Stagonosporopsis chrysanthemi(MYCOLG)
EPPO Datasheet: Stagonosporopsis chrysanthemi
IDENTITY
Authority: (F. Stevens) P.W. Crous, N. Vaghefi & P.W.J. Taylor
Taxonomic position: Fungi: Ascomycota: Pezizomycotina: Dothideomycetes: Pleosporomycetidae: Pleosporales: Didymellaceae
Other scientific names: Ascochyta chrysanthemi F. Stevens, Didymella ligulicola (Baker, Dimock & Davis) von Arx, Mycosphaerella ligulicola Baker, Dimock & Davis, Phoma ligulicola var. ligulicola Boerema, Stagonosporopsis ligulicola var. ligulicola (Baker, Dimock & Davis) Aveskamp, Gruyter & Verkley
Common names in English: flower blight of chrysanthemum, ray blight of chrysanthemum, stem canker of chrysanthemum
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Notes on taxonomy and nomenclature
Not to be confused with Didymella chrysanthemi (Tassi) Garibaldi & Gullino (syn. Sphaerella chrysanthemi Tassi = Mycosphaerella chrysanthemi (Tassi) Tomilin). The name Didymella chrysanthemi has been erroneously used as synonymous to Didymella ligulicola (for example, see Punithalingam, 1980). This was based on a study by Garibaldi & Gullino (1971) that reported morphological identity of the ray blight pathogen described in the USA (at the time known as Didymella ligulicola) and a little-known fungus described in Italy (at the time known as Sphaerella chrysanthemi). Later, Walker & Baker (1983) studied the type specimen of both species and demonstrated that the American species was morphologically distinct and rejected the synonymy of Didymella ligulicola and Didymella chrysanthemi suggested by Garibaldi & Gullino (1971). For more information see Walker & Baker (1983) and Vaghefi et al. (2016b).
Taxonomy of the ray blight pathogen was revised in 2012 based on multi-locus phylogenetic analyses (Vaghefi et al., 2012). Currently, three Stagonosporopsis species are recognized as the cause of ray blight disease on Asteraceae. S. chrysanthemi is identified as the cause of ray blight on chrysanthemums while S. inoxydabilis and S. tanaceti are recognized as the cause of ray blight on other Asteraceae species in Europe and Australia, respectively (Vaghefi et al., 2016b).
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EPPO Code: MYCOLG
HOSTS 2023-11-20
The principal hosts of S. chrysanthemi are florists' chrysanthemums, Chrysanthemum x morifolium and C. indicum, the former being the original host on which this pathogen was described (Stevens, 1907). Later, annual chrysanthemum (Chrysanthemum carinatum), endives (Cichorium endivia), Dahlia (Dahlia pinnata and D. variabilis), globe artichokes (Cynara scolymus), lettuces (Lactuca sativa var. crispa), rudbeckia (Rudbeckia hirta), sunflowers (Helianthus annuus), and Zinnia (Zinnia elegans) were reported to be infected by artificial inoculation (Chesters & Blakeman, 1967). Pyrethrum (Tanacetum cinerariifolium) is also established as an experimental host for S. chrysanthemi after in vitro inoculation (Vaghefi et al., 2016b). Records established prior to the molecular reclassification of the pathogen in 2012 (Vaghefi et al., 2012) should be treated with caution as the identity of the isolates are not confirmed. For example, DAR 28714 previously identified as Didymella ligulicola (syn. S. chrysanthemi) on Zinnia (Walker & Baker, 1983) was later reclassified as Stagonosporopsis caricae (Marin-Felix et al., 2019).
Within the EPPO region the main potential hosts of S. chrysanthemi would be chrysanthemums grown under protected conditions and outdoors.
Host list: Chrysanthemum indicum, Chrysanthemum lavandulifolium, Chrysanthemum x morifolium, Lactuca sativaGEOGRAPHICAL DISTRIBUTION 2023-11-20
Though Boerema & Van Kesteren (1974) have suggested that S. chrysanthemi (D. ligulicola at the time) was present in Italy before introduction from the USA, Walker & Baker (1983) consider the fungi involved to be distinct (see Notes on taxonomy and nomenclature). On this basis, the fungus causing ray blight of chrysanthemum originated in North America and is believed not to have reached Europe until the 1960s (Walker & Baker, 1983).
Records dated prior to the molecular characterization of the pathogen (Vaghefi et al., 2012) should be treated with caution as the identity of the isolates cannot be confirmed and some reports of S. chrysanthemi were subsequently found to belong to other Stagonosporopsis species.
