spray-irrigated with effluent from sewage treatment plants.

Symptoms

Young lesions are small, elongated, and purple; as they enlarge they become dark brown to reddish-brown, usually with pointed ends and buff centers. If numerous, the lesions often coalesce to form large areas of dead tissue. Lesions are present on infected plants throughout the growing season. Leaves, leaf sheaths, and culms may be attacked by S. foliicola (Zeiders, 1975).

Etiology

In the spring, pycnidia of S. foliicola develop on diseased leaves infected the previous year. The erumpent pycnidia are scattered over the dead leaves and are not confined to the diseased tissue. Spores are released by rupture or disintegration of the pycnidial wall. These spores comprise the inoculum that causes new infections in the spring. They are probably disseminated from plant to plant by rain and wind-blown fragments of dead plant parts.

Host Range

The fungus is confined to Phalaris spp.

Control

Disease severity can be significantly reduced by mowing the stand three times a year, preventing build-up of disease. Inoculation experiments and field evaluations have revealed strong resistance to tawny blotch in some genotypes. It should be possi­ble to develop varieties with good resistance to this disease (Zeiders and Sherwood, 1977a, b).

Literature

Zeiders, K.E. 1975. Stagonospora foliicola, a pathogen of reed canarygrass spray-irrigated with municipal sewage effluent. Plant Dis. Rep. 59:779-783.

________, and R.T. Sherwood. 1977a. Effect of irriga­tion with sewage wastewater, cutting manage­ment, and genotype on tawny blotch of reed canarygrass. Crop Sci. 17:594-596.

________, and 1977b. Reaction of reed canarygrass genotypes to the leafspot pathogens Stagonospora foliicola and Helminthosporium catenarium. Crop Sci. 17:651-653.

Anthracnose of Grasses

caused by Colletotrichum graminicola (Ces.) Wils.

Anthracnose, one of the most common and widely distributed diseases of forage grasses, is particularly noticeable in mid-summer to early fall. Anthracnose occurs on practically all of the cultivated warm- and cool-season forage grasses in humid areas of the northern United States. Sudangrass is attacked in mid-summer at the height of vegetative growth. C. graminicola is primarily a high temperature organism (optimum 28C), which accounts for its association with maturity of grass in mid-summer (Kreitlow et al., 1953). Bruehl and Dickson (1950) found some host specificity among isolates. In general, isolates from warm-season grasses, such as Sorghum spp., were more pathogenic at higher temperatures. If con­ditions are favorable for development of the disease, the fungus causes stunting, wilting, and sometimes death of seedlings. Although relatively few seedlings may be killed, the root system is usually damaged and yield is reduced, even though plants appear to recover as the season progresses. In older plants, the culm or leaf sheath is attacked, and the fungus may spread into the crown and roots. Perennial grasses, so infected, frequently die out in the second or third years. Death of plants occurs more rapidly in areas of low soil fertility. Early attacks cause a general reduc­tion in vigor, shriveling of seed, premature ripening, or death of the plant (Kreitlow et al., 1953).

Symptoms

Lesions that develop on the sheath or stem are usually light tan with a darker border of red or brown. Black acervulii of the fungus usually de­velop within the bleached center of a lesion or on leaf blades of dead plants, particularly when moisture is plentiful.

Etiology

Infection occurs from diseased seed or from spores and mycelia that develop saprophytically on old crop residues. Infection in mature culms is com­mon in the vicinity of nodes. After the pathogen is es­tablished and lesions develop, secondary spread oc­curs from spores or mycelia.

Host Range

A lengthy list of Gramineous hosts has been compiled by Sprague (1950).

Control

General control measures include crop rota­tion, particularly to avoid a sequence of closely re­lated crops; timely cutting or grazing; maintaining soil fertility; plowing under all plant residues; and de­velopment of resistant varieties (Kreitlow et al., 1953).

Literature

Bruehl, G.W., and J.G. Dickson. 1950. Anthracnose of cereals and grasses. U.S. Dep. Agric. Tech. Bull. No. 1005. 37 p.

Kreitlow, K.W., J.H. Graham, and R.J. Garber. 1953. Diseases of forage grasses and legumes in the northeastern states. Pa. Agric. Exp. Stn. Bull. 573.

Sprague, R. 1950. Diseases of Cereals and Grasses in North America. Ronald Press Co. New York. 538 p.

Septoria Leafspots of Forage Grasses

Numerous species of Septoria Fr. em. Sacc. cause leaf blotch and leafspots on grasses and cereals through­out the temperate and subtropical zones. These irreg­ular blotches are straw-colored to brown, with dark brown to black pycnidia occurring in the older portions of the lesions. The blotches are often similar to those produced by Stagonospora or Ascochyta; mor­phological distinctions between the three genera on the grasses are not clearly defined. Septoria species are generally more aggressively parasitic on grasses than are those of closely related genera. Infected grasses may be the source of Septoria inoculum from which cereal crops become infected.

In Septoria, pycnidia are subepidermal, slightly erumpent, and are formed in the older portions of the blotches or spots on the leaves, culms, and inflores­cences of the host. The pycnidia are lens-shaped, brown to black, ostiolate, and parenchymatous.

The etiologies of the Septoria diseases discussed here are basically similar. Spores overwinter in pyc­nidia on dead leaves and are dispersed via wind or rain, and infect healthy plants. Germination will occur over a wide temperature range. Under moist conditions, spores are extruded in a gelatinous mass from the ostiole and are disseminated to adjacent healthy leaves by rain and wind-borne infected plant parts.

Leafspot of Bromus Species

caused by Septoria bromi Sacc.

S. bromi is widely distributed on Bromus species and may cause moderate to severe damage on smooth bromegrass.

Symptoms

Leafspots are elongate to elliptical, tan to brown, with infected leaves often yellow, eventually becoming brown (Sprague, 1950). The disease occurs early in spring, decreases in prevalence during sum­mer, and reappears again in autumn.

Control

Some reduction in disease incidence may be achieved by removing or plowing-under plant residues.

Literature

Sprague, R. 1950. Diseases of Cereals and Grasses in North America. Ronald Press. New York. 538 p.

Speckled Leaf Blotch

caused by Septoria tritici var. lolicola Sprague & A.G. Johnson

Although Septoria species are common on many grasses and occur worldwide, damage caused by this pathogen is most severe on the ryegrasses.

Symptoms

The earliest symptoms usually appear as diffuse oval lesions along the leaf margins. Pycnidia are prominent and numerous within the lesions, usu­ally aligned in rows parallel to the leaf veins. They are most numerous on the upper surface of the leaf (O'Rourke, 1976). Pycnidia are globose to sub­globose and are light brown.

Host Range

The fungus is restricted to species of Lolium.

Control

There are no reports of cultivars with resis­tance to S. tritici var. lolicola.

Literature

O'Rourke, C.J. 1976. Diseases of grasses and forage legumes in Ireland. An Foras Taluntais. Oak Park Res. Centre. Carlow. 115 p.

Leafspot/Blotch

caused by Septoria bromi (Sacc.) var. phalaricola Sprague.

This disease is not as common or prevalent on reed canarygrass as is tawny blotch, but has the potential to cause serious damage. The pathogen is confined almost entirely to reed canarygrass.

Symptoms

The young lesions are narrow, water­ soaked or dark gray spots up to 4 mm in length, surrounded by a yellow halo. Mature lesions are buff­ colored, 2 to 4 mm long, and often confined between veins of the leaf. Lesions may coalesce to form large buff-colored necrotic areas. The disease may occur from May to October.

Etiology

Pycnidiospores, produced in globose or bulb-shaped pycnidia, are disseminated by wind and wind-driven rain to adjacent plants.

Host Range

The pathogen is confined to reed canarygrass.

Control

Zeiders (1979) identified genotypes with moderate resistance to S. bromi var. phalaricola that could be used in breeding resistance into a new cultivar.

Literature

Zeiders, K E. 1979. A Septoria disease of reed canarygrass in Pennsylvania. Plant Dis. Rep. 63:796-800.

Ascochyta Leafspot

caused by Ascochyta sorghi Sacc. (Syn. A. graminicola Sacc.).

A. sorghi is probably the most widely distributed of several Ascochyta species that cause leafspot diseases on forage grasses.

Symptoms

A. sorghi attacks a wide range of the Gra­mineae and causes brownish-tan lesions, up to 20 mm in length, with brown to reddish margins on leaves and culms. Light brown lens-shaped-to­ globose ostiolate pycnidia form in groups within the leaf lesions and then become erumpent.

Etiology

The fungus overwinters as mycelium in dis­eased plant tissue. In late spring and early summer, globose pycnidia form in infected tissue. Pycnidiospores are extruded, disseminated by wind and wind-blown rain, and infect other plants.

Host Range

The host range of A. sorghi has been re­corded by Sprague (1950).

Control

The tetraploid 'Sabrina,' Lolium x hyuridum Hausskn., has resistance to A. sorghi (Braverman, 1986).

Literature

Braverman, S.W. 1986.Disease resistance in cool sea­son forage, range and turf grasses II. Bot. Rev. 52:1-112.

Sprague, R. 1950. Diseases of Cereals and Grasses in North America. Ronald Press. New York. 538p.

