© 2012 Plant Management Network. Accepted for publication 15 November 2012. Published 26 November 2012.

Anthracnose of Centipedegrass Turf Maria Tomaso-Peterson, Department of Biochemistry, Molecular Biology, Entomology, and Plant Pathology, Mississippi State University, Mississippi State, MS 39762; Jo Anne Crouch, Systematic Mycology and Microbiology Laboratory, USDA-ARS, Beltsville, MD 20705; and Clarissa Balbalian, Plant Disease and Nematode Diagnostic Service, Mississippi State University, Mississippi State, MS 39762 Corresponding author: Maria Tomaso-Peterson. [email protected] Tomaso-Peterson, M., Crouch, J. A., and Balbalian, C. 2012. Anthracnose of centipedegrass turf. Online. Applied Turfgrass Science doi:10.1094/ATS-2012-1126-01-MG.

Host: Centipedegrass [Eremochloa ophiuroides (Munro) Hack]. Disease: Anthracnose. Pathogen: Colletotrichum eremochloae J.A. Crouch & Tomaso-Peterson. Taxonomy Eukaryota; Fungi; Dikarya; Ascomycota; Pezizomycotina; Sordariomycetes; Hypocreomycetidae; Glomerellales; Glomerellaceae; Mitosporic Glomerellaceae; Colletotrichum; Colletotrichum eremochloae J.A. Crouch & Tomaso-Peterson. C. eremochloae was recently described based on DNA sequence data of modern cultures and archival fungarium specimens. The identification of C. eremochloae sp. nov., a pathogen of centipedegrass, was made based on phylogenetic evidence from four sequence markers, Apn2, Apn2/Mat1, Sod2, and ITS (1). C. eremochloae is closely related to C. sublineola, the pathogen of sorghum [Sorghum halapense (L.) Pers] and johnsongrass (S. vulgaris Pers.), but genealogical concordance supported their distinction as phylogenetic species (1). Currently, centipedegrass is the only host from which C. eremochloae has been isolated and confirmed as a pathogen. No teleomorph of C. eremochloae has been identified. Symptoms and Signs In home lawns, centipedegrass anthracnose occurs during the spring, March through early May, when the grass is transitioning out of winter dormancy. Turf recovery is observed in June when centipedegrass is growing under optimal conditions. Anthracnose symptoms have yet to be observed during fall transition. The crowns and oldest leaf sheaths of affected plants are necrotic while young leaf sheaths and leaves are chlorotic (Fig. 1A). Leaf blight progresses from the leaf tip to the base in older leaves. Foliar lesions are absent, and stolons and roots are visually unaffected. Various sizes of bright yellow or chlorotic, diffuse areas to irregular patches develop within centipedegrass lawns (Fig. 1B-C). Initially, the affected areas have a reduction in turf density and the leaves appear yellow or chlorotic (Fig. 1D). Necrotic areas subsequently progress within the sward. Bare areas develop and persist into the summer as the plants die (Fig. 1E-F). As a result, re-sodding of affected areas may be necessary to maintain turf uniformity and aesthetics. Unlike anthracnose in cool-season turfgrass species caused by C. cereale Manns, acervuli and setae may not be present on plant tissues affected by centipedegrass anthracnose. In recent collections of the fungus, no setae were observed in planta, but setae have been noted from preserved fungarum specimens such as those curated at the US National Fungus Collections (e.g., BPI 397271). Light gray, prominently septate mycelia of C. eremochloae colonize

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Fig. 1. (A) Necrotic crowns and leaf sheaths associated with foliar chlorosis are symptoms of centipedegrass anthracnose. (B, C) Home lawns exhibiting symptoms of bright yellow or chlorotic plants in irregular patches or diffuse areas. (D) Initial symptoms of centipedegrass anthracnose in a home lawn. (E, F) Disease progression results in the loss of centipedegrass.

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the epidermal tissue of the inner leaf sheaths of affected centipedegrass. Abundant hyphal appressoria are produced in association with the mycelium (Fig. 2) and may be used as a diagnostic feature.

Fig. 2. Hyphal appressoria microscopically observed on affected leaf sheaths of centipedegrass. Bar = 10 µm.

