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Plasmopara Halstedii

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Plasmopara Halstedii Bulletin OEPP/EPPO Bulletin (2014) 44 (3), 350–359 ISSN 0250-8052. DOI: 10.1111/epp.12160 European and Mediterranean Plant Protection Organization Organisation Europe´enne et Me´diterrane´enne pour la Protection des Plantes PM 7/85 (2) Diagnostics Diagnostic PM 7/85 (2) Plasmopara halstedii Specific scope Specific approval and amendment This Standard describes a diagnostic protocol for Plasmopara Approved in 2008–09. Revised in 2014–06 halstedii.1 Synonyms: Plasmopara helianthi Novotel’ nova Introduction Taxonomic position: Oomycetes; Peronosporales; The oomycete Plasmopara halstedii, which causes downy EPPO code: PLASHA mildew on cultivated sunflower, is known on every conti- Phytosanitary categorization: EU Annex designation:II/A2 nent of the world. It is considered as a major disease in all sunflower producing countries in Europe (Sackton, 1981). Detection In addition, fungicide tolerant isolates as well as new path- ogenic races have appeared and are spreading, and repre- Plasmopara halstedii may induce symptoms of various senting a threat to sunflower crops (Roeckel-Drevet et al., kinds depending on the age of tissue, level of inoculum, 2003). cultivar reaction and environmental conditions (moisture The significant hosts are the wild and cultivated species and temperature). Symptoms result from systemic or local of Helianthus, including sunflower (Leppik, 1966; Novo- infections; those being caused by systemic infections being tel’nova, 1966), but over 100 host species from a wide more typical. range of genera in the family Asteraceae have been Systemically infected sunflower plants may have some reported. Wild Asteraceae hosts (e.g. species of the genera degree of stunting and the leaves show pale green or chlo- Helianthus, Artemisia, Xanthium, etc.) may also occur rotic mottling which spreads along the main veins and over widely in the corresponding areas but their potential as res- the lamella (Fig. 1). Young leaves of severely affected ervoirs of the pest is not yet confirmed (Viranyi, 1984; plants often become entirely chlorotic, curl downward, and Walcz et al., 2000). become rigid and thick (leaf chlorosis is illustrated in This pathogen is mainly soil-borne and seed-borne (Basa- Fig. 2). Under moist conditions, a white downy growth varju et al., 2004) but may also be wind-borne. composed of sporangiophores and sporangia of the oomyce- In the EPPO region the pathogen is present wherever te develop on the lower leaf surface and observations have sunflowers are grown. Further information on the biology shown that their extent strictly corresponds to the chlorotic and epidemiology of the pest can be found in the EPPO areas on the upper leaf surface (Fig. 3). Due to internode data sheet on P. halstedii (EPPO/CABI, 1997). Details on shortening, an infected sunflower may have a cabbage-like its geographical distribution can be found in the EPPO appearance. Plant Quarantine Retrieval System (PQR) (2007). Heads of infected sunflower plants have a reduced size and face upwards (Fig. 1), bearing no or a limited number of seeds, the viability of which is poor. The economical Identity loss can be considerable. Name: Plasmopara halstedii (Farlow) Berlese & de Toni The root system of infected sunflower is underdeveloped, with significant reductions in secondary root formation and 1Use of names of chemicals or equipment in these EPPO Standards with a dark brown appearance on their surface. implies no approval of them to the exclusion of others that may also be suitable. 350 ª 2014 OEPP/EPPO, Bulletin OEPP/EPPO Bulletin 44, 350–359 Plasmopara halstedii 351 ised by white sporulation on the cotyledon leaves and the stems of seeds at cotyledon stage (Figs 4 and 5). Local leaf infections causing angular leaf spotting may also be observed (small, angular, pale green spots, delim- ited by the veins). Systemic infection (pathogen growing through the petioles into the stem) may subsequently occur when environmental conditions are favourable and the infection pressure is high (Ratai-Vida, 1996). Before fructification, P. halstedii may be confused with herbicide phytotoxicity. After fructification, it may be con- fused with Albugo tragopogonis but in this case typical large chlorotic blister-like pustules form on the upper side of leaves. Such deformation of the foliage does not occur with P. halstedii. Fig. 1 Field symptoms of P. halstedii on sunflower plant. Leaf chlorosis, stunting and upturned head (Photo: Hedvig Komjati). Identification This oomycete is an obligatory biotrophic plant pathogen and morphological identification is possible only in vivo,on the host plant. Consequently, morphological identification from seed and soil samples requires a bioassay to be con- ducted. PCR tests have also been developed for identifica- tion from seed and may be adapted for identification of in planta grown structures (sporangiophores bearing sporan- gia). An ELISA test is described in the literature (Bouterige et al., 2000), but there is little