Leafspot

caused by Ascochyta brachypodii (perfect state: Didymella sp.).

Ascochyta leafspot was the most important disease on three perennial warm-season grasses (big bluestem, little bluestem, and indiangrass) in Penn­ sylvania over a five-year period (Zeiders, 1982). The disease also occurs on these and several other species in the western United States.

Symptoms

The disease, caused by A. brachypodii, is characterized by reddish-brown elongated spots or blotches with tan centers, 2 to 6 by 1 to 1.5mm in di­mension, on leaves and leaf sheaths. In advanced stages, symptoms are sometimes streak-like.

Etiology

Symptoms of the disease caused by A. brachypodii usually appear in mid to late June. Infection increases in severity until about October 1 on grasses not cut or grazed. The fungus overwinters as mycelium in infected leaves. In late spring and sum­mer, brown, globose, beaked pycnidia develop in this tissue. Spores (conidia) of A. brachypodii are produced in these fruiting bodies and then extruded through the pycnidial beak in a viscous mass. Other plants are infected by spores disseminated by wind and wind-blown rain. Zeiders (1982) found the sever­ity of Ascochyta leafspot to be related to the duration of high relative humidity and leaf wetness in the growing area.

Host Range

Additional hosts of A. brachypodii in­clude maize, sudangrass, and oats (Zeiders, 1982).

Control

The severity of the disease can be reduced by cutting or grazing the grasses at least twice per growing season to prevent disease build-up. Differ­ences in susceptibility of big bluestem varieties to A. brachypodii in the field have been observed (Zeiders, 1982). In Pennsylvania, severity of leafspot on the cultivar NY-1145 was consistently less severe than on cultivars Kaw or Pawnee (Zeiders, 1982).

Literature

Zeiders, K.E. 1982. Leaf spots of big bluestem, little bluestem, and indiangrass caused by Ascochyta brachypodii. Plant Dis.66:502-505.

Leafspot of Smooth Bromegrass

caused by Selenophoma bromigena (Sacc.) Sprague & A.G. Johnson.

Figure 19. Selenophoma leafspot, Selenophoma donacis, on timothy.

Selenophoma leafspot is an important disease that is general in distribution, appearing on smooth brome­ grass wherever it is grown (Allison, 1945). Severe leaf infection may almost completely defoliate the plant. S. donacis (Pass.) Sprague & A.G. Johnson is an im­portant pathogen of timothy.

Symptoms

Soon after new spring growth, chlorotic oblong lesions 8 to 15 mm in length appear on the lower leaves of infected plants. After about 7 days, the light-brown lesions have a light-red border and are dry at the center. After about 12 to 14 days, nu­merous pycnidia appear in these dried areas. The in­fected leaves tum yellow and often die. Almost total defoliation of severely infected plants is common. Or­dinarily, infection is confined to localized spots on the leaves, but if the environment is cool and damp, infected areas may coalesce and cover large portions of the leaf surface. At such times the pathogen often spreads to the sheath, stem, rachis, panicle, and glumes; severe attacks stunt growth and may kill the plants.

Etiology

Presumably, the fungus overwinters as pycnidia in infected tissue. Pycnidiospores ooze from the pycnidium during the spring and are spread to adjacent plants by wind-driven rain. Penetration of the host tissues is direct.

Host Range

Smooth bromegrass, mountain brome­ grass, fringed brome (B. ciliatus L.), Canada brome (B. purgans L.), and B. arvensis L. are hosts (Sprague, 1950).

Control

Smooth bromegrass plants resistant to S. bromigena are reported to be common; thus, selection for resistance and breeding offers a means of reduc­ing disease severity. Specialized races of S. bromigena have also been observed.

Literature

Allison, J.L. 1945. Selenophoma bromigena leaf spot on Bromus inermis. Phytopathology 35:233-240.

Sprague, R. 1950. Diseases of Cereals and Grasses in North America. Ronald Press Co. New York. 538 p.

Rhizoctonia Blight

caused by Rhizoctonia solani Kuehn (perfect state: Thanatephorus cucumeris (Frank) Dank).

Rhizoctonia solani, a soil-borne organism, is particu­larly abundant in acid soils. The pathogen has a wide host range among the forage grasses and is severe on smooth bromegrass, orchardgrass, and tall fescue. It is important also in subtropical climates. The fungus may attack all parts of the plant-roots, crowns, leaves, and stems-but damage usually becomes most evident after stands are well established. The fungus causes damping-off and root rots of many grasses and is most destructive on seedlings or older weakened plants. In addition to causing damping-off and root and crown rots, R. solani causes a foliar blight that is extremely destructive to dense stands during hot, humid weather.

Symptoms

Large, irregular, tan-to-whitish blotches with narrow, dark brown borders form on leaves and leaf sheaths. The blotches often coalesce and girdle the stems. Under warm, moist conditions in dense stands, the fungus spreads rapidly by leaf-to-leaf contact, producing large patches of grayish matted dead tissue covered with a web of mycelium. Even so, the crowns of mature plants often survive.

Etiology

R. solani produces small, tan to brown scle­rotia on dead infected tissues, enabling the fungus to survive for many years in the soil. The sexual stage of the fungus, Thanatephorus cucumeris (Frank) Donk, has been reported to occur on tall fescue as effuse white patches of mycelium, usually on the lower sur­face of the leaf (Luttrell, 1962).

Host Range

Sprague (1950) has compiled an exten­sive list of gramineous hosts of the fungus.

Control

The pathogen is soil-borne, attacks a wide range of hosts, and persists in the soil indefinitely. Timely cutting or grazing of the grass is effective in curtailing the spread of Rhizoctonia blight, since its development is retarded when exposed to direct sun­ light and good air drainage. Crop rotation is ineffect­ive. Parasitic races of R. solani exist, but attempts to identify germplasm of forage grasses with good field resistance have met with little success (Luttrell, 1962).

Literature

Luttrell, E.S. 1962. Rhizoctonia blight of tall fescue grass. Plant Dis. Rep. 46:661-664.

Sprague, R. 1950. Diseases of Cereals and Grasses in North America. Ronald Press Co. New York. 538 p.

Take-all

caused by Gaeumannomyces (Ophiobolus) graminis (Sacc.) Arx & Oliver.

Figure 20. Take-all. Gaeumannomyces gramlnis var. avenae, on perennial ryegrass.

Gaeumannomyces graminis, the causal agent of the take-all and white diseases of cereals, parasitizes a large number of forage and turf grasses as well. Take­-all disease is widely distributed in western Europe, Ireland, and Great Britain. In the United States, the disease occurs mainly in the Pacific Northwest (O'Rourke, 1976).

Symptoms

The fungus attacks the roots, crown, and basal culm tissues of cereals and grasses. Infection is not as prominent in the grasses. Under moist condi­tions leaf color fades and bleaching of leaves and culms follows. The main roots, crown, and culm (par­ticularly the tissue between the leaf sheath and culm base), show a distinct rot and dark runner hyphae that form a mat of thick-walled coarse mycelia. Under dryer conditions, plant tillering is reduced and the mycelial mat is less pronounced.

Etiology

The pathogen is soil-borne in a rather direct association with culms and roots of grasses. Infection occurs from mycelia in the crop residue; the mycelia invade the roots, culm, and sheath. Slender infection hyphae grow from the runner hyphae into the root tissues as far as the stele (O'Rourke, 1976). Perithecia form on the dark fungal mats between the leaf sheaths and the culm base. In a moist environment, asci subsequently develop and mature ascospores are discharged and initiate new infection sites. Grasses are important economic hosts of G. graminis, as they provide the inoculum sources for the infection of ce­real hosts. Each is required for the continued spread and survival of the pathogen.

Host Range

In Great Britain and Ireland, the fungus is prevalent on Agrostis, Agropyron, Festuca, Lolium, and Poa spp. (O'Rourke, 1976). While G. graminis readily attacks wheat, barley, and rye, oats are nearly immune. However, oats are very susceptible to G. graminis var. avenae Turner. Brooks (1965) found that pathogen survival was optimum on orchardgrass, red fescue, and annual and perennial ryegrasses. He also noted survival on Arrhenatherum sp. and Agrostis sp.

Control

Brooks (1965) noted that tall oatgrass was the least susceptible of 15 grasses when inoculated artificially with G. graminis, but the most susceptible when the inoculum source was G. graminis var. avenae. Nilsson (1969) reported that orchardgrass showed resistance to G. graminis var. avenae, but was susceptible to G. graminis. Perennial ryegrass was re­ sistant to G. graminis. In general, specific resistance to take-all disease by economically important grasses will vary within a country and from country to country.

Nilsson (1969) noted that perennial ryegrass was resistant to take-all, but documented conflicting re­ports of resistance to the disease in other Lolium spp. However, Walker (1975) reviewed the take-all dis­eases of the Poaceae and indicated that Lolium spp. were resistant.

Nitrogen in ammonia compounds in a slightly acid soil appears to reduce the incidence of take-all dis­ease (O'Rourke, 1976).

Literature

Brooks, D.H. 1965. Wild and cultivated grasses as carriers of the take-all fungus (Ophiobolus graminis). Ann. Appl. Biol. 57:307-316.

