Plasmopara halstedii (downy mildew of sunflower)
- Taxonomic Tree
- Notes on Taxonomy and Nomenclature
- Distribution Table
- Risk of Introduction
- Habitat List
- Hosts/Species Affected
- Growth Stages
- List of Symptoms/Signs
- Biology and Ecology
- Seedborne Aspects
- Plant Trade
- Detection and Inspection
- Prevention and Control
- Distribution Maps
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PicturesTop of page
IdentityTop of page
Preferred Scientific Name
- Plasmopara halstedii (Farl.) Berl. & De Toni
Preferred Common Name
- downy mildew of sunflower
Other Scientific Names
- Plasmopara helianthi Novot.
International Common Names
- Spanish: mildiu del girasol
- French: mildiou du tournesol
Local Common Names
- Germany: Falscher Mehltau: Sonnenblume
- Hungary: napraforgo-peronoszpora
- Italy: Peronospora del Girasole
- Russian Federation: lozhnaga muchinistaya rosa podsolnechnika
- PLASHA (Plasmopara halstedii)
Taxonomic TreeTop of page
- Domain: Eukaryota
- Kingdom: Chromista
- Phylum: Oomycota
- Class: Oomycetes
- Order: Peronosporales
- Family: Peronosporaceae
- Genus: Plasmopara
- Species: Plasmopara halstedii
Notes on Taxonomy and NomenclatureTop of page The fungus was originally described by Farlow in 1882 as Peronospora halstedii, the name referring to Halsted, who first collected it on Eupatorium purpureum. After a revision of the genus Peronospora, the fungus was renamed Plasmopara halstedii in 1888 (Berlese et De Toni), and this name has become generally accepted and conventionally used in many parts of the world (Sackston, 1981). The name refers to a closely related group of fungi, the P. halstedii complex (Leppik, 1966), attacking cultivated sunflowers, other annual and perennial species of Helianthus, as well as a number of other composites (members of the Asteraceae).
As another concept, on the basis of pathological assessments and of morphological examinations, Novotel'nova (1966) differentiated between species and forms, giving the name Plasmopara helianthi to the fungus, thought to be confined to the genus Helianthus, with further specialization on intrageneric taxa as formae speciales: f.sp. helianthi (downy mildew of sunflower), f.sp. perennis, and f.sp. patens. The species name Plasmopara helianthi is now regarded as taxonomically invalid, because its introduction by Novotel'nova did not adhere to the rules of the International Code of Botanical Nomenclature (Gulya et al. 1997). Novotel'nova's (1966) differentiation between species and forms of this fungus on the basis of minor morphological traits is not convincing when facing the great variability of biometric characters observed even among sporangiophores and sporangia of single isolates of the pathogen (Delanoe 1972). Finally, the extremely narrow host range reported from pathogenicity tests with local Russian populations of sunflower downy mildew was not found in isolates from other countries. For example, in Hungary the fungus population of P. halstedii was found to be wider in host range than was indicated by Novotel'nova for P. helianthi (Virányi, 1984; Bohár and Vajna, 1996). Consequently, Novotel'nova's concept of classification appears not to be valid for regions other than the Krasnodar area of Russia (Sackston, 1981; Virányi, 1984).
DescriptionTop of page P. halstedii is characterized by its monopodially branched, slender sporangiophores, usually ending in three sterigmata and bearing oval to elliptical sporangia, each with an apical papilla. Size of sporangia varies, as does the number of biflagellate zoospores released by each sporangium. Interestingly, sporangiophores formed on sunflower roots differ in shape from those emerging on the leaves (Novotel'nova, 1966). The vegetative thallus of the fungus is composed of intercellular, hyaline, aseptate hyphae containing granular cytoplasm. Hyphae produce globular haustoria in adjacent host cells.
Sexual reproduction is by means of oogamy; oogonia, spherical in shape, and club-shaped antheridia form on distal hyphal branches separately, those hyphae being of different mating types. Oospores are spherical, hyaline to light brown, thick-walled, temporarily surrounded by oogonial remnants (Hall, 1989).