Africa: Kenya, Malawi, Tanzania, Tunisia, Zimbabwe
Asia: India (Andhra Pradesh), Israel, Japan (Honshu), Korea, Republic
North America: Canada (Ontario), Mexico, United States of America (California, Florida, Mississippi, New York, North Carolina, Ohio, Pennsylvania, South Dakota)
Oceania: Australia (New South Wales, Queensland, Tasmania), New Zealand, Papua New Guinea
BIOLOGY 2023-11-20
A principal source of primary inoculum contributing to ray blight epidemics is ascospores, which mature during the winter and early spring in pseudothecia on diseased host tissue (Baker et al., 1949). Ascospores are discharged throughout the season and carried in air currents (McCoy, 1973) causing scattered infections through much of the flowering season (Baker et al., 1949). Airborne ascospores are forcibly discharged (McCoy, 1973) and may disperse over longer distances and infect neighbouring fields (Blakeman & Hadley, 1968). Under experimental conditions on chrysanthemum stem and leaf tissue, pseudothecia developed in seven days at 20°C from the time of inoculation (McCoy et al., 1972). Lower temperatures delayed pseudothecia formation (25 days at 10°C) while temperatures higher than 25°C inhibited development of pseudothecia (McCoy et al., 1972). In naturally infected plants, pseudothecia were reported to be less abundant, but once plant samples were dried and incubated in the laboratory, an increase in the number of pseudothecia was observed within six to eight weeks (Baker et al., 1949).
Pycnidia, which form readily and abundantly on infected flower buds and peduncles and less so on stems and leaves, are also overwintering structures that can survive temperatures as low as -29°C (Baker et al., 1949). Upon artificial inoculation of chrysanthemum leaves and stems in the laboratory, pycnidia developed in three days at 26°C (optimal growth temperature) while pycnidial formation was inhibited at 30°C (McCoy et al., 1972). Pycnidia are produced under both dry and humid conditions (relative humidity of 2 to 85%) but splash-dispersal of pynidiospores occurs only in humid conditions (Baker et al., 1961). Pynidiospores exude in gelatinous drops (cirri) and are spread by rain splash, misting, infested tools and clothing or hands of workers. Infection by waterborne pynidiospores result in localized streaks of infection in the crop (Baker et al., 1949).
Given sufficient moisture, spores can infect petals within 6 hours and over a wide temperature range (6-30°C) but the optimal temperature for infection is 20-26°C (Baker et al., 1949). pynidiospores penetrate directly through or between epidermal cells, and a characteristic, much branched, short-celled mycelium quickly grows through the tissue, both intra- and intercellularly, causing a moist, brown decay (Baker et al., 1949). S. chrysanthemi produces a phytotoxin that is able to induce leaf symptoms similar to those caused by the pathogen itself (Schadler & Bateman, 1974). For more information see Stevens (1907), McCoy (1971), Vaghefi et al. (2016b).
The fungus can survive as epiphytic mycelium on chrysanthemum cuttings for 12 weeks (Chesters & Blakeman, 1966) and, more importantly, as pseudothecia and pycnidia, which are the overwintering organs and can survive over a broad range of temperatures on infected plants outside (-29 to 30°C) (Baker et al., 1949). Sclerotia have been reported to survive in the soil for eight weeks (Blakeman and Hornby, 1966), however, the identity of the isolates is not confirmed. Therefore, this record should be treated with caution as the original descriptions of the ray blight pathogen (Stevens, 1907; Walker & Baker, 1983) did not mention production of sclerotia.
DETECTION AND IDENTIFICATION 2023-12-05
Symptoms
Ray blight caused by S. chrysanthemi affects chrysanthemum under favorable conditions on potted plants, stock beds and cuttings grown in the field and protected areas (Engelhard 1984; Baker et al. 1949). Flowers and cuttings are particularly susceptible, but all plant parts are affected and present symptoms (Baker et al., 1949, 1961).
On cuttings
Cuttings are usually attacked at the terminal bud, from which point the infection spreads downwards to the whole plant. Unopened buds, bracts and stem tissue become darkened. On leaves, the fungus causes irregular brownish-black blotches, 2-3 cm across (Baker et al., 1949). Under favourable conditions, these rapidly coalesce and the leaf-rots. On stems, symptoms are associated with the points at which the diseased leaves adjoin, with wounds, or at the cutting base. During rooting, symptom development may be arrested, but diseased tissues remain on the plant and constitute a source of inoculum.
On adult plants
Stem lesions, which may girdle the stem and are often localized at the base or nodes, are associated with an abnormal appearance in the corresponding shoots, without the latter being contaminated by the fungus. This is due to production of a phytotoxin which induces drooping of terminal growth, makes leaves smaller, chlorotic and more or less brittle, and causes slight dwarfing.