Nilsson, H.E. 1969. Studies on the foot and foot rot diseases of cereals and grasses I. On the resistance to Ophiobolus graminis Sacc. Ann. Agric. Col. Swe. 35:275-807.

O'Rourke, C.J. 1976. Diseases of grasses and forage legumes in Ireland. An Foras Taluntais. Oak Park Res. Centre. Carlow. 115 p.

Walker, J. 1975. Take-all diseases of Gramineae: A re­view of recent work. Rev. Plant Pathol. 54:113-144.

Red Thread

caused by Corticium fuciforme (Berk.) Wakef.

Figure 21. Red thread, Corticium fuciforme, on perennial ryegrass.

Red thread, or pink patch, is common on turfgrass species in areas of low soil fertility. The disease is par­ticularly noticeable on red fescue and perennial ryegrass.

Symptoms

The disease first appears as small irregu­lar reddish or pinkish patches of infected plants; the infection may expand to several meters. The pinkish discoloration is due to (i) anthocyanins produced in affected grass foliage and (ii) to a pink, slimy mycelial growth. The fungus produces pink stromata on dried-out leaves which, in turn, fade to a tan to red­dish color.

Etiology

In dry weather, stromata break off from the diseased tissue. Stromata may survive under this condition up to two years, and can be wind-blown, disseminating the pathogen to initiate new areas of infection. Basidia are produced on infected host tis­sues and on the stromata. Fungal spread within a diseased area is by mycelial growth from plant to plant. According to Couch (1973), basidiospores are not im­portant in the spread of this fungus. The disease is most evident in autumn, particularly during periods of low temperature and high humidity.

Host Range

In Britain, the disease is widespread on red fescue, Agrostis spp., perennial ryegrass, and an­nual meadowgrass (Poa annua L.) (O'Rourke, 1976).

Control

Red thread is prevalent in soils of low fertil­ity, thus application of nitrogenous fertilizers reduces disease incidence (O'Rourke, 1976). However, exces­sive nitrogen fertilization may enhance Fusarium patch disease. Several cultivars of perennial ryegrass have been tested for resistance to C. fuciforme (Braver­man, 1986).

Literature

Braverman, S.W. 1986. Disease resistance in cool sea­son forage, range and turf grasses IL Bot. Rev. 52:1-112.

Couch, H.B. 1973. Diseases of Turfgrasses. Reinhold Publishing Corp. New York. 348 p.

O'Rourke, C.J. 1976. Diseases of grasses and forage legumes in Ireland. An Foras Taluntais. Oak Park Res. Centre. Carlow. 115 p.

Snow Molds

caused by Typhula incarnata Lasch. ex Fr., T. ishikariensis Imai, T. itoana Imai, Fusarium nivale (Fr.) Ces, and Sclerotinia borealis (Bubak & Vleugel) Kohn.

Several species of Typhula, Fusarium, and Sclerotinia are the causal agents of a ubiquitous disease in tem­perate, cool regions of the world. The disease is quite severe in regions where winters are cold and snow cover predominates. F. nivale usually requires a snow cover of at least 60 days; T. incarnata needs 90 days; T. ishikariensis requires about 120 days, and severe dam­age results with a snow cover in excess of 150 days. S. borealis needs at least 180 days' snow cover to cause extensive damage. The disease caused by F. nivale and T. incarnata can also occur without snow cover (Weibull, 1979b, 1983).

Symptoms

Fusarium patch caused by F. nivale (Mi­cronectriella nivalis (Schaffn.) Booth) appears on snow­ free areas as small, circular water-soaked patches, be­coming yellow to orange-brown. The small patches often coalesce, and under moist conditions a fringe of pale pink mycelium is visible; this mycelium tends to mat down the aerial plant parts. The disease prog­resses under prolonged snow cover, with new infec­tions showing an abundant aerial mycelium which may tum pink, orange, or beige if exposed to light.

At times, minute gelatinous masses of conidia may be seen on the matted leaves. The dead tissue, after dry­ing, forms a thick pink mat, providing the name "pink snow mold." The pink color distinguishes this disease from other low-temperature maladies, such as those caused by Typhula spp. and Sclerotinia sp. (Weibull, 1979b).

Typhula blight or gray or speckled snow mold, caused by T. incarnata, is widespread and may se­verely damage grasses. Although fungal attacks will occur on grasses without snow cover, damage is most severe after at least 90 days of cover. The disease first appears as small patches, 2 to 5 cm in diameter, which at first light are yellowish-brown to a grayish­ white. Under snow, the diseased patches coalesce to a considerable size. Initially, a white-grayish myce­lium develops on the grass substrate. Pink to pinkish­ orange followed by reddish-brown to dark brown sub-globose sclerotia develop on or within infected tissue (Weibull, 1979b).

Gray or speckled snow mold, also called Typhula blight, is caused by T. ishikariensis and is a major dis­ease on overwintering grasses with prolonged snow cover. Much more virulent than T. incarnatum (Wei­ bull, 1979b), T. ishikariensis becomes a major incitant of snow mold on grasses covered 150 days or longer. Symptoms of this snow mold are bleached patches of dead or dying plants sparsely covered with a grayish­ white mycelium. The decaying tissue is speckled with numerous dark, globose to slightly flattened sclerotia.

A third causal agent of the Typhula speckled snow mold or snow scald is T. itoana. The fungus is preva­lent on grasses snow-covered during winter and early spring. As the snow melts, those grasses at­tacked by the fungus appear as flattened mats of bleached dead foliage and mycelium. Numerous brown sclerotia develop on the dead leaves and grass stems. T. itoana is apparently the most widely distrib­uted of the three incitants of Typhula snow molds.

Sclerotinia snow mold is also a major disease on overwintering grasses. It is restricted to those regions of extreme cold where snow cover lasts for at least 180 days. The disease causes severe damage to grasses. Symptoms first appear, after snow melt, as bleached patches of dead and/or dying plants. Dis­eased areas are covered with grayish-white mycelium. Elongated and flattened grayish-white sclerotia form on or within the infected tissue and blacken upon maturity.

Etiology

Fusarium patch disease may occur through­ out the year, while pink snow mold occurs only un­der a snow cover or very shortly after a thaw. Both Fusarium patch and pink snow mold are favored by cool, moist conditions. The conidial stage of F. nivale is the primary inoculum source. The disease is favored by cool, moist weather and high relative hu­midity. Conidia and mycelia produced under these climatic conditions invade healthy tissue. Perithecia of M. nivalis are abundant on cereals, but rarely ob­served on grasses in the field and do not form in cul­tures of F. nivale isolated from grass. Chlamydo­ spores are absent (Weibull, 1979b).

The etiologies of the three diseases caused by Typhula spp. are similar. Each fungus produces scle­rotia that remain dormant on the soil surface during the summer months. As temperatures become cooler and moist conditions prevail, the sclerotia germinate and produce one to three sporophores that bear basiodiospores. While the basidiospores may cause infections, the most infections are from mycelial growth into weakened tissues. Sclerotia buried in the soil may survive for several years.

Sclerotinia borealis produces sclerotia that are dor­mant during the summer months, and become a source of inoculum in autumn. Elongated asci, hav­ing eight ascospores, develop during moderately low temperatures and high moisture. Ascospores dis­charge onto surrounding foliage and are the main in­ oculum source. Infection may also occur by direct mycelial growth in weakened host tissues. The scle­rotia may survive burial in the soil for years.

Host Range

Snow molds occur on a wide variety of grasses and are particularly prevalent on Agrostis spp., Festuca spp., and Poa spp. During periods of prolonged snow cover, Lolium spp., timothy, and Agropyron spp. may be extensively damaged.

Control

Good management practices offer a measure of control for snow mold-susceptible grass species. Cutting low in autumn to reduce excessive top growth, avoiding heavy top dressings or winter mulches, and eliminating heavy or late nitrogen ap­plications may be effective. Abundant late-season nitrogen contributes to a dense host growth and creates a microclimate favorable to fungal development. Good drainage is important and fungicide applica­tions may be advisable.

Resistant cultivars offer a good measure of snow mold resistance in forage grasses (Abe and Matsu­ moto, 1981; Duich et al., 1972; Funk et al., 1969, 1973, 1974; Jamalanien, 1974; O'Rourke, 1976; Schmidt, 1976a, b; and summarized by Braverman, 1967, 1986).

Literature

Abe, J., and N. Matsumoto. 1981. Resistance to snow mold disease caused by Typhula spp. in cocksfoot. J. Jap. Soc. Grassl. Sci. 27:152-158. (Abstr. Rev. Appl. Plant Pathol. 61:148. 1982.)

Braverman, S.W. 1967. Disease resistance in cool sea­son forage, range and turf grasses. Bot. Rev. 33:329-378.

_________. 1986. Disease resistance in cool season for­age, range and turf grasses II. Bot. Rev. 52:1-112.

Duich, J.M., A.T. Perkins, and H. Cole. 1972. Regis­tration of Pennfine perennial ryegrass (Reg. No. 26). Crop Sci. 12:257.

Funk, C.R., R.W. Duell, and P.M. Halisky. 1969. Registration of Manhattan perennial ryegrass (Reg. No. 18). Crop Sci. 9:679-680.

