Dead Spot of Creeping Bentgrass and Hybrid Bermudagrass

John E. Kaminski, Department of Plant Science, University of Connecticut, Storrs 06269: and Peter H. Dernoeden, Department of Natural Resource Sciences and Landscape Architecture, University of Maryland, College Park 20742

Corresponding author: John Kaminski. john.kaminski@uconn.edu

Disease: Dead spot (syn.= Bentgrass dead spot, Bermudagrass dead spot).

Primary Hosts: Creeping bentgrass (Agrostis stolonifera L.) and hybrid bermudagrass (Cynodon dactylon (L.) Pers. × C. transvaalensis Burtt-Davy).

Pathogen: Ophiosphaerella agrostis Dernoeden, M. P. S. Câmara, N. R. O’Neill, van Berkum et M. E. Palm.

Taxonomy

Ophiosphaerella agrostis Dernoeden, M. P. S. Câmara, N. R. O’Neill, van Berkum et M. E. Palm was first recognized as a new species within the genus Ophiosphaerella in 1999 and described in 2000 (1,2). Molecular and morphological studies revealed differences between this new species and three other Ophiosphaerella spp. (i.e., O. herpotricha, O. korrae, and O. narmari) associated with diseases of turf (1).

Symptoms and Signs

Dead spot is a disease of relatively young stands (≤ 6 years old) of Agrostis spp. and hybrid bermudagrass (Cynodon dactylon (L.) Pers. × C. transvaalensis Burtt-Davy) turf grown on high sand content golf greens and tees in various regions of the United States (6). The disease has been observed in the spring on bermudagrass greens that were overseeded with roughstalk bluegrass (Poa trivialis L.) (8), but the pathogenicity of O. agrostis on roughstalk bluegrass has not been documented formally. Dead spot also can develop on older, sand-based putting greens that have been seeded following fumigation with methyl bromide. The disease is not known to affect turfgrass grown on native soils.

On close-mown Agrostis or hybrid Cynodon golf greens, dead spot initially appears as small, reddish-brown spots approximately 1 cm in diameter, which can increase up to 10 cm in diameter (Fig. 1) (2,5). In Agrostis spp., initial infection centers often are confused with other turfgrass diseases such as dollar spot (Sclerotinia homoeocarpa F. T. Bennett), copper spot (Gloeocercospora sorghi Bain and Edgerton ex Deighton), and Microdochium patch (Microdochium nivale [Fr.] Samuels and I. C. Hallett). Spots also can mimic damage from black cutworms (Agrotis ipsilon Hufnagel) or ball-marks, which commonly are found on cool- and warm-season turf maintained as putting greens (Fig. 2). As the disease progresses, grass in the center of infected spots becomes tan, while leaves in the active, outer edge remain reddish-brown. Dead rings with living turf in the center (frog-eyes) seldom occur. Removal of leaf sheaths reveals that the crown and roots of dying plants are discolored or necrotic. Patches may be distributed throughout greens or localized, and the spots or patches generally only coalesce during severe epidemics (Fig. 3). Recovery of bentgrass is slow, as stolon growth into dead patches appears restrained or inhibited. Recovery occurs from tillering of healthy plants along the periphery of patches. On hybrid bermudagrass putting greens, turf recovers quickly when temperatures become suitable for vigorous bermudagrass growth.


Fig. 1. Ophiosphaerella agrostis infection centers on a creeping bentgrass green with tan or brown dead tissue surrounded by a reddish-brown active edge.		Fig. 2. Dead spot often is confused with damage from other diseases and insect pests including black cutworms (arrow) surviving in an aerification hole.

Fig. 3. Infection centers on creeping bentgrass can be found throughout the putting surface, but generally only coalesce under conditions of severe disease pressure.