Host Range Current to this report, centipedegrass is the only known host of C. eremochloae. Geographic Distribution At present, anthracnose of centipedegrass caused by C. eremochloae is known from the US and China. The first affirmative diagnosis of centipedegrass anthracnose was from a home lawn in Hattiesburg, MS, in late March 2007 (1). During that same period, diseased centipedegrass from a lawn in coastal North Carolina was diagnosed with anthracnose at Mississippi State University (unpublished data). Four centipedegrass samples with anthracnose symptoms collected between 1923 and 1973 have also been identified as C. eremochloae. One sample (BPI 397271) was intercepted by US port authorities in Washington, DC, from a shipment of centipedegrass from China in 1923, two samples were collected in Quincy, FL, in 1944 (BPI 398409, BPI 596723), and one sample from Kailua, HI collected in 1973 (BPI 398410). Nucleotide sequences generated from DNA of these fungarium specimens shared 100% identity with modern, living samples of C. eremochloae (1). Therefore, it appears that centipedegrass anthracnose was brought into the United States via infected centipedegrass from China and may occur where centipedegrass is cultured in the southern US and Hawaii. Pathogen Isolation Centipedegrass home lawn samples were submitted to the Mississippi State University Plant Disease and Nematode Diagnostic Laboratory (http://msucares.com/lab) in March 2007. When the plants were examined microscopically, mycelium and hyphal appressoria characteristic of Colletotrichum sp. were observed. Infected leaf sheath segments (5 mm²) were surface disinfested in 0.6% NaOCl, rinsed three times in sterile, distilled water, and air dried in a laminar flow hood. Tissue segments were transferred to water agar (1.5% w/v). Hyphal tips emerging from leaf tissue were transferred with a sterile needle to potato dextrose agar (PDA, Fisher Scientific, Waltham, MA) and maintained in 24-h cool white fluorescent light at room temperature (~22°C). Pathogen Identification Morphology-based identification of C. eremochloae can be made directly from plant specimens, or from cultured isolates of the fungus grown on PDA at 20°C with a 12-h photoperiod for 7 days. In culture, the young colony has a moderate growth rate measuring 6.75 cm diameter at 7 days. The inner half of the colony appears gray and the outer half is white. The perimeter of the colony has a distinct fluorescent yellow pigment when cultured in 12 to 16 h of light per

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day (Fig. 3A). As the colony matures, the perimeter becomes whitish-gray with the majority of the colony appearing gray at 21 days (Fig. 3B). Conidia are produced from short, solitary conidiophores arising from mycelium (Fig. 4). Conidia production may be enhanced when axenic cultures are incubated in continuous light. Hyaline, single-celled, guttulate conidia appear falcate to fusiform, averaging 23.4 × 5.6 µm, similar in size to conidia of C. sublineola (4) (Fig. 4). The conidia produced by C. caudatum, the causal agent of tanso-byo disease on centipedegrass, produce a long filiform appendage from one of the conidial apices (3), readily distinguishing them from the conidia of C. eremochloae. Acervuli and setae have not been observed in culture. Abundant hyphal appressoria are produced both in planta and in vitro and are readily observed on the underside of the culture dish (Fig. 5). Hyphal appressoria are grayish-brown to dark brown in color, the edges are moderately lobed or irregular and average size dimensions are 12.5 × 12.5 µm (Fig. 5). The type strain of C. eremochloae is available through Centraalbureau voor Schimmelcultures culture collection, accessioned as CBS 129661 (1).

Fig. 3. Colony morphology of Colletotrichum eremochloae growing on potato dextrose agar in the presence of continual fluorescent light: (A) seven-day-old culture; (B) twenty-one-day-old culture.

Fig. 4. Single-celled conidia of Colletotrichum eremochloae. Bars = 10 µm.

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Fig. 5. Hyphal appressoria of Colletotrichum eremochloae. Observed in planta (left) and in vitro (right). Bar = 10 µm.