_________, R.E. Engle, G.W. Pepin, A.M. Radko, and R.J. Peterson. 1974. Registration of Bonnieblue Kentucky bluegrass (Reg. No. 10). Crop Sci. 14:906.

_________, R.E. Engle, G.W. Pepin, A.M. Radko, and A.R. Russell. 1973. Registration of Adelphi Kentucky bluegrass (Reg. No. 9). Crop Sci. 13:580.

Jamalainen, E.A. 1974. Resistance in winter cereals and grasses to low temperature parasitic fungi. Ann. Rev. Phytopathol. 12:281-302.

O'Rourke, C.J. 1976. Diseases of grasses and forage legumes in Ireland. An Foras Taluntais. Oak Park Res. Centre. Carlow. 115 p.

Schmidt, D. 1976a. Diseases affecting persistence in Italian ryegrass. Arbeiten aus dem Gebiete des Futterbaues 20:50-59.

_________. 1976b. Observations sur la pourriture des neiges affectant les graminees. Revue Suisse d'Agriculture 8:8-14.

Weibull, Peter. 1979b. Descriptions of grass diseases. Weibull's Gras-tips 22:unpaginated.

___________. 1983. Descriptions of grass diseases. No. 12. Weibull's Gras-tips. p. 22-25, December 1983.

Diseases incited by Viruses

Viruses are submicroscopic infective entities that re­produce only in living cells. When purified, many viruses form crystals of characteristic sizes and shapes permitting classification. Dimensions of virus particles are measured in millimicrons; the particles may be rigid or flexuous rods, spheres, or pinwheels.

Symptoms of virus infections include a wide range of host reactions, some similar to those incited by bacteria or fungi. Two distinct symptoms commonly encountered in the Gramineae are a mosaic (or mottle) and a yellowing of leaf tissues. In the former, chlorophyll development is patchy, causing an une­venness in green coloring of the foliage and forma­tion of a yellow and green mosaic or mottle pattern. In leaves that yellow, there is a nearly uniform reduc­tion of chlorophyll, with little or no mosaic pattern. Other symptoms encountered in the Gramineae in­clude: veinbanding and veinclearing, streak, necro­sis, and tissue malformations. Quite often, virus­ infected plants are stunted (Walker, 1970).

Several methods of virus transmission are found in the Gramineae: mechanical, seed, insects, mites, nematodes, and soil (Slykhuis, 1976).

Many viruses infecting the Gramineae, such as bar­ley yellow dwarf virus, have a broad host range while the host range of others may be restricted to a few species (Rochow, 1970; Catherall, 1971b). With some viruses, the host range is dependent upon the strain of the virus (Catherall, 1971b).

Literature

Catherall, P.L. 1971b. Virus diseases of cereals and grasses. p. 308-309. In Diseases of Crop Plants. J.H. Western Ed. Macmillan. London. 404 p.

Rochow, W.F. 1970. Barley yellow dwarf virus. Com­ monwealth Mycological Institute/Association of Applied Biologists. Descriptions of Plant Viruses No. 32.

Slykhuis, J.T. 1976. Virus and virus-like diseases of cereal crops. Annu. Rev. Phytopathol. 14:189-210.

Walker, J.C. 1970. Plant Pathology. McGraw-Hill Book Co. New York. 819 p.

Ryegrass Mosaic.

Figure 22. Ryegrass mosaic virus on annual ryegrass, mild strain.

Figure 26. Ryegrass mosaic virus on annual ryegrass (severe strain).

Ryegrass mosaic virus (RMV) causes a light green to yellow mosaic on Lolium spp. and has been reported from North America and Europe (Slykhuis, 1972). In Great Britain, RMV is probably the most damaging disease of annual and perennial ryegrasses (Heard et al., 1974). The virus infects only members of the Poaceae and naturally occurs on annual and peren­nial ryegrasses, although several additional genera have been infected by mechanical inoculation.

Ryegrass mosaic virus is transmitted by the erio­phyid mite, Abacarus hystrix (Nelpa), and by manual sap inoculation to other species of the Poaceae. All in­stars of A. hystrix transmit RMV, but lose their infectivity within 24 hours after removal from a virus source (Slykhuis, 1972).

RMV is readily transmitted by sap inoculation in the ryegrasses. The virus is not seed- or soil-borne; nor has transmission through dodder (Cuscuta spp.) been reported. In the field, RMV is probably intro­ duced and spread within the crop by mites and by grazing.

Symptoms

RMV is systemic and causes pale green streaks in ryegrass foliage, most evident in the upper leaves of the flowering stem. On older leaves, the streaks become yellow or brown and distinct. On an­nual ryegrass, the entire lamina occasionally becomes dark brown. In perennial ryegrass, the virus reduces tillering; in late stages of infection, the plants are also stunted. In general, annual ryegrass is more suscepti­ble and more severely affected than perennial ryegrass.

Control

The severity of RMV in Great Britain and Europe has stimulated progress toward producing disease resistant cultivars in both species (Braver­man, 1986). Resistance in the ryegrasses is polygenic (controlled by more than one gene). Interspecific crosses between perennial ryegrass and annual rye­grass combine the longevity of perennial ryegrass and the yield of annual ryegrass. Resulting lines are intermediate in reaction to RMV (Braverman, 1986).

Host Range

In addition to infections of annual and perennial ryegrass, RMV has been transmitted mechanically to orchardgrass, meadow fescue, annual meadowgrass, Kentucky bluegrass, rough bluegrass (Poa trivialis L.), smooth bromegrass, colonial bent­ grass, Bromus arvensis L., and B. sterilis L. (Mulligan, 1960).

Literature

Braverman, S.W. 1986. Disease resistance in cool sea­son forage, range and turf grasses II. Bot. Rev. 52:1-112.

Heard, A.J., J.A. Brook, E.T. Roberts, and R.J. Cook. 1974. The incidence of ryegrass mosaic virus in crops of ryegrass grown for seed in some southern counties of England. Plant Pathol. 23:119-127.

Mulligan, T.E. 1960. The transmission by mites, host­ range and properties of ryegrass mosaic virus. Ann. Appl. Biol. 48:575-579.

Slykhuis, J.T. 1972. Ryegrass mosaic virus. Common­wealth Mycological Institute/Association of Applied Biologists. Descriptions of Plant Viruses No. 86.

Cocksfoot Mottle.

Figure 23. Cocksfoot mottle virus on orchardgrass.

Cocksfoot mottle virus (CfMV) causes a severe mott­ling and "dying out" of orchardgrass. The virus, not reported outside of central and southern England (Catherall, 1971), infects only a few species in the Poaceae.

CfMV is transmitted by adults and larvae of a Chrysomelid beetle, Oulema melanopus L. The adults are the more efficient vectors, and may be infective up to two weeks after acquiring the virus from in­fected tissue. CfMV is readily transmitted by sap in­ oculation in the orchardgrasses (Catherall, 1971a).

The virus is not seed-borne; transmission by dod­der has not been tested. CfMV may be spread in the field by mowing implements and (presumably) by grazing animals.

Symptoms

Symptoms of CfMV are similar to those of cocksfoot streak virus (CSV). The virus is systemic, and the most striking symptoms occur in spring and early summer. Young leaves display a yellow streak­ing or mottling which often becomes white or necrotic as the leaf matures. Infected leaves die pre­maturely; severely infected tussocks appear flat­tened, with young mottled tillers standing erect among a mass of streaked, yellow, and dying leaves. Infected plants occasionally flower, but set few viable seed. Although infected plants will form a few new tillers, the plants from these tillers are stunted and mottled and do not flower (Catherall, 1971b; Serjeant, 1964, 1967).

Control

Considerable variation in genotype re­sponse exists in orchardgrass. 'Conrad' and 'Cam­bria,' released by the Welsh Plant Breeding Station, Aberysthwyth, Wales, are highly resistant. The Japa­nese cultivar Okamidori is also CfMV resistant. One hybrid, D. mariana Borill x D. glomerata, has shown good CfMV resistance (Wilkins, 1977).

Host Range

The virus infects orchardgrass, other Dactylis spp., wheat, oats, and barley (Catherall, 1971a). Several additional species were identified as hosts in inoculation studies (Catherall et al., 1977).

Literature

Catherall, P.L. 1971a. Cocksfoot mottle virus. Com­monwealth Mycological Institute/Association of Applied Biologists. Descriptions of Plant Viruses No. 23.

__________. 1971b. Virus diseases of cereals and grasses. p. 316-317. In Diseases of Crop Plants. J.H. Western Ed. Macmillan. London. 404 p.

__________, P.A. Andrews, and J.A. Chamberlain. 1977. Host ranges of cocksfoot mottle and cyno­surus mottle viruses. Ann. Appl. Biol. 87:233-235.

Setjeant, E.P. 1964. Cocksfoot mottle virus. Plant Pathol. 13:23-24.

__________. 1967. Some properties of cocksfoot mottle virus. Ann. Appl. Biol. 59:31-38.

Wilkins, P.W. 1977. Herbage viruses. p. 176-177. In Welsh Plant Breeding Station Report for 1977. Aberysthwyth.

Cocksfoot Streak.