DistributionTop of page
The downy mildew found in Australia and New Zealand on Arctotheca and Arctotis is thought to be caused by a new species, Plasmopara majewskii sp. nov. rather than by P. halstedii (Constantinescu and Thines, 2010). P. halstedii is absent from Australia and New Zealand.
See also CABI/EPPO (1998, No. 235).
Distribution TableTop of page
The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.
|Continent/Country/Region||Distribution||Last Reported||Origin||First Reported||Invasive||Reference||Notes|
|China||Restricted distribution||CABI/EPPO, 2014; EPPO, 2014|
|-Jilin||Present||Hua and Ma, 1996|
|-Liaoning||Present||Yang and Wei, 1988; CABI/EPPO, 2014; EPPO, 2014|
|-Shandong||Present||Hua and Ma, 1996|
|Georgia (Republic of)||Widespread||CABI/EPPO, 2014; EPPO, 2014|
|India||Widespread||CABI/EPPO, 2014; EPPO, 2014|
|-Andhra Pradesh||Present||Moses, 1989; Patil et al., 1993; CABI/EPPO, 2014; EPPO, 2014|
|-Indian Punjab||Present||CABI/EPPO, 2014; EPPO, 2014|
|-Karnataka||Present||Patil et al., 1993; CABI/EPPO, 2014; EPPO, 2014|
|-Madhya Pradesh||Present||Agrawal et al., 1991; CABI/EPPO, 2014|
|-Maharashtra||Present||Mayee and Patil, 1987; CABI/EPPO, 2014; EPPO, 2014|
|-Tamil Nadu||Present||CABI/EPPO, 2014|
|-Uttar Pradesh||Present||CABI/EPPO, 2014|
|Iran||Present||Rahmaini & Madjidieh-Ghassemi, 1975; CABI/EPPO, 2014; EPPO, 2014|
|Iraq||Present||CABI/EPPO, 2014; EPPO, 2014|
|Israel||Widespread||CABI/EPPO, 2014; EPPO, 2014|
|Japan||Present||CABI/EPPO, 2014; EPPO, 2014|
|Kazakhstan||Present||CABI/EPPO, 2014; EPPO, 2014|
|Korea, Republic of||Present||Choi et al., 2009; CABI/EPPO, 2014|
|Pakistan||Present||CABI/EPPO, 2014; EPPO, 2014|
|Taiwan||Absent, formerly present||CABI/EPPO, 2014; EPPO, 2014|
|Turkey||Widespread||****||Onan and Onogur, 1989; CABI/EPPO, 2014; EPPO, 2014|
|Egypt||Absent, unreliable record||EPPO, 2014|
|Ethiopia||Present||CABI/EPPO, 2014; EPPO, 2014|
|Kenya||Present||CABI/EPPO, 2014; EPPO, 2014|
|Morocco||Widespread||Achbani and Tourvieille, 1993; CABI/EPPO, 2014; EPPO, 2014|
|South Africa||Restricted distribution||Gulya et al., 1996; CABI/EPPO, 2014; EPPO, 2014|
|Uganda||Present||CABI/EPPO, 2014; EPPO, 2014|
|Zimbabwe||Present||Gulya et al., 1996; CABI/EPPO, 2014; EPPO, 2014|
|Canada||Present||CABI/EPPO, 2014; EPPO, 2014|
|-Manitoba||Present||Rashid, 1991; Rashid, 1993; CABI/EPPO, 2014; EPPO, 2014|
|-Nova Scotia||Present||CABI/EPPO, 2014; EPPO, 2014|
|-Ontario||Present||CABI/EPPO, 2014; EPPO, 2014|
|-Quebec||Present||CABI/EPPO, 2014; EPPO, 2014|
|-Saskatchewan||Present||CABI/EPPO, 2014; EPPO, 2014|
|Mexico||Present||Greathead and Greathead, 1992; CABI/EPPO, 2014|
|USA||Present||CABI/EPPO, 2014; EPPO, 2014|
|-California||Present||Gulya et al., 1991a; CABI/EPPO, 2014; EPPO, 2014|
|-Florida||Present||CABI/EPPO, 2014; EPPO, 2014|
|-Kansas||Present||Jardine & Gylya, 1994; CABI/EPPO, 2014; EPPO, 2014|
|-Maryland||Present||CABI/EPPO, 2014; EPPO, 2014; Rivera et al., 2014|
|-Minnesota||Present||Garcia and Gulya, 1991; CABI/EPPO, 2014; EPPO, 2014|
|-New Jersey||Present||CABI/EPPO, 2014|
|-New Mexico||Present||CABI/EPPO, 2014|
|-New York||Present||CABI/EPPO, 2014|
|-North Carolina||Present||CABI/EPPO, 2014|