experience in the EPPO region with using this method. Therefore ELISA is not rec- ommended. Detection from seed is difficult because seed infection in lots is highly variable and sometimes very low due to blending/dilution of seeds with various infection levels. The bioassay tests described in this section and the PCR tests give an indication of the number of seeds that can be tested Fig. 2 Leaf chlorosis on sunflower cotyledon leaves – plants 5–6 days in a single test. The total number of seeds to be tested in older than those from Fig. 4 (Photo: Hedvig Komjati). order to detect the pest at different infection levels in a lot needs to be determined statistically (useful guidance is given in table 1 and 2 of ISPM 31 Methodologies for sam- Other, less common symptoms associated with systemic pling of consignments (IPPC, 2008)). infection include damping-off of seedlings, pith discolor- ation of the stem and/or capitulum, disturbance of inflores- Bioassay cence, twisted leaves and basal gall (Sackton, 1981). Furthermore, systemic downy mildew infection may be Detection from seed localized to lower stem tissues (cotyledon or hypocotyl Detection from seed is based on the observation of signs of infections (Ljubich & Gulya, 1988; Viranyi & Gulya, sporulation on germinated seedlings at cotyledon leaf stage. 1996)). In moist conditions, such infections are character- The appearance of any other symptoms such as distortion Fig. 3 White downy growth developed on the lower leaf surface (A) and corresponding to the chlorotic areas on the upper leaf surface (B) Photo Courtesy The Food and Environment Research Agency (Fera), Crown A B Copyright. ª 2014 OEPP/EPPO, Bulletin OEPP/EPPO Bulletin 44, 350–359 352 Diagnostics water saturated. In such conditions motile zoospores migrate through soil-pores to the roots (Gulya et al., 1997). A fast and comparatively simple bioassay method was developed for diagnostic purposes, based on the method of Gulya & Radi (2002). Soil samples from the suspected field should be taken after the vegetative period (late autumn at the earliest) or before sowing during springtime the follow- ing year. Samples should be taken from the rhizosphere level (up to 20 cm) as potentially infected plant debris may have been incorporated onto the soil surface. Detection from soil is based on the observation of signs of sporulation in germinated seedlings at cotyledon leaf stage. The method is presented in Appendix 1. Morphology Fig. 4 Sporulation of Plasmopara halstedii of surface of artificially inoculated sunflower cotyledon leaves (Photo: Hedvig Komjati). Morphological identification is possible by microscope. or discoloration of young plantlets is not a specific sign of Mycelium infection. In addition, ensuring good growth conditions Mycelium is composed of intercellular, colourless, aseptate (temperature of 16–17°C and relative humidity of 95 to hyphae 6–20 lm diameter, often irregularly shaped and 100%) is critical for the detection of P. halstedii. swollen, bearing small, rounded, vesicular haustoria, The method is presented in Appendix 1. 5–10 lm in diameter, growing in all plant tissues. Detection from soil Sporangiophores This test is recommended for testing soil from fields where Sporangiophores are hypophyllous, or occasionally epiphyl- infestation is suspected. A soil sample with a standard rate lous, arborescent, 300–450 (750) lm 9 7–14 lm, obconi- À1 of at least 1 500 mL soil ha should be collected from at cal or pointed at the base, branching in the upper half, with À1 least 100 cores ha preferably in a rectangular grid of not the apex of the branching axis frequently swollen: branches <5 m in width and not more than 20 m in length between in the form of a whorl of 7–8, produced monopodially at sampling points covering the entire field. The whole sample right angles to the main sporangiophore axis, each with 2–5 should be used for the test. The detection from soil is based secondary branches 40–86 lm long, bearing 3–5 tips, on the ability of the pathogen to infect young sunflower 8–15 lm long, diverging at right angles (Figs 6 and 7). seedlings when the roots are 1–2 cm long and the soil is Sporangia Sporangia are ovoid to ellipsoid; 18–30 lm 9 14–20 lm papillate; germinating to give 20 reniform, biflagellate zoospores (Fig. 8). Fig. 5 Sporulation of the oomycete at soil level on the stem of Fig. 6 Sporangiophores of Plasmopara halstedii with sporangia (Photo: sunflower seedling in a climatic chamber. (Photo: Aranka Kormany) Aranka Kormany). ª 2014 OEPP/EPPO, Bulletin OEPP/EPPO Bulletin 44, 350–359 Plasmopara halstedii 353 oomycete in sunflower seed samples by the Polymerase Chain Reaction (PCR). This method may be adapted for identification of in planta grown structures (sporangiophores bearing sporan- gia). A duplex real-time PCR has also been developed by Ioos et al. (2012). Details of the tests are presented in Appendi- ces 2 and 3. Reference material P. halstedii pathotypes designated as 100, 330, 700, 710 and 730 can be obtained from the Szent Istvan University culture collection on request (R.
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