Cocksfoot streak virus (CSV) is an aphid-stylet-borne (nonpersistent) virus which infects the Grarnineae in Britain. It is also widespread in Germany, France, The Netherlands, and Sweden. According to Catherall (1971b), a similar (but possibly distinct) virus, orchardgrass mosaic, occurs in North America. While in Britain, CSV is restricted to Dactylis and Lo­lium spp., the virus has been transmitted to a multi­tude of grasses. Many are symptomless carriers.

CSV is transmitted primarily by five species of aphids: Myzus persicae (Sulz.); Macrosiphum euphorbiae (Thos.); M. avenae (F.); Hyalopteroides humulis (Wik.), and Metopolophium dirhodum (Wik.). The aphids be­ come infective shortly after feeding on infected hosts, but the virus persists for less than one hour. With dif­ficulty, CSV can also be transmitted mechanically by sap inoculation. However, this mode of transmission is probably of minor consequence in spreading the virus.

Initial symptoms of CSV on orchardgrass and rye­ grass are pale green or yellow streaks on the young leaves. As infection progresses, the streaks increase in intensity and eventually spread throughout the host. Infected plants are not stunted to any extent, but vegetative tiller production is usually reduced by 40 percent. Seed yield is slightly reduced (Catherall, 1971b).

Control

Orchardgrass cultivars resistant to the virus have not been found (Catherall, 1971b).

Host Range

In Britain, CSV infects only orchardgrass and ryegrass; in Germany it has been transmitted to several grasses, some of which are symptomless car­riers (Catherall, 1971b).

Literature

Catherall, P.L. 1971b. Virus diseases of cereals and grasses. p. 308-309. In Diseases of Crop Plants. J.H. Western Ed. Macmillan. London. 404 p.

Cocksfoot Mild Mosaic.

Figure 25. Cocksfoot mild mosaic virus on orchardgrass.

Cocksfoot mild mosaic (CMM) has been reported in Germany and England and infects only species in the Gramineae. In orchardgrass, CMM causes a diffuse mild mosaic or mottle, and at times may induce a strong chlorotic streaking (Huth and Paul, 1972). Lo­lium persicum Boiss. & Hohen ex Boiss. and foxtail millet are additional diagnostic hosts. Perennial rye­ grass and meadow fescue, while not susceptible to the type strain of CMM, are susceptible to a different virus strain. CMM is transmitted by Myzus persicae, but transmission by seed or by dodder has not been reported.

CMM and Phleum mottle viruses have distinct host ranges, but are serologically related. CMM does not infect timothy, and Phleum mottle virus does not in­fect orchardgrass.

Literature

Huth, W., and H.L. Paul. 1972. Cocksfoot mild mo­saic virus. Commonwealth Mycological Institute/ Association of Applied Biologists. Descriptions of Plant Viruses No. 107.

Agropyron Mosaic.

Agropyron mosaic virus (AMV), which causes a mo­saic in the leaves of many Gramineae, has been re­ ported in the United States, Canada, and Europe. Ac­cording to Slykhuis (1973), only species of the Gramineae have been found to be systemically in­fected, although the virus does produce local lesions in Chenopodium quinoa Willd., a dicotyledonous species.

AMV is transmitted by an eriophyid mite, Abacerus hystrix, and also is sap transmissible. The virus per­sists in the rhizomes of its respective hosts, but is not seed-borne. Transmission by dodder has not been re­ported. In the field, AMV is most likely spread by mowing implements and presumably by the trampling of virus-infected foliage by grazing animals.

Symptoms

Systemically infected species in the Gra­mineae develop a light green to yellow·mosaic and striping on the foliage. Some stunting is evident.

Host Range

In North America, wheat appears to be highly susceptible to the virus. Several Agropyron species are moderately susceptible. They include A. elongatum (Host.) Beauv., beardless wheatgrass (A. inerme (Scribn. & Smith) Rydb.), A. intermedium (Host.) Beauv., quackgrass, and fescue. Wild rye­grass and annual ryegrass are less susceptible. A ma­jority of the reported hosts are found in Europe (Slykhuis, 1973).

Literature

Slykhuis, J.T. 1973. Agropyron mosaic virus. Com­monwealth Mycological Institute/Association of Applied Biologists. Descriptions of Plant Viruses. No. 118.

Bromegrass Mosaic.

Bromegrass mosaic virus (BMV) causes a mild mosaic in most species of Gramineae in the United States and Europe. The monocotyledonous host range is large, but the dicotyledonous host range consists of a few genera in five families (Bancroft, 1970). BMV sur­vives in air-dried leaf tissue for one year or longer.

Schmidt et al. (1963) reported transmission of BMV by two nematodes- Xiphinema paraelongatum Atherr. (= X. diversicaudatum (Micoktzky) Thome) and X. coxi Tarjan -- in plants grown in a greenhouse, but field transmission of the virus by those nematodes has not been reported. The virus is also transmitted by ani­mals (McKinney, 1953). BMV is not seed-borne (Lane, 1974). Its transmission by dodder has not been reported. Attempts to transmit BMV by selected in­sects have been unsuccessful (Lane, 1974).

Symptoms

Bromegrass mosaic virus causes a chlo­rotic mottling and striping, and brown necrotic stripes and blotches. Culms are shortened and the in­florescences are frequently sterile. Occasionally, ex­cessive tillering may occur.

Although BMV is widely distributed on smooth bromegrass, the virus has a broad host range in the Gramineae. This is based on mechanical inoculations including other grasses. The disease is generally mild and of little economic importance. The virus appears to lack the appropriate insect vector (Gibson and Kenten, 1978).

Literature

Bancroft, J.B. 1970. Bromegrass mosaic virus. Com­monwealth Mycological Institute/Association of Applied Biologists. Descriptions of Plant Viruses No. 3.

Gibson, R.W., and R.H. Kenten. 1978. The occur­rence of brome mosaic virus in Britain. Plant Pathol. 27:66-67.

Lane, L.C. 1974. The brome viruses. Advances in Virus Research 19:151-220.

McKinney, H.H. 1953. Virus diseases of cereal crops. p. 350-360. In USDA Yearbook of Agriculture - 1953.

Schmidt, H., R. Fritzsche, and W. Lehmann. 1963. Die Ubertragung des Weidelgrasmosaik-Virus Nematoden durch Nematoden. Die Naturwis­ senschaften 50:386.

Barley Yellow Dwarf.

Figure 24. Barley yellow dwarf virus on annual ryegrass (yellowing and dwarfing).

Barley yellow dwarf virus (BYDV), which causes a stunting and chlorosis of a wide range of monocotyle­donous species, is worldwide in distribution (Rochow, 1970; Panayotou, 1982). While BYDV is probably more widespread than any other virus in­fecting cereals, Doodson (1967) concludes that a large reservoir of the virus exists in England and Wales.

Barley yellow dwarf virus is transmitted by about 14 species of aphids, including Macrosiphum avenae, Rhopalosiphum maidis F., R. padi (L.), and Schizaphis gramineum.

The virus is not seed-borne. However, BYDV has been recovered by aphids feeding on dodder estab­lished on infected barley (Rochow, 1970).

Symptoms

Barley yellow dwarf virus is systemic; however, some infected grasses show no symptoms. In perennial ryegrass, infected foliage is character­ized by yellow or red leaf discolorations and/or stunting. Catherall (1963) and Doodson (1967) have emphasized the importance of infected perennial rye­grass as a reservoir of virus-infected material from which growing cereal crops could become infected by migrating aphids.

Control

Breeding for BYDV resistance in perennial ryegrass not only serves to develop a virus-free cultivar(s) but also to reduce a potential BYDV reser­voir for future cereal crop infections. BYDV resistance in Lolium perenne is a complex problem (Catherall and Wilkins, 1977) due in part to the lack of symptom ex­pression in susceptible cultivars and excessive till­ering to compensate for plant dwarfing due to infection.

Host Range

The virus infects a wide range of mono­cotyledonous species (Rochow, 1970).

Literature

Catherall, P.L. 1963. Transmission and effect of bar­ley yellow dwarf virus isolated from perennial rye­grass. Plant Pathol. 12:157-160.

_________, and P.W. Wilkins. 1977. Barley yellow dwarf virus in relation to the breeding and assess­ment of herbage grasses for yield and uniformity. Euphytica 26:385-391.

Doodson, J.K. 1967. A survey of barley yellow dwarf virus in S.24 perennial ryegrass in England and Wales. Plant Pathol. 16:42-45.

Panayotou, P.C. 1982. Some aspects on barley yellow dwarf virus host range. Zeitschrift fur Pflanzen­ krankheiten und Pflanzenschutz 39:595-603.

Rochow, W.F. 1970. Barley yellow dwarf virus. Com­monwealth Mycological Institute/Association of Applied Biologists. Descriptions of Plant Viruses No. 32.

Phleum Mottle.

Catherall (1966) proposed the name phleum mottle virus (PMV) for a Gramineae-infecting virus distinct from other grass-infecting viruses. The physical properties of PMV confirm that it belongs in the southern bean mosaic group of beetle-transmitted plant viruses (Walters, 1969). PMV is reported to occur only in central and southeastern Britain. PMV includes five distinct strains, as follows: PM (phleum mottle); HTM (holcus transitory mottle); FM (festuca mottle); CMM (cocksfoot mild mosaic), and BSM (brome stem leaf mottle).