|-North Dakota||Present||Garcia and Gulya, 1991; Gulya, 1996; CABI/EPPO, 2014; EPPO, 2014|
|-South Dakota||Present||Garcia and Gulya, 1991; Gulya, 1996; CABI/EPPO, 2014; EPPO, 2014|
|-Virginia||Present||Hong, 2006; CABI/EPPO, 2014|
|-West Virginia||Present||CABI/EPPO, 2014|
Central America and Caribbean
|Dominican Republic||Present||CABI/EPPO, 2014; EPPO, 2014|
|Argentina||Present||Gulya et al., 1991a; CABI/EPPO, 2014; EPPO, 2014|
|Brazil||Present||CABI/EPPO, 2014; EPPO, 2014|
|-Minas Gerais||Present||Duarte et al., 2013; EPPO, 2014|
|-Parana||Present||Leite et al., 2007; CABI/EPPO, 2014|
|-Rio Grande do Sul||Present||Schuck and Jobim, 1988; CABI/EPPO, 2014|
|-Sao Paulo||Present||CABI/EPPO, 2014; EPPO, 2014|
|Chile||Present||CABI/EPPO, 2014; EPPO, 2014|
|Paraguay||Present||CABI/EPPO, 2014; EPPO, 2014|
|Uruguay||Present||CABI/EPPO, 2014; EPPO, 2014|
|Albania||Present||Kola, 1980; CABI/EPPO, 2014; EPPO, 2014|
|Austria||Present, few occurrences||CABI/EPPO, 2014; EPPO, 2014|
|Belgium||Absent, no pest record||EPPO, 2014|
|Bosnia-Hercegovina||Present||Batinica et al., 1973; CABI/EPPO, 2014|
|Bulgaria||Widespread||****||CABI/EPPO, 2014; EPPO, 2014|
|Croatia||Restricted distribution||CABI/EPPO, 2014; EPPO, 2014|
|Czech Republic||Restricted distribution||CABI/EPPO, 2014; EPPO, 2014; Sedlárová et al., 2016|
|Estonia||Restricted distribution||CABI/EPPO, 2014; EPPO, 2014|
|France||Restricted distribution||Lafon et al., 1996; CABI/EPPO, 2014; EPPO, 2014|
|Germany||Present, few occurrences||1986||Spring et al., 1994; CABI/EPPO, 2014; EPPO, 2014|
|Greece||Restricted distribution||Thanassoulopoulos and Mappas, 1992; CABI/EPPO, 2014; EPPO, 2014|
|Hungary||Restricted distribution||1949||Virányi and Gulya, 1996; Bán et al., 2014; Bán et al., 2014; CABI/EPPO, 2014; EPPO, 2014|
|Italy||Present||Zazzerini, 1983; CABI/EPPO, 2014; EPPO, 2014|
|Moldova||Present||CABI/EPPO, 2014; EPPO, 2014|
|Netherlands||Present, few occurrences||NPPO of the Netherlands, 2013; CABI/EPPO, 2014; EPPO, 2014|
|Poland||Absent, formerly present||Kucmierz, 1976; CABI/EPPO, 2014; EPPO, 2014|
|Portugal||Absent, confirmed by survey||EPPO, 2014|
|Romania||Restricted distribution||CABI/EPPO, 2014; EPPO, 2014|
|Russian Federation||Widespread||Novotel'nova, 1966; CABI/EPPO, 2014; EPPO, 2014|
|-Central Russia||Restricted distribution||CABI/EPPO, 2014; EPPO, 2014|
|-Southern Russia||Restricted distribution||CABI/EPPO, 2014; EPPO, 2014|
|-Western Siberia||Present||CABI/EPPO, 2014; EPPO, 2014|
|Serbia||Present||CABI/EPPO, 2014; EPPO, 2014|
|Slovakia||Restricted distribution||CABI/EPPO, 2014; EPPO, 2014|
|Spain||Restricted distribution||Melero-Uara et al., 1982; CABI/EPPO, 2014; EPPO, 2014|
|Switzerland||Widespread||CABI/EPPO, 2014; EPPO, 2014|
|UK||Transient: actionable, under eradication||IPPC, 2010; CABI/EPPO, 2014; EPPO, 2014|
|-England and Wales||Transient: actionable, under eradication||CABI/EPPO, 2014; EPPO, 2014|
|Ukraine||Restricted distribution||CABI/EPPO, 2014; EPPO, 2014|
|Yugoslavia (Serbia and Montenegro)||Present||Masirevic, 1992|
|Australia||Absent, invalid record||CABI/EPPO, 2014; EPPO, 2014|
|-New South Wales||Absent, invalid record||EPPO, 2014|