On flowers
Following infection, spots develop, initially on one side of the blossom only. The spots appear reddish on light-coloured cultivars and brownish on darker ones. Infection subsequently spreads rapidly, and complete rotting of the flower head may occur, the infected florets sticking together. The fungus then grows down the peduncle, blackening and weakening the tissue, so that the head eventually droops and wilts (Baker et al., 1949).
Flower and leaf symptoms may be confused with those due to Botrytis cinerea, while rotting of cuttings resembles that due to B. cinerea or Pythium infection. In case of doubt, reproductive structures should be carefully examined. B. cinerea is distinguishable by the copious grey spores it produces. Septoria leaf spots (caused by Septoria spp.) have more definite lesion margins and the central areas have a characteristic sheen. For more information, see Stevens (1907), Nillsson (1963), Sauthoff (1963).
Morphology
Pycnidia are visible with a x15 hand lens as depressed, thin-walled, globose bodies of two sizes: small (72 x 180 µm) which are aggregated on the petals, and large (111 x 325 µm) which are scattered on the stems and leaves.
Pycnidiospores exude in short columns, are hyaline, continuous (10-40%) and septate (60-90%, usually with one septum, occasionally with more), ovoid to cylindrical with a pronounced tendency to irregularity and an extreme variability in dimensions; continuous spores 6-22 x 2.5-8 µm, mostly 8.5-13 x 3.5-5.5 µm; septate spores 9-23 x 3-6.5 µm, mostly 13-15.5 x 4-5 µm. For more information, see Sauthoff (1963), Blakeman & Hadley (1968), Boerema & Bollen (1975).
S. chrysanthemi shows phialidic ontogeny. Septation of the spores is a secondary process, related to temperature, and is probably a function of spore size. In culture, on oatmeal agar at 20-22°C, with a variable light-dark cycle, the majority of the pycnidiospores remain one-celled, 3.5-15 x 1.5-3.5 µm, mostly 4-8.5 x 2-3 µm.
S. chrysanthemi is homothallic and produces abundant pseudothecia in culture and on infected plants incubated in the laboratory for three to eight weeks (Baker et al., 1949; Vaghefi et al., 2016a,b). Pseudothecia are less commonly found on infected stems and flowers exposed outdoors during the winter, and are round and more erumpent than pycnidia, have dark-brown, thick-walled outer cells and are 96-224 µm in diameter.
Ascospores are hyaline to greyish, fusiform to elliptical, uniseptate, 12-16 x 4-6 µm.
Detection and inspection methods
Visual examination in a place or site of production is not effective at detecting S. chrysanthemi as the disease may be latent and symptoms may be confused with other diseases (EFSA, 2013). If S. chrysanthemi is suspected, samples should be sent to the laboratory for identification.
Isolation can be performed from all plant parts showing symptoms, but also seeds can be plated and the obtained culture characterized and identified. S. chrysanthemi can be easily isolated on pea agar and cherry decoction agar (Stevens, 1907; Boerema et al., 2004). For morphological studies, potato dextrose agar (Baker et al., 1949), oat meal agar or malt agar can be used (de Gruyter et al., 2002; Boerema et al., 2004).
Colony diameter on OA approximately 7 cm after seven days, regular to slightly irregular, colourless/greenish olivaceous to dull green olivaceous, often in a zonate pattern; with sparse to abundant, felted to floccose, white to pale olivaceous grey aerial mycelium; reverse grey olivaceous to fawn/hazel or olivaceous grey, more or less discolouring due to a diffusible pigment. On MA, slower growth, 5–6 cm in diameter after seven days, sometimes with a pale luteous pigment production. Only the anamorph of the pathogen is produced in culture (Van der Aa et al., 1990) with pycnidia developing on and in the agar medium (OA).
It should be noted that identification of S. chrysanthemi based on micro-morphological features requires expertise and is not always possible due to morphological similarity to other Stagonosporopsis spp. that cause ray blight on Asteraceae, thus, molecular sequencing is necessary for correct identification (Vaghefi et al., 2016b). A multiplex PCR test capable of differentiating the three Stagonosporopsis species associated with ray blight symptoms on Asteraceae has been developed (Vaghefi et al., 2016a).
An EPPO diagnostic protocol on Stagonosporopsis chrysanthemi is under development.