PMV is transmitted by two species of cereal leaf beetle, Lema melanopa L. and L. lichensis Weise. Adult beetles of either species are more efficient vectors than larvae. Some beetles retain the PMV virus for as long as two weeks after feeding on an infected plant. Attempts to transmit PMV with insects other than beetles, including aphids, were unsuccessful.

The virus is readily sap-transmissible and can be spread by farm machinery and presumably by graz­ing animals.

Symptoms

Initial infection appears as pale green or yellow streaks that form at the base of new growth and gradually extend to the leaf tip. Streaks coalesce to form a noticeable mottle, eventually covering the entire leaf (Catherall, 1970). Infection reduces the number of tillers and fresh green weight, but appar­ently does not affect plant height or number of inflorescences.

Control

Resistance to PMV in P. pratense cultivars has not been reported. Alpine timothy (P. alpinum L.) and P. commutatum Guad ( = P. alpinum) do show re­sistance to PMV (Catherall, 1970).

Host Range

Suscepts include Agrostis spp., Bromus spp., orchardgrass, meadow fescue, Lolium persicum, reed canarygrass, timothy, and Poa spp.

Literature

Catherall, P.L. 1966. Phleum mottle virus. Rep. Welsh Plant Breeding Stn. for 1965. p. 94.

___________. 1970. Phleum mottle virus. Plant Pathol. 19:101-103.

Walters, H.J. 1969. Beetle transmission of plant viruses. Adv. Vir. Res. 15:339-363.

Mineral Deficiencies

This section describes characteristic effects on plants of various mineral disorders but does not attempt to be comprehensive. For a more extensive coverage of minerals and their role in plant growth and composi­tion, the reader is referred to the volumes edited by C. Bould, E.J. Hewitt, and P. Needham titled "Diag­nosis of Mineral Disorders in Plants," Volume 1, "Principles," and Volume 2, "Vegetables," published by Chemical Publishing, N.Y., 1984, from which the following information has been summarized.

Macronutrient Deficiencies

Calcium.

In plant tissue, the concentration of calcium is important to growing points and young leaves and is usually related to the distribution of specific symp­toms, but does not entirely explain the appearance of others. In cereals and grasses, emerging young leaves remain trapped in subtending leaves. Leaves which have emerged remain rolled, chlorotic, and have circular constrictions a few centimeters behind the apex. The distal portion wilts and withers. Death of the stem apex occurs in calcium-deficient plants, but multiple apical shoots or axillary nodal shoots may follow to fill the void.

Magnesium.

Magnesium is very important in the makeup of chlorophyll. Therefore, magnesium defi­ciency is most frequently indicated first by the loss of chlorophyll. As a rule, the chlorosis usually appears first in the oldest leaves and is progressive. In several species, the chlorosis is generally interveinal within a persistent green margin of the leaf.

Symptoms of magnesium deficiency may be diffi­cult to distinguish from the symptoms of potassium deficiencies at certain stages of plant growth. The principal distinction is that magnesium deficiency af­fects first the oldest leaves, while potassium defi­ciency usually is first noticed in younger leaves.

Nitrogen.

Nitrogen-deficient plants are much smaller than normal, but the root growth can be extensive and root length may increase to compensate for the absence of N. The angle between petioles and stems is more acute. Tillering is suppressed, bud produc­tion or expansion is decreased, and flowering is de­layed. Bud dormancy can be prolonged. Foliage is pale green and leaf senescence is accelerated. De­pending upon species, leaves can develop purple, red, or orange anthocyanin tints in addition to the yellowishness caused by the loss of chlorophyll. lnterveinal areas and older leaves are first to express symptoms. Leaf bases and stems become red-purple.

Phosphorus.

Symptoms of phosphorus deficiency frequently resemble those of nitrogen deficiency. There is a diminutive or spindly habit, suppression of tillering, acute leaf angles, decreased size and num­bers of flowers, prolonged dormancy, and early se­nescence. The leaves lack luster; leaf color changes, but may be either paler or darker than normal. Again, depending on species, leaf color may range from deep purple to red, or be absent.

Potassium.

Potassium deficiency frequently appears as short telescoped shoots, caused by a shortening of stem internodes. Apical dominance of the viable ter­minal bud is sometimes suppressed, causing exces­sive basal shoots or tillers. Severe deficiency causes death of the terminal bud and a typical dieback. Strong light intensity accentuates and weak light of­ten reduces or eliminates these conditions. Leaf scorch, preceded by irregular marginal or interveinal chlorosis, appears first in the oldest leaves. Scorching may be pale brown to almost black. Chlorosis almost always occurs and first appears in oldest leaves, which often curve downwards or become convex.

Sulphur.

Decreased leaf size, red or purple pigmen­tation, and general chlorosis symptoms of sulphur deficiency resemble those of nitrogen. However, there are important differences. Usually young leaves are more sensitive to sulphur deficiency than older ones. In sulphur deficient soils, new leaves are fre­quently uniform golden yellow and stiff and erect.

Micronutrient Deficiencies

Chlorine.

Little has been reported in the literature as to the effect of chlorine deficiency on plants in the Gramineae. Chlorine deficiency has been reported to limit growth and cause wilt, chlorosis, and promi­nent raised veins and clubbed tips of roots in dicoty­ledons. Deficiency will keep barley leaves rolled, as does copper deficiency.

Copper.

In many plant species, the young leaves are most severely affected. Leaves are often rolled or curled, and may be coiled in a spiral which may re­verse direction along its length. These leaves are of­ten white. The emerging leaves may be trapped in the subtending leaf, producing symptoms known as white tip or wither tip. Floral meristems are sensitive to copper deficiency. Head formation is suppressed, and the grains are shriveled or the glumes do not fill. Often the pollen cells are sterile.

Iron.

Iron deficiency is first apparent as chlorosis of rapidly expanding leaves. Chlorosis is usually inter­veinal and produces a contrasting tramline effect in the leaves. The glumes may be more chlorotic than the flag leaf. In cereals and grasses, bleached or brown lesions develop more frequently in the interveinal areas and leaves collapse transversely.

Manganese.

A variety of symptoms (including some form of chlorosis) are produced by manganese deficiency. Older leaves are usually affected first. Chloro­sis is usually interveinal and produces a bold pattern of dark green major veins which contrasts with the fine reticulate pattern observed with iron deficiency. Manganese deficiency is also distinguished from iron deficiency symptoms in leaves by the appearance of varied but characteristic necrotic spotting or lesions. In the Gramineae, these lesions vary in appearance from dark brown spots along interveinal areas to chlorotic beading. In oats, the necrotic lesions are elongated, generally ivory to pale brown with "blue­ green or grey" halo areas, and often coalesce into broad lesions that may collapse leaves transversely.

Ice and Water Damage

Ice and water damage prevail in areas with alternate freezing and thawing and in maritime locales of pro­longed high humidity. Such areas may be character­ized by poor soil drainage or the soil may be frozen, thereby preventing or substantially reducing water flow. Damage increases during prolonged ice cover.

Symptoms

Small differences in topography influ­ence the extent and severity of damage. The grass, usually green after initial ice melt in the spring, soon becomes dark brown and subsequently light brown upon drying. At this stage, the crowns and roots have been damaged and the entire plant begins to rot (Weibull, 1979c). Species forming rhizomes or stolons may regenerate later in the growing season to fill in killed patches. While ice and water damage may af­fect any grass species, cultivars differ in susceptibil­ity. Kentucky bluegrass is tolerant and perennial rye­grass is sensitive, for example.

Literature

Weibull, Peter, 1979c. Descriptions of grass diseases No. 11. Ice and water damage. Weibull's Gras-tips 22:unpaginated.

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Glossary

Morphological definitions are based in part on Kreitlow, J.W., et al., 1953, and O'Rourke, C.J., 1976.

Abiotic

[A disease] not caused by a biological agent.

Acervulus

(i). A small, open fruiting body that has ruptured the host epidermis and consists of a mass of hyphae bearing conidiophores and conidia.

Aeciospore

A spore formed in chains within the ae­cium of a rust.

Aecium

(a). A cup-shaped fruiting structure charac­teristic of the rusts in which aeciospores are borne.

Alternate Host

Either of two unrelated plant spe­cies necessary for certain rust fungi to complete their life cycle.

Amphigenous

Making growth all around or on two sides.

Apothecium

(a). An open cup or saucer-shaped fruiting body whose concave surface is lined with asci.

Ascigerous

Having asci.

Ascocarp

The ascus-bearing structure of the Ascomycetes.

Ascospore

One of normally eight spores borne in an ascus.

Ascus

(i). A sac-like structure in which normally eight ascospores are formed as a result of the sex­ual process.

Asexual Reporduction

Reproduction without fu­sion of gametes.

Autoecious

Referring to a parasite which completes its life cycle on one host.

Bacterium

(a). A single-celled microscopic organism lacking a well-defined nucleus or nuclear membrane.

Basidospore

A spore produced at the apex of a basidium.

Basidium

A fruiting structure which bears the basidiospores.

Biotic

[A disease] caused by a biological agent. Blind Seed. Sterile seed.