|-South Australia||Absent, invalid record||EPPO, 2014|
|New Zealand||Absent, invalid record||Hall, 1989; CABI/EPPO, 2014; EPPO, 2014|
Risk of IntroductionTop of page RISK CRITERIA CATEGORY
ECONOMIC IMPORTANCE High
SEEDBORNE INCIDENCE Low
SEED TRANSMITTED Yes
SEED TREATMENT Yes
OVERALL RISK Moderate
Notes on Phytosanitary Risk
P. halstedii is listed as an A1 quarantine pest by IAPSC, but not by any other regional plant protection organization (EPPO, 1996). However, Australia treats it as a major quarantine pest. Possible quarantine status arises from the existence of several pathogenic forms (pathotypes), the distribution of which is limited to geographical regions. Therefore, quarantine restrictions may be needed for sunflower seed imported into areas where the pathotype(s) in question have not yet been reported. The pathogen is potentially dangerous everywhere that sunflowers are grown, except where high soil temperature (above 25°C) and/or drought is a limiting factor. If control measures are lacking and conditions are favourable, downy mildew can be devastating to sunflower production.
Habitat ListTop of page
Hosts/Species AffectedTop of page
P. halstedii has been reported to inhabit over 100 host species from a wide range of genera in the Asteraceae, including annual and perennial Helianthus spp., as well as a number of other wild or cultivated Asteraceae (Berlese et Toni, 1888; Leppik, 1966; Novotel'nova, 1966). Unfortunately, few experimental data of infection studies are available on species-specificity of the isolated pathogen populations. Under laboratory conditions, at least six wild species of Helianthus (H. annuus, H. argophyllus, H. debilis, H. divaricatus, H. grosseseratus and H. petiolaris) and the composite Artemisia vulgaris have successfully been infected and the fungus sporulated on cotyledon leaves (Virányi, 1984). In contrast, Zimmer (1974) reported that as with other perennial Helianthus species, H. grosseseratus was immune to artificial inoculations by P. halstedii.
The only host crop known so far is the cultivated sunflower (H. annuus). Another cultivated member of the Asteraceae, Senecio sp., a pot-cultivated flower plant, has been reported from Italy as a host of P. halstedii (Gullino and Garibaldi, 1988), but cross infection studies with H. annuus were not conducted. As for a significant wild host, Xanthium strumarium (Virányi, 1984; Komajati et al., 2007) and more recently Ambrosia artemisifolia (Walcz et al., 2000) were found to contribute as a potential reservoir of this fungus in Hungary. Both isolates readily attacked sunflower, and the isolate from A. artemisifolia was characterized on sunflower differential lines as virulence type 730 (equally to former race 4), a common pathotype in Hungary (Walcz et al., 2000).
Growth StagesTop of page Flowering stage, Pre-emergence, Seedling stage, Vegetative growing stage
SymptomsTop of page The symptoms induced by P. halstedii in sunflower are diverse, they depend on the age of tissue, level of inoculum, cultivar reaction and environment (moisture and temperature). There are three types of infection: systemic, local and latent (Spring, 2001).