Foliar mycelium is not observed in the field, but when diseased plants are incubated under high humidity for 3 to 5 days, a white to pale-pink foliar mycelium may develop (Fig. 4). Numerous sexual fruiting bodies (pseudothecia) often can be found embedded in necrotic leaf, sheath and stolon tissues (Fig. 5). When pseudothecia are not present, the disease can presumptively be identified by the presence of dark-brown to black hyphal masses found near or on the nodes of creeping bentgrass stolons (Fig. 6). Isolation of the pathogen in pure culture, however, is needed for a positive diagnosis. In bermudagrass, hyphae generally are limited to the interior cells of roots (8).

Fig. 4. Mycelium is not found in the field. Rose-quartz colored mycelia can develop following several days of incubating diseased plants under high humidity (left) or on agar media (right).


Fig. 5. Sexual fruiting bodies (pseudothecia) often are found embedded within necrotic leaves, leaf sheaths, and stolon tissues of infected plants (creeping bentgrass shown).		Fig. 6. Hyphal mats often can be found on or adjacent to the nodes of bentgrass stolons, but generally are less prominent in the internodal region of stolons.

In bentgrass grown in the Mid-Atlantic region, the disease may be active between May and December. However, new infection centers most often appear during the summer months when soil temperatures reach ≥ 20°C and mean daily relative humidity is low (≤ 80%) (5). On bermudagrass putting greens the disease may appear as early as March, but generally disappears during the summer months when conditions are optimum for growth of this warm-season turfgrass. Active dead spot infection centers generally appear in areas with full sun and good air circulation. In particular, O. agrostis infection centers often appear first along ridges, on mounds, and on south-facing slopes of greens. These areas are particularly prone to higher soil temperatures and often are the first to exhibit symptoms of drought stress. While extended leaf wetness durations appear to influence O. agrostis infection, heat and drought stress are the primary environmental factors that promote symptom expression and disease severity. Active disease symptoms are not observed during the winter months; however, infection centers that have not fully recovered in the autumn will remain visible until the following summer.

Host Range

Several cool-season and a single warm-season turfgrass species have been identified to date as hosts of O. agrostis (6,8). Cool-season hosts for O. agrostis include creeping bentgrass, colonial bentgrass (A. capillaris L.), velvet bentgrass (A. canina L.), and roughstalk bluegrass (6,8). While all bentgrass cultivars may be infected by O. agrostis, varying levels of cultivar susceptibility within and among bentgrass species have been reported, and no cultivar is immune (6). In southern regions of the United States, O. agrostis causes a similar springtime disease on hybrid bermudagrass putting greens that usually were overseeded with Poa trivialis prior to winter (6,8) (Fig. 7). In the aforementioned situation, the disease may first appear on overseeded roughstalk bluegrass during spring green-up of the bermudagrass (8). Although creeping bentgrass and hybrid bermudagrass are considered the primary hosts for the pathogen, the other turfgrass species listed above may also be severely damaged. Observations in the field indicate that the disease can be extensive on velvet and colonial bentgrasses as well as roughstalk bluegrass; however, the pathogenicity of O. agrostis on these turfgrass species has not been examined under controlled experiments (i.e., Koch’s postulates). Isolates from various turfgrass species do not appear to be species specific and O. agrostis strains are capable of infecting both hybrid bermudagrass and creeping bentgrass (5).

Fig. 7. Dead spot symptoms on a hybrid bermudagrass putting green overseeded with roughstalk bluegrass (photos courtesy Bruce Martin).

Geographic Distribution

Dead spot first was reported as a disease of creeping bentgrass after being found on a golf course putting green in Maryland in 1998. By the end of that year, the disease had been identified on ten golf courses in five states including Illinois, Maryland, Ohio, Pennsylvania, and Virginia (2). Dead spot has been found on creeping bentgrass as far north as Michigan, as far west as Missouri, and along the eastern seaboard of the United States from Massachusetts to Georgia. In addition to creeping bentgrass, the pathogen has been confirmed in or isolated from diseased hybrid bermudagrass putting greens in Florida, South Carolina and Texas (6,8; B. Martin, personal communication). Dead spot was documented in at least fifteen states between 1998 and 2004 (5; L. Burpee and B. Martin, personal communication), and O. agrostis likely can be found throughout the eastern half of the United States (Fig. 8).