Chlamydospore

A thick-walled asexual resting body formed by the rounding up of a mycelial segment.

Chlorotic

Deficient in chlorophyll.

Cleistothecium

(a). A fungal fruiting body, con­taining asci, which has no specific opening (e.g., in Erysiphaceae).

Conidium

(a). An asexual spore.

Conidiophore

A specialized branch of the myce­lium bearing cells from which conidia are borne.

Culm

The stem of a grass.

Cuticle

The outermost layer of leaf tissue.

Disease

An interaction between a causal agent and host which alters the morphological and physiolog­ical development of the host.

Disease Development

The sequence of events from time of infection to symptom expression.

Disease Incitant

An abiotic or biotic entity which may cause disease.

Endemic

The occurrence of a disease from year to year in moderate to severe form.

Epiphytic

A large-scale disease outbreak in plants.

Forma (ae) specialis (es)

A subdivision within a species, distinguished by physiological traits rather than by morphological characteristics.

Germination

The process by which a hypha emerges from a spore.

Guttation

The process of the escape of liquid water from uninjured plants.

Haustorium

(a). A specialized mycelial branch, es­pecially one within a living cell of a host.

Heteroecious

Referring to a parasite that completes its life cycle on unrelated hosts.

Host Plant

A plant morphologically and physiolog­ically altered by a disease incitant.

Host Range

Those plant species known to be sus­ceptible to a pathogenic organism.

Hydathode

A pore-like structure in a plant leaf through which guttation occurs.

Hypha

(ae). A single strand or strands of mycelium.

Infection

The establishment of a disease incitant within a host.

Lesion

An area of tissue showing disease symptoms.

Life Cycle

A sequential series of forms and relation­ships assumed by an organism from a primary stage to a resumption of that stage.

Macroconidium

(a). The larger conidium of a fun­gus which also has microconidia.

Microconidium

(a). The smaller conidium of a fun­gus which also has macroconidia.

Morphology

A study dealing with the form and structure of organisms.

Mycelium

(a). The vegetative body of the fungus comprising a mass of hyphae.

Necrotic

Dead or dying.

Obligate Parasite

An organism that grows only on its host and which usually cannot be cultured on artificial media.

Ostiole

An opening from which spores extrude from an ascigerous or pycnial fruiting body.

Parasite

An organism which derives its nourishment from another organism, the host.

Pathogen

A disease-causing organism.

Pathovar

A specialized variety of a pathogenic or­ganism based on host range or its characteristic growth on a specific culture medium.

Penetration

An initial invasion of a host by a dis­ease incitant.

Perithecium

(a). A round-oval, flask-shaped fruiting structure containing asci and an apical aperture.

Physiologic Race

A pathogen similar in morpholog­ical characteristics but differing in ability to parasi­tize certain varieties of the host.

Polygenic

Having more than one gene.

Primary Inoculum

The initial infective disease­ inciting agent (pathogen) causing infection in the host plant.

Pycnidium

(a). A small globose or flask-shaped fruit­ing body containing asexual spores.

Pycnidiospore(s)

A spore or spores developed in a pycnidium.

Saprophyte

An organism which derives its nourish­ment from dead organic matter.

Sclerotium

(a). A firm-bodied resting structure formed by certain fungi consisting of a mass of hy­phae surrounded by a darker outer ring.

Secondary Inoculum

Spores or infective bodies produced after the host has been colonized.

Secondary Infection

An infection resulting from inoculum produced by the pathogen during pri­mary infection or during the production of second­ary inoculum.

Seta

(ae). A small, slender, usually rigid bristle or hair.

Sexual Reproduction

Reproduction requiring nu­clear fusion and meiosis.

Sorus

(i). A spore-containing body of rusts and smuts which become exposed at maturity upon rupturing the epidermis of the host.

Spore

A specialized cell(s) adapted for dissemina­tion and capable of germinating to perpetuate the species.

Sporidium

(a). One of several spores borne on a spe­cialized hypha (promycelium) which develop from teliospores of rusts and smuts.

Spordochium

A compact conidial body; mass of sporophores.

Strain

An organism with similar morphologic char­acteristics to another but with different physiologic characters.

Stromatic Body (stroma)

A mass of vegetative hy­phae in or on which fruiting structures develop. Substrate. The base upon which an organism de­rives its nourishment.

Symptom

The expression by the host resulting from the host-pathogen disease-incitant interaction.

Systemic

Spread throughout the host tissues; not lo­calized or confined within specific boundaries.

Teliospore

A spore produced by rusts and smuts which germinates to form a hypha (promycelium) on which sporidia are abstricted.

Telium

(a). A spore-bearing body or sorus in which teliospores are produced.

Tussock

A compact tuft (clump) of a grass.

Uredium

(a). A spore-bearing body or sorus in which urediospores are produced.

Urediospore(s)

A spore characteristic of the rust fungi capable of initiating secondary infection by reinfecting the same host.

Vector

A living organism capable of transmitting a disease incitant.

Virus

A submicroscopic infective entity which can reproduce only in living cells.

Yellows

A foliage condition caused by destruction of the chlorophyll.

Diseases Indexed by Host

Bentgrass and Red Top

• Anthracnose
• Ascochyta leafspot (A. sorghi)
• Bipolaris foot rot
• Blister smuts (leafspot smuts)
• Cocksfoot mottle
• Covered smuts (kernel smuts)
• Crown rust
• Drechslera leafspot
• Ergot
• Kernel smuts (covered smuts)
• Leaf rust
• Leafspot smuts (blister smuts)
• Mastigosporium leaf fleck
• Phleum mottle
• Powdery mildew
• Red thread
• Rhizoctonia blight
• Stem rust (black stem rust)
• Stripe rust (yellow rust)
• Stripe smut
• Snow molds
• Take-all
• Uromyces leaf rust

Bluegrass

• Agropyron mosaic
• Anthracnose
• Ascochyta leafspot (A. sorghi)
• Bacterial stripe
• Bacterial wilt
• Bipolaris foot rot
• Blind seed
• Blister smuts (leafspot smuts)
• Bromegrass mosaic
• Cocksfoot mottle
• Crown rust
• Ergot
• Flag smut
• Leaf rust
• Leafspot (Drechslera poae)
• Leafspot smuts (blister smuts)
• Melting out
• Powdery mildew
• Red thread
• Rhizoctonia blight
• Ryegrass mosaic
• Snow molds
• Stem rust (black stem rust)
• Stripe rust (yellow rust)
• Stripe smut
• Take-all
• Uromyces leaf rust
• Yellow leaf rust

Crested wheatgrass

• Agropyron mosaic
• Anthracnose
• Ascochyta leafspot (A. sorghi)
• Bacterial brown stripe
• Bacterial stripe
• Bipolaris foot rot
• Ergot
• Flag smut
• Head smuts (loose smuts)
• Loose smuts (head smuts)
• Powdery mildew
• Rhizoctonia blight
• Snow molds
• Stem rust (black stem rust)
• Stripe rust (yellow rust)
• Take-all

Agropyron species

• Agropyron mosaic
• Anthracnose
• Ascochyta leafspot (A. sorghi)
• Bacterial brown stripe
• Bipolaris foot rot
• Brown spot/halo blight
• Crown rust
• Ergot
• Halo blight/brown spot
• Head smuts (loose smuts)
• Loose smuts (head smuts)
• Powdery mildew
• Rhizoctonia blight
• Snow molds
• Stem rust (black stem rust)
• Stripe rust (yellow rust)
• Stripe smut
• Take-all
• Translucent leaf stripe

Fescues

• Agropyron mosaic (red fescue)
• Anthracnose
• Ascochyta leafspot (A. sorghi)
• Bacterial brown stripe
• Bacterial wilt
• Barley yellow dwarf
• Bipolaris foot rot
• Blister smuts (leafspot smuts)
• Brown blight (meadow fescue)
• Brown spot/halo blight
• Crown rust
• Ergot
• Flag smut (red fescue)
• Halo blight/brown spot
• Leaf blotch (Rhynchosporium)
• Leaf rust
• Leaf scald
• Leafspot smuts (blister smuts)
• Leaf streak (Rhynchosporium)
• Leaf streak (Scolecotrichum)
• Net blotch
• Phleum mottle
• Powdery mildew
• Red thread (red fescue)
• Rhizoctonia blight
• Ryegrass mosaic
• Snow molds
• Stem rust (black stem rust)
• Stripe rust (yellow rust)
• Take-all (red fescue)
• Uromyces leaf rust
• Yellow leaf rust

Orchardgrass

• Anthracnose
• Ascochyta leafspot (A. sorghi)
• Bacterial wilt
• Bipolaris foot rot
• Blind seed
• Brown blight
• Cocksfoot mild mosaic
• Cocksfoot mottle
• Cocksfoot streak
• Ergot
• Flag smut
• Leaf blotch (Rhynchosporium)
• Leaf scald
• Leafspot (Drechslera catenaria)
• Leafspot/blotch (Drechslera dactylidis)
• Leaf streak (Rhynchosporium)
• Leaf streak (Scolecotrichum)
• Mastigosporium leaf fleck
• Powdery mildew
• Phleum mottle
• Purple leafspot
• Rhizoctonia blight
• Ryegrass mosaic
• Snow molds
• Stem rust (black stem rust)
• Stripe rust (yellow rust)
• Stripe smut
• Translucent leaf stripe
• Uromyces leaf rust
• Yellow slime

Reed canarygrass

• Anthracnose
• Crown rust
• Ergot
• Drechslera leafspot (D. catenaria)
• Leafspot/blotch (Septoria bromi)
• Phleum mottle
• Powdery mildew
• Rhizoctonia blight
• Stripe smut
• Tawny blotch

Phalaris spp.