Systemically infected plants are stunted to a lesser or greater extent and the leaves of affected plants show pale green or chlorotic mottling which spreads along the main veins and over the lamella. The biomass production of vegetative and generative plant parts is reduced drastically (Spring et al., 1991). Young leaves of severely affected plants often become chlorotic as a whole, curl downward and are rigid and thick. Under moist conditions, a white downy growth composed of sporangiophores and sporangia of the fungus develops on the underside of leaves, their extent strictly corresponding to the chlorotic areas on the upper leaf surface. Due to shortening of internodes, a mildewed sunflower often has a cabbage-like appearance.
The heads of infected sunflower plants are reduced in size and face upward (horizontal head), bearing no or a limited number of seeds of poor viability. The root system of mildewed sunflowers is underdeveloped, with a significant reduction in secondary root formation and with a dark brown appearance on their surface. Other, less familiar, symptoms associated with systemic infection include damping-off of seedlings, pith discoloration of stems and/or capitulum, disturbance of inflorescence, twisted leaves and basal gall (Sackston, 1981). Furthermore, systemic downy mildew infection may be localized in root and lower stem tissues (cotyledon- or hypocotyl-limited infection) with some pathogen-host combinations, under certain conditions (Ljubich and Gulya, 1988; Virányi and Gulya, 1996).
Local infection of leaves occur frequently, but often do not attract much attention. As a result, small, angular, pale green spots, delimited by the veins, appear on the leaves. Under conditions of high relative humidity, downy growth (sporulation) develops on the lower leaf surface. At a later stage, the host tissue dies leaving brownish leaf lesions. For a long time it was widely ignored that these local infections may give rise to systemic infection, the fungus growing through the petioles into the stem (Sackston, 1981). In the past years in Hungary, a local-to-systemic infection extensively occurred in some sunflower fields due to favourable weather conditions coupled with unusually high infection pressure (Vida, 1996). Similar observations were made in Germany in 1999 where in up to 8% of the locally infected plants the pathogen succeeded to invade the petiole and propagated systemically into the upper plant parts (Spring 2001).
The latent type of infection appears symptomless and affected plants can not easily be recognized from outside. It may derive either from below ground infections on plants which are able to control the pathogen by prohibiting its expansion from the roots and the hypocotyl into the epicotyl or it may derive from very late infections during flowering stage, when the growth of vegetative parts has finished and therefore no symptoms become visible. The former type is typical for some so-called resistant sunflower genotypes (which sometimes allow sporulation on hypocotyls and cotyledons, known as HLI/CLI-type infection; Tourvieille de Labrouhe et al., 2000). The latter mostly leads to seed contamination and, due to the interaction with the plant's phytohormone system, may be recognized by retarded decomposition of the chlorophyll and inhibited gravitropic reaction of the aging flower head (Spring 2001). It should be noted that in both cases the pathogen stays alive and is able to complete its life cycle through sexual reproduction by the formation of oospores (Heller et al. 1997).
List of Symptoms/SignsTop of page
|Leaves / abnormal colours|
|Leaves / fungal growth|
|Roots / fungal growth on surface|
|Roots / necrotic streaks or lesions|
|Seeds / discolorations|
|Seeds / empty grains|
|Stems / stunting or rosetting|
|Whole plant / dwarfing|
|Whole plant / early senescence|
|Whole plant / plant dead; dieback|
Biology and EcologyTop of page
Starting from a single oospore that germinates and gives rise to a zoosporangium, zoospore differentiation and release are the subsequent steps of development. In the presence of free water, the zoospores remain in motion for hours but tend to move to infection sites (root, hypocotyl) soon if available. Following encystment and germ tube elongation (the latter usually terminates in an appressorium against a host cell) the fungus develops an infection structure (infection peg) for direct penetration. Under experimental conditions it was demonstrated that germ tubes do not usually form appressoria in water but they do so in the presence of host cells (Gray et al., 1985). After penetration, the fungus grows intracellularly and then intercellularly, and once being established in a susceptible host (compatible) it starts to colonize the entire plant systemically by growing preferably toward the shoot apex and, to a lesser extent, in the direction of the root. When conditions are favourable, asexual sporulation takes place on affected leaves and occasionally on below-ground tissues. Fully developed sporangia disseminate by wind and, since they are short-lived and sensitive to drought and direct sunshine, their survival depends on the current weather situation. Oospores are also produced in infected plant parts, primarily in root and lower stem tissues, whereas leaves and upper plant parts, except seeds, are free from these resting spores (Sackston, 1981; Virányi, 1988; Onan and Onogur, 1991). The most susceptible stage of host development is between germination and emergence (Meliala et al. 2000).