Fig. 8. Confirmed cases of dead spot on creeping bentgrass and hybrid bermudagrass.

Pathogen Isolation

Isolation of O. agrostis from various tissues may be difficult depending on the season. During periods of peak disease activity (i.e., summer for bentgrass species or spring for hybrid bermudagrass and roughstalk bluegrass), the pathogen may be isolated from all plant parts including leaves, sheaths, crowns, roots, and stolons. When disease activity is arrested due to cold temperatures, however, the pathogen is most easily isolated from nodes on discolored stolons (Fig. 9). Isolation from infected plant tissue may be accomplished by sectioning (3 to 6 mm) tissue, surface disinfesting in 0.5% sodium hypochlorite (household bleach) for 60 sec and washing three times for 30 sec in sterile distilled water. Sectioned tissue should be dried on sterile filter paper and incubated on antibiotic (streptomycin sulfate at 0.5 g/liter) water agar in a dark growth chamber maintained at 25°C to 30°C (7). Fungal colonies free from bacterial and fungal contaminants should be transferred to potato dextrose agar (PDA) and incubated for 3 days (7). Single ascospore isolates also may be obtained using methods previously described (7).

Fig. 9. The pathogen can be isolated from nodes of discolored bentgrass stolons throughout the summer and winter months.

Pathogen Identification

In the absence of fungicide use, pseudothecia often are produced in abundance and serve as the primary means for identifying the pathogen. Pseudothecia are black and range from 130 to 350 µm in diameter, with necks 40 to 160 µm long (Fig. 10) (1). Ascospores are long and slender, pale-brown, and range from 70 to 190 × 2.0 to 2.5 µm with 6 to 15 septations (Fig. 11) (1,4). Ascospores are spirally twisted near the middle of bitunicate (two wall layers) asci and appear light-brown en masse (Fig. 12) (1). Dark-brown to black, ectotrophic hyphae and mycelial mats appear near or on stolon nodes and stem bases of Agrostis (Fig. 13). Dark-brown hyphae can be observed within roots of Cynodon, but may not be evident on the surface of roots or stolons (8).


Fig. 10. Pseudothecia can be found in abundance within various parts of infected plants and hundreds or thousands of ascospores (arrows) can be produced within a small area.		Fig. 11. Ascospores vary in length from 70 to 190 µm and may have 6 to 15 septations (ascospore stained with acid fuchsin).

Fig. 12. Ascospores are spirally twisted (A) within bitunicate asci (B) and appear light brown en masse (C).

Fig. 13. Hyphal mats often are found near the nodes of stolons. Hyphae produce simple hyphopodia (arrow), which can directly penetrate tissues.

Colonies of O. agrostis typically appear rose-quartz or pink, and less frequently gray, olive-gray or buff when incubated for 3 to 10 days on PDA (Fig. 14). The optimum O. agrostis growth rate is between 4 and 6 mm/day at temperatures between 25°C and 30°C (7). No asexual stage has been observed, therefore, conidia are not produced in culture. Pseudothecia are seldom produced in pure culture. The production of pseudothecia can be induced on an O. agrostis-infested mixture of tall fescue (Festuca arundinacea Schreb.) seed and wheat (Triticum aestivum L.) bran (1:1 v/v) (6). Pseudothecia develop within 4 to 7 days when incubated under constant light at 25°C (5,7).

Fig. 14. When grown on potato dextrose agar, mycelium characteristics vary among isolates, but most isolates exhibit a rose-quartz colony color, while others appear gray or olive-gray.

When pseudothecia are not present, a molecular-based method for rapid identification of O. agrostis is available (5). This polymerase chain reaction (PCR)-based technique can detect O. agrostis DNA from isolates grown in pure culture and from infected bentgrass or bermudagrass field samples. For labs equipped to perform this molecular procedure, a positive identification can be accomplished in as little as four hours.