PATHWAYS FOR MOVEMENT 2023-11-20
S. chrysanthemi has a relatively low dispersal potential on its own. Pathways for movement include host plants for planting (including seed), cut flowers of host plants and soil and other growing media (EFSA, 2013). Infected cuttings of host plants can be a pathway for movement, as the pathogen is capable of surviving epiphytically on symptomless plant cuttings. Soil and other growing media attached to roots can also be a source of inoculum and therefore potted plants of hosts, and non-hosts with infected soil, can be a pathway for movement.
PEST SIGNIFICANCE 2023-11-20
Economic impact
There are no recent reports of the effects of S. chrysanthemi on chrysanthemum crops (EFSA, 2013). Historically, the disease has caused economic impacts, and these are recorded for North America. The disease was recorded in North Carolina (USA) in 1904 and remained localized and of little importance until the late 1940s when, concurrent with the intensification of chrysanthemum flower and pot plant production, it began to cause serious losses throughout the distribution range. In North America from 1946-47, the disease was described as destructive (EFSA, 2013). In 1975, in Connecticut, the disease was reported to be particularly important on chrysanthemum cuttings in propagating benches under mist and 50% losses occurred.
S. chrysanthemi is present in the EU sporadically but there are no reports of the pathogen causing damage on chrysanthemum crops in the EU (EFSA, 2013). Additionally, EFSA (2013) conclude that the overall impact of S. chrysanthemi in the EU is minor.
Control
Cultural practices in places of production can reduce the spread and infection by S. chrysanthemi. EFSA (2013) highlight that the use of use of pathogen-free propagation material, monitoring of the crop, avoiding overhead irrigation, proper management of waste plant material, soil disinfestation with steam, use of clean irrigation water and general hygiene measures can be used to avoid occurrence and spread.
Rapid removal of infected plants can help reduce disease spread.
In Europe, historically, the disease has been controlled successfully with benomyl. However, repeated and excessive usage of this fungicide (and thiophanate-methyl) over a number of years has led to the development of resistant isolates (Punithalingam, 1980). EFSA (2013) note that recent information on fungicides for S. chrysanthemi is poor.
Currently there is no biological control method available (CABI, 2012).
Phytosanitary risk
There is the potential for further establishment and spread of S. chrysanthemi in the EPPO region (EFSA, 2013). Host plants of S. chrysanthemi are popular ornamentals in the EPPO region and are widely grown in public and private gardens. The crop is cultivated in open fields and greenhouses.The Netherlands and Italy are the main European producers of chrysanthemum cut flowers whereas potted plants are produced throughout the EPPO region. Maintaining the current import procedures, production practices and control measures can keep the phytosanitary risk to a minimal level.
PHYTOSANITARY MEASURES 2023-11-20
Certified host plant material should be used (see EPPO, 2002). In countries where the disease occurs, growing-season inspections should be carried out, especially during rooting of cuttings, but also on mother plants and at flowering. Rooted or unrooted cuttings should come from rooting beds or plants, respectively, which were found free from S. chrysanthemi during the last growing season. Symptoms may develop in transit on blooms which are apparently healthy when cut. For imported consignments and inspections of places of production of chrysanthemum plants, visual inspection should be conducted and if suspected symptoms are detected, a sample should be sent for laboratory testing.
REFERENCES 2023-12-05
Baker KF, Dimock A & Davis LH (1949) Life history and control of the Ascochyta ray blight of chrysanthemum. Phytopathology 39, 789-805.
Blakeman JP & Hadley G (1968) The pattern of asexual sporulation in Mycosphaerella ligulicola. Transactions of the British Mycological Society 51, 643-651.
Blakeman JP & Hornby D (1966) The persistence of Colletotrichum coccodes and Mycosphaerella ligulicola in soil, with special reference to sclerotia and conidia. Transactions of the British Mycological Society 49, 227-240.
Boerema GH, de Gruyter J, Noordeloos ME & Hamers MEC (2004) Phoma identification manual. Differentiation of specific and infra-specific taxa in culture. CABI publishing, Wallingford, UK, 470 pp.
Boerema GH & Bollen GJ (1975) Conidiogenesis and conidial septation as differentiating criteria between Phoma and Ascochyta. Persoonia 8, 111-144.
Boerema GH & Van Kesteren HA (1974) [Some special fungal infections VI]. Gewasbescherming 5, 119-125.
CAB International (Centre for Agricultural Bioscience International), online. Datasheets: Didymella ligulicola. Crop Protection Compendium. http://www.cabi.org/cpc/ (accessed on 13 of December 2012).
Chesters C & Blakeman J (1967) Host range and variation in virulence of Mycosphaerella ligulicola. Annals of Applied Biology 60, 385-90.