Survival and Source of Inoculum.
With respect to the primary infection, P. halstedii is a soilborne pathogen. Its oospores serve as primary inoculum to underground tissues of young sunflower seedlings. It may also be windborne, causing secondary infection of leaves and/or inflorescence. If the latter is the case, the fungus might also be seedborne: the affected seeds carrying mycelium and/or oospores internally. Oospores develop mainly in root and lower stem tissues of mildewed plants, with or without visible symptoms and, with plant residues of the preceding sunflower crop, they come into the soil. Oospores are long-lived and are able to survive for at least 6-8 years (Sackston, 1981; Virányi, 1988). It is generally thought that oospores mainly germinate under wet conditions. However, only a few results on the germination dynamics have been available so far. A low-temperature shock prior to wetness and the presence of host exudates released by roots were shown to enhance the germination process (Delanoe, 1972). In another report (Spring & Zipper 2000), no such temperature effects could be observed and freshly developed oospores were reported to germinate spontaneously in water within a period of 10-30 days, but at a highly variable rate (1-17%).
Novotel'nova (1966) stated that the fungus was present in a significant percentage of seeds from naturally infected plants. Later on, seed infection by artificial inoculation was confirmed (Sackston, 1981). Such seeds gave rise to apparently healthy seedlings with no typical symptoms (latent type of infection) but the pathogen sporulated more often on the roots of these symptomless plants from infected seed.These observations were confirmed by field studies on late infected plants that occur frequently under favorable weather conditions during the flowering stage. The crucial role of such symptomless and nonsystemic infections for the distribution of the pathogen was discussed by Spring (2001).
The significance of windborne sporangia in disease initiation has long been regarded to be low. However, secondary infection is considered as an important factor in the spreading of the disease in certain regions under favourable environmental conditions. As an example, secondary infection by zoosporangia was numerous in some Hungarian fields in 1995 and 1996 due to unusual weather and a high infection pressure (Vida, 1996). Apart from the fact that secondary infection of inflorescence may give rise to latent infection of seeds by P. halstedii (Sackston, 1981), from local leaf lesions the fungus is able to proceed and grow into the stem causing systemic infection (Spring 2001).
The nature of the inoculum (oospore or zoospore), weather variables (relative humidity, temperature), infection site (age of tissue), as well as cultivar reaction are factors that influence or determine the infection process, disease incidence, and severity. Zoospores, originating from either sexual or asexual sporulation, require free water for retaining viability and capability of moving toward infection sites. Consequently, rainfall or intensive irrigation will be a prerequisite for the initiation of infection. It was shown by several studies that if there was enough rain or corresponding water supply during the first two weeks after sowing, the incidence of primary infection from the soil increased. However, the duration of time that favours infection is relatively short and even susceptible sunflowers become resistant with age (Sackston, 1981). Tourvieille et al. (2008a) found that the risk of downy mildew attack appeared greatest if there was heavy rainfall when sunflower seedlings were at their most susceptible stage, whereas heavy rainfall before sowing or after emergence had no effect on the percentage of diseased plants. Göre (2009) that low temperature and extensive spring rains in approximately 85% yield loss and lower quality of sunflower production in the Marmara region of Trace. Besides environmental conditions, disease intensity may also be influenced by the aggressiveness of the pathogen population. Sakr et al. (2009) were able to differentiate the two pathogen strains in terms of their aggressiveness based on the population’s latent period and sporulation density.