Pathogen Storage

Isolates of the pathogen have been stored for up to 5 years on PDA slants incubated at 4°C without significant loss of viability. Another effective method includes the storage of O. agrostis isolates on PDA agar plugs (3 mm in diameter) in sterile, distilled water at 4°C. Although sterile filter paper commonly is used for storing other plant pathogens (3), this is not a viable long-term method of O. agrostis storage. Retrieval of O. agrostis isolates stored on filter paper generally was unsuccessful after incubation for only a few months (Kaminski, unpublished).

Pathogenicity Tests

Limited testing has revealed that O. agrostis isolates appear to be non-host specific. For instance, isolates from A. stolonifera were capable of infecting A. canina and A. capillaris under field conditions (6); and an isolate from hybrid bermudagrass was capable of infecting and killing creeping bentgrass in a greenhouse (Kaminski, unpublished). Differences in disease severity were reported among various Agrostis spp. and cultivars, and various cultivars within the same bentgrass species exhibited varying levels of disease severity (6). Although the impact of different isolates or strains of the pathogen on different host species is unknown, DNA fingerprinting of O. agrostis isolates revealed no correlation between isolate grouping and host species (7).

Inoculation of turfgrasses with tall fescue seed/wheat bran mixture (1:1 v/v) infested with O. agrostis has been successful in growth chamber and field situations (6). In growth chambers, inoculated plants should be placed in plastic bags to encourage high levels of relative humidity and incubated at 25°C. After 2 to 5 days of incubation, mycelia will begin to grow radially from the inoculum to plants resulting in infection and death of plants in about 7 days. The fungus may directly penetrate tissues or gain ingress through open stomata. Under the aforementioned conditions, pseudothecia may develop within 3 weeks in the presence of light. Field inoculations can be accomplished by placing the aforementioned inoculum (0.5 g) in a small hole (1 to 2 cm in diameter) in the thatch layer (Fig. 15). In field studies conducted on bentgrass stands less than three years old, disease symptoms developed regardless of whether the area was inoculated in the spring or the previous autumn (5).

Fig. 15. Field inoculation can be performed in the autumn or spring and is accomplished by placing an O. agrostis-infested tall fescue/wheat bran mixture in the thatch layer or at the soil surface.

Acknowledgments

We thank Bruce Martin for providing photos of dead spot on bermudagrass.

Literature Cited

1. Câmara, M. P. S., O’Neill, N. R., van Berkum, P., Dernoeden, P. H., and Palm, M. E. 2000. Ophiosphaerella agrostis sp. nov. and its relationship to other species of Ophiosphaerella. Mycologia 92:317-325.

2. Dernoeden, P. H., O’Neill, N. R., Câmara, M. P. S., and Feng, Y. 1999. A new disease of Agrostis palustris incited by an undescribed species of Ophiosphaerella. Plant Dis. 83:397.

3. Harmon, P. F., Dunkle, L. D., and Latin, R. 2003. A rapid PCR-based method for the detection of Magnaporthe oryzae from infected perennial ryegrass. Plant Dis. 87:1072-1076.

4. Kaminski, J. E. 2001. Growth, pseudothecia production, and ascospore germination of Ophiosphaerella agrostis and cultivar susceptibility and geographic distribution of bentgrass dead spot. M.S. Thesis. University of Maryland, College Park.

5. Kaminski, J. E. 2004. Biology of Ophiosphaerella agrostis, epidemiology of dead spot and a molecular description of the pathogen. Ph.D. Diss. University of Maryland, College Park.

6. Kaminski, J. E., and Dernoeden, P. H. 2002. Geographic distribution, cultivar susceptibility, and field observations on bentgrass dead spot. Plant Dis. 86:1253-1259.

7. Kaminski, J. E., Dernoeden, P. H., O’Neill, N. R., and Momen, B. 2002. Reactivation of bentgrass dead spot and growth, pseudothecia production, and ascospore germination of Ophiosphaerella agrostis. Plant Dis. 86:1290-1296.

8. Krausz, J. P., White, R. H., Foerster, W., Tisserat, N. A., and Dernoeden, P. H. 2001. Bermudagrass dead spot: A new disease of bermudagrass caused by Ophiosphaerella agrostis. Plant Dis. 85:1286.