Morphology-based identification of C. eremochloae should be confirmed using sequence data. The concurrent use of molecular and morphological identification is recommended due to the similarity of spore, appressoria, and colony appearance to the closely related fungus C. sublineola. Nucleotide sequence analysis using the Apn2 or Sod2 markers is sufficient to differentiate C. eremechloae from C. sublineola, but the ITS marker cannot distinguish between these two species (1) (Table 1). Table 1. Molecular references for identification of Colletotrichum eremochloae causing centipedegrass anthracnose in Mississippi. Accession numberx

Sequence markery

Isolate

JQ478446

ITS

C01 (CBS 129661z)

JQ478448

Sod2

C01

JQ478461

Apn2/Mat1

C01

JQ478475

Apn2

C01

JQ478447

ITS

C05 (CBS 129664z)

JQ478449

Sod2

C05

JQ478462

Apn2/Mat1

C05

JQ478476

Apn2

C05

x Deposited in NCBI GenBank. y Nuclear rDNA internal transcribed spacer region (ITS), superoxide dismutase 2 z

(Sod2), DNA lyase gene (Apn2), mating gene (Mat1). Available from the Centraalbureau voor Schimmelcultures culture collection, Utrecht, The Netherlands.

Pathogen Storage Colonies of C. eremochloae maintain viability on PDA plates that are sealed with parafilm (American National Can, Greenwich, CT) and stored in plastic containers or sealed bags on the bench top. If the culture should become contaminated, appressoria growing on the bottom of the culture plate may be axenically transferred to fresh PDA to initiate new growth. To ensure fungal viability for extended periods of time, C. eremochloae isolates can be maintained at −20°C on filter paper (5). A hyphal plug of C. eremochloae, placed mycelium-side down, is transferred to sterile glass fiber filter paper resting on the surface of PDA. When mycelial growth colonizes the filter paper (10 to 14 days), it is removed from the PDA and air-dried in a laminar flow hood to maintain sterility (36 to 48 h). The colonized filter paper is then axenically cut into 5-mm² pieces and stored in parafilm-wrapped petri dishes or glass vials at −20°C. To initiate fungal growth, transfer an infested piece of filter paper onto PDA, mycelium-side down. The fungus grows equally well in the presence or absence of light at room temperature, with light inducing conidia production. C. eremochloae isolates described herein have maintained viability under these conditions since 2007.

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Pathogenicity Tests Pathogenicity tests are used to confirm C. eremochloae as the causal agent of centipedegrass anthracnose. To determine pathogenicity, a 5-ml spore solution (0.1× potato dextrose broth) 5 × 104 conidia/ml (2) is sprayed onto healthy ‘Common’ centipedegrass seedlings. Inoculated centipedegrass plants are covered for at least 10 days to maintain high humidity (Fig. 6) and incubated at 30°C with a 16-h day length in a growth chamber at which time leaf chlorosis may become evident (Fig. 7). After 21 days, hyphal appressoria may be observed on infected leaf sheaths and leaves using low magnification.

Fig. 6. Plant inoculation chamber used to incubate Colletotrichum eremochloae on centipedegrass to fulfill Koch’s Postulates.

Fig. 7. Foliar chlorosis of centipedegrass is evident 10 days following inoculation with a spore solution of Colletotrichum eremochloae. Uninoculated control plants (left) and inoculated plants (right).

Literature Cited 1. Crouch, J. A., and Tomaso-Peterson, M. 2012. Anthracnose disease of centipedegrass turf caused by Colletotrichum eremochloae, a new fungal species closely related to Colletotrichum sublineola. Mycologia 104:1085-1096. 2. Crouch, J. A., Clarke, B. B., White J. F. J., and Hillman, B. I. 2009. Systematic analysis of falcate-spored graminicolous Colletotrichum and a description of six new species of the fungus from warm season grasses. Mycologia 101:717-732. 3. Fuke, K., Hozumi, N., Enami, Y., Matsuura, K., and Tajimi, A. 2006. Anthracnose of centipedegrass caused by Colletotrichum caudatum. J. Gen. Plant Pathol. 72:7475. 4. Sutton, B. C. 1980. The Coelomycetes. Commonwealth Mycological Institute, Kew, London, UK. 5. Young, J. R., Tomaso-Peterson, M., Tredway, L. P., and de la Cerda, K. 2010. Occurrence and molecular identification of azoxystrobin-resistant Colletotrichum cereale isolates from golf course putting greens in the southern United States. Plant Dis. 94:751-757.

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