EFSA Panel on Plant Health (PLH) (2013) Scientific opinion on the risks to plant health posed by Stagonosporopsis chrysanthemi (Stevens) Crous, Vaghefi and Taylor [Didymella ligulicola (Baker, Dimock and Davis) Arx var. ligulicola; syn. Didymella ligulicola (Baker, Dimock and Davis) Arx] in the EU territory, with identification and evaluation of risk reduction options. EFSA Journal 11(10), 3376. https://efsa.onlinelibrary.wiley.com/doi/epdf/10.2903/j.efsa.2013.3376
EPPO (2002) EPPO PM 4/6(2) Production of healthy plants for planting, Certification scheme for chrysanthemum. EPPO Bulletin 32, 49-53.
Garibaldi A & Gullino G (1971) Brevi notizie sulla presenza in Italia dell’ascochitosi del crisantemo. L’Agricoltura Italiana 71 (N.S. 26), 21-290.
Hahn W & Schmatz R (1980) Diagnosis and control of Ascochyta disease of Chrysanthemum. Nachrichtenblatt für den Pflanzenschutz in der DDR 34, 189-192.
IMI (1993) Distribution Maps of Plant Diseases No. 406 (edition 3). CAB International, Wallingford, UK.
Kamemoto H & Nakasone HY (1953) Controlling chrysanthemum flowering by altering daylength.
Marin-Felix Y et al. (2019) Genera of phytopathogenic fungi: GOPHY 3. Studies in Mycology 94, 1-124.
McCoy RE (1971) Epidemiology of chrysanthemum Ascochyta blight. PhD Thesis, Cornell University, Ithaca, USA.
McCoy RE, Horst RK, Dimock AW (1972) Environmental factors regulating sexual and asexual reproduction by Mycosphaerella ligulicola, Phytopathology 62, 1188-1195.
McCoy R (1973) Ballistics of Mycosphaerella ligulicola ascospore discharge. Phytopathology 63, 793-794.
Nillsson L (1963) [Black rot (ascochyta rot) of chrysanthemum]. Växtskyddsnotiser 27, 8-15.
Punithalingam E (1980) Didymella chrysanthemi. CMI Descriptions of Pathogenic Fungi and Bacteria No. 662. CAB International, Wallingford, UK.
Sauthoff W (1963) [Didymella ligulicola (Baker, Dimock et Davis) v. Arx as a pathogen of chrysanthemum in Germany]. Phytopathologische Zeitschrift 48, 240-250.
Schadler DL & Bateman DF (1974) Ascochyta chrysanthemi toxin: production and properties, Phytopathology 64, 779-784.
Vaghefi N, Pethybridge SJ, Ford R, Nicolas ME, Crous PW & Taylor PWJ (2012) Stagonosporopsis spp. associated with ray blight disease of Asteraceae. Australasian Plant Pathology 41, 675-686.
Vaghefi N, Hay FS, Pethybridge SJ, Ford R & Taylor PWJ (2016a) Development of a multiplex PCR diagnostic assay for the detection of Stagonosporopsis species associated with ray blight of Asteraceae. European Journal of Plant Pathology 146, 581-595.
Vaghefi N, Pethybridge SJ, Hay FS, Ford R, Nicolas ME & Taylor PWJ (2016b) Revisiting Stagonosporopsis species associated with chrysanthemum and pyrethrum ray blight, Australasian Plant Pathology 45, 561-570.
Van der Aa H, Noordeloos M & de Gruyter J (1990) Species concepts in some larger genera of the Coelomycetes. Studies in Mycology 32, 3-19.
Walker J & Baker KF (1983) The correct binomial for the chrysanthemum ray blight pathogen in relation to its geographical distribution. Transactions of the British Mycological Society 80, 31-38.
ACKNOWLEDGEMENTS 2023-11-20
This datasheet was extensively revised in 2023 by Niloofar Vaghefi, University of Melbourne. Her valuable contribution is gratefully acknowledged.
How to cite this datasheet?
Datasheet history 2023-11-20
This datasheet was first published in the EPPO Bulletin in 1983 and revised in the two editions of 'Quarantine Pests for Europe' in 1992 and 1997, as well as in 2023. It is now maintained in an electronic format in the EPPO Global Database. The sections on 'Identity', ‘Hosts’, and 'Geographical distribution' are automatically updated from the database. For other sections, the date of last revision is indicated on the right.
CABI/EPPO (1992/1997) Quarantine Pests for Europe (1st and 2nd edition). CABI, Wallingford (GB).
EPPO (1982) Data Sheet on Quarantine Organisms no 66: Didymella chrysanthemi. EPPO Bulletin 12(1), 41-46. https://onlinelibrary.wiley.com/doi/abs/10.1111/j.1365-2338.1982.tb01954.x