• Anthracnose
• Ascochyta leafspot
• Crown rust
• Ergot
• Rhizoctonia blight
• Tawny blotch

Annual ryegrass

• Bacterial wilt
• Take-all

Perennial ryegrass

• Red thread

Annual and perennial ryegrasses

• Agropyron mosaic
• Anthracnose
• Bacterial brown stripe
• Barley yellow dwarf
• Bipolaris foot rot
• Blind seed
• Bromegrass mosaic
• Brown blight
• Brown spot/halo blight
• Cocksfoot streak
• Crown rust
• Ergot
• Halo blight/brown spot
• Leaf blotch (Rhynchosporium)
• Leaf scald
• Leaf streak (Rhynchosporium)
• Leaf streak (Scolecotrichum)
• Mastigosporium leaf fleck
• Net blotch
• Phleum mottle
• Powdery mildew
• Rhizoctonia blight
• Ryegrass mosaic
• Snow molds
• Speckled leaf blotch
• Spermospora leafspot
• Stem rust (black stem rust)
• Stripe rust (yellow rust)
• Stripe smut

Smooth bromegrass

• Ascochyta leafspot (A. sorghi)
• Bacterial brown stripe
• Barley yellow dwarf
• Bipolaris foot rot
• Blackish-brown stripe
• Bromegrass mosaic
• Brown leafspot
• Brown spot/halo blight
• Ergot
• Flag smut
• Halo blight/brown spot
• Leaf blotch (Rhynchosporium)
• Leaf scald
• Leaf streak (Rhynchosporium)
• Rhizoctonia blight
• Selenophoma leafspot (S. bromigena)
• Septoria leafspot (S. bromi)
• Stagonospora leafspot (S. bromi)
• Translucent leaf stripe

Bromus spp.

• Ascochyta leafspot (A. sorghi)
• Bacterial brown stripe
• Bipolaris foot rot
• Bromegrass mosaic
• Brown leafspot
• Brown spot/halo blight
• Cocksfoot mottle
• Crown rust
• Ergot
• Halo blight/brown spot
• Head smuts (loose smuts)
• Leaf scald
• Leaf blotch (Rhynchosporium)
• Leaf streak (Rhynchosporium)
• Leaf streak (Scolecotrichum)
• Loose smuts (head smuts)
• Phleum mottle
• Powdery mildew
• Ryegrass mosaic
• Selenophoma leafspot (S. bromigena)
• Septoria leafspot (S. bromi)
• Stagonospora leafspot (S. bromi)
• Stripe smut
• Translucent leaf stripe

Sorghum

• Bacterial stripe
• Bacterial brown stripe
• Bacterial leafblight
• Powdery mildew

Sudangrass

• Anthracnose
• Ascochyta leafspot (A. sorghi)
• Bacterial stripe
• Bacterial leafblight
• Ergot
• Powdery mildew

Tall oatgrass

• Anthracnose
• Ascochyta leafspot (A. sorghi)
• Bacterial brown stripe
• Bacterial wilt
• Bipolaris foot rot
• Ergot
• Head smuts (loose smuts)
• Leaf streak (Scolecotrichum)
• Loose smuts (head smuts)
• Powdery mildew
• Rhizoctonia blight
• Yellow leaf rust

Timothy

• Anthracnose
• Bacterial wilt
• Barley yellow dwarf
• Blind seed
• Blister smuts (leaf spot smuts)
• Bipolaris foot rot
• Bromegrass mosaic
• Brown spot/halo blight
• Crown rust
• Drechslera leaf streak (D. phlei)
• Ergot
• Eyespot
• Flag smut
• Halo blight/brown spot
• Leaf spot smuts (blister smuts)
• Leaf streak (Scolecotrichum)
• Mastigosporium leaf fleck
• Phleum mottle
• Powdery mildew
• Rhizoctonia blight
• Snow molds
• Stem rust (black stem rust)
• Stripe rust (yellow rust)
• Stripe smut
• Translucent leaf stripe

Wild ryegrass

• Agropyron mosaic
• Anthracnose
• Ascochyta leafspot (A. sorghi)
• Bipolaris foot rot
• Cocksfoot mottle
• Crown rust
• Ergot
• Flag smut
• Head smuts (loose smuts)
• Loose smuts (head smuts)
• Powdery mildew
• Rhizoctonia blight
• Stripe smut

Warm-season Grasses

Big bluestem

• Ascochyta leafspot (A. sorghi)
• Stem rust (black stem rust)

Little bluestem

• Ascochyta leafspot (A. sorghi)
• Switchgrass Spot blotch

Indiangrass

• Ascochyta leafspot (A. sorghi)

Illustrations

Figure 1. Cells of Xanthomonas campestris pv. graminis exuding from the vascular system of annual ryegrass (Lolium multiflorum Lam.)

Figure 2. Yellow slime, Corynebacterium rathayi, on orchardgrass.

Figure 3. Bacterial stripe, Pseudomonas andropogonis, on sudangrass.

Figure 4. Halo blight, Pseudomonas avenae, on oats.

Figure 5. Brown stripe, Pseudomonas avenae, on mountain bromegrass.

Figure 6. Halo blight, Pseudomonas syringae pv. coronafaciens, on mountain bromegrass.

Figure 7. Halo blight, Pseudomonas syringae pv. coronafaciens, on annual ryegrass (early spring symptoms).

Figure 8. Halo blight, Pseudomonas syringae pv. coronafaciens, on annual ryegrass (late summer symptoms).

Figure 9. Bacterial wilt, Xanthomonas campestris pv. graminis, on annual ryegrass.

Figure l0. Crown rust, Puccinia coronata, on perennial ryegrass

Figure 11. Stem rust, Puccinia graminis, on perennial ryegrass.

Figure 12, Stem rust, Puccinia poarum, on Kentucky bluegrass.

Figure 13. Powdery Mildew, Erysiphe graminis, on orchardgrass.

Figure 14. Ergot, Claviceps purpurea, on orchardgrass.

Figure 15. Mastigosporium leaf fleck, Mastigosporium spp., on orchardgrass.

Figure 16. Brown leafspot, Pyrenophora bromi, on smooth bromegrass.

Figure 17. Net blotch, Drechslera dictyoides, on annual ryegrass.

Figure 18. Eyespot, Heterosporium phlei, on timothy.

Figure 19. Selenophoma leafspot, Selenophoma donacis, on timothy.

Figure 20. Take-all. Gaeumannomyces gramlnis var. avenae, on perennial ryegrass.

Figure 21. Red thread, Corticium fuciforme, on perennial ryegrass.

Figure 22. Ryegrass mosaic virus on annual ryegrass, mild strain.

Figure 23. Cocksfoot mottle virus on orchardgrass.

Figure 24. Barley yellow dwarf virus on annual ryegrass (yellowing and dwarfing).

Figure 25. Cocksfoot mild mosaic virus on orchardgrass.

Figure 26. Ryegrass mosaic virus on annual ryegrass (severe strain).

The Authors

S. W. Braverman is affiliate associate professor of plant pathology and F. L.Lukezic is professor of plant pathology at The Pennsylvania State Univer­sity. K. E. Zeiders is research plant pathologist, Agri­cultural Research Service, United States Department of Agriculture. J. B. Wilson is adjunct professor of plant pathology at The Pennsylvania State University and former director of the U.S. Regional Pasture Re­search Laboratory, Agricultural Research Service, United States Department of Agriculture at Univer­sity Park.

The authors wish to express their appreciation to the scientists and others who have provided photo­graphs or otherwise contributed to the preparation of this publication. Dr. T. Tominaga, Sayama-Chi, Japan; Dr. T. Egli, CIBA-Geigy, Ltd., Basel, Switzerland; and Dr. D. Schmidt, Swiss Federal Re­search Station for Agronomy, Nyon, Switzerland provided photographs of bacterial diseases. Dr. C. J. O'Rourke, The Agricultural Institute, Dublin, Ire­land; and Dr. P. Weibull, Landskrona, Sweden, pro­vided photographs of fungus diseases. Photographs of the virus diseases are courtesy of Dr. P. L. Catherall, Welsh Plant Breeding Station, Aberystwyth, Dyfed County, Wales. Mrs. Teri-Anne Jordan assisted in preparation of the manuscript for the authors and editors.

Research reported in this publication is supported by funds from the Pennsylvania State Legislature, the United States Congress, and other government and private sources. Published by The Pennsylvania State Agricultural Experiment Station in cooperation with the U.S. Regional Pasture Research Laboratory, Uni­versity Park, Pennsylvania, and agricultural experi­ment stations of the Northeast Region. Authorized for publication 21 November, 1985.