The existence of pathogenic forms within P. halstedii became evident soon after the release of the first mildew-resistant sunflower cultivars. Initially, two pathotypes (races) of the fungus were differentiated: pathotype 1, originally referred to as the European race, and pathotype 2, known as the Red River race (referring to the Red River Valley of North America). Although additional new pathotypes have been described from the North American continent, it was in 1988 that pathotypes other than race 1 were found in Europe. By now, more than 20 different pathotypes of P. halstedii have been recorded due to different pathogenicity against sunflower genotypes and are designated by numbering in a sequence of order of appearance (Gulya et al., 1991a; Molinero-Ruiz et al., 2002). Accordingly, pathotypes 1 to 10 occur in both North America and Europe, pathotypes 2, 3, and 7 in South America, pathotype 1 in India, and pathotypes 1, 4, 8 and 9 in Africa (Henning and Franca Neto, 1985; Gulya et al., 1991a; Masirevic, 1992; Rashid, 1993; Spring et al., 1994; Virányi and Gulya, 1995a, 1995b; Gulya et al., 1996; Melero-Vara et al., 1996; Lafon et al., 1996; Molinero-Ruiz et al., 2002). With the increasing number of new pathotypes, and with the aim at providing the investigators with a more accurate and comparable designation, Gulya et al. (1998) suggested the use of a coded triplet notation that is based on the virulence patterns of P. halstedii isolates determined on a 3 x 3 set of differentials. This new system of classification is likely to enhance communication between workers interested in, and responsible for, identifying and monitoring pathotypes of P. halstedii worldwide. At present, in France, there is another identification system still in use and, as a result, pathotypes designated with capital letters have been published, making international comparison difficult (Lafon et al., 1996).
Recently, Gulya (2007) listed as many as 35 pathotypes (virulence phenotypes). He found a considerable variation by continent on the number of pathotypes; Asia and South America having the fewest number of pathotypes identified followed by Africa, Europe and North America. The countries with the highest numbers of pathotypes are Canada, France and the USA. There could be various explanations for the appearance of new virulent forms (Delmotte et al. 2008), but the increase in pathotypes during the last decade is probably due to the introduction of new cultivars with different genetic pedigrees and to the intensity of production, particularly in Europe.
A further difficulty is caused by the presence of various pathotypes in the same field. Testing such field isolates with sunflower differential lines makes it difficult to detect the proportion of the less virulent genotypes in a mixture. For that reason, single spore or single sporangium strains should be tested and methods to create them were recently developed (Spring et al. 1998). As an alternative to characterize genotypes of P. halstedii, various molecular approaches were recently attempted (Borovkova et al., 1992; Borovkov et al., 1993; Roeckel-Drevet et al., 1997; Intelmann and Spring, 2002). However, none of these techniques has so far been able to show a correlation of DNA markers with the virulence behavior of the tested strain, thus prohibiting its use for pathotype studies. Based on differences in partial sequence analysis of the nuclear ITS region, Spring et al. (2006) succeeded in differentiating between certain pathotypes of sunflower downy mildew originating from different geographic regions. Molecular genetic studies based on single nucleotide polymorphisms (SNPs) and size variation in expressed sequence tags (ESTs) were performed using sunflower downy mildew isolates from France and Russia (Giresse et al. 2007).
Seedborne AspectsTop of page
P. halstedii has been found to occur in sunflower seeds from naturally infected plants, either as mycelium or oospores (Novotel'nova, 1966). Doken (1989) reported that the mycelium was only found in the testa and in the inner layer of the pericarp; it was absent from the embryo. Following artificial inoculation, Cohen and Sackston (1973) confirmed that sunflower buds inoculated with P. halstedii became systemically infected and produced infected seeds. Oospores were observed in seeds of inoculated and naturally infected plants in the field. Other records of seed infection are known from Iran (Zad, 1978), Turkey (Döken, 1989) and Germany (Spring, 2001). The fungus usually invades the ovary and the pericarp, but fails to grow into the embryo (Novotel'nova, 1966; Döken, 1989). Seed infection regularly occurs in systemically infected plants if they survive up to the flowering stage. In such cases the development of the embryo is often retarded or inhibited. Moreover, such plants are dwarf and will seldom be harvested. They may increase the local stock of oospores in a field, but for the seed-derived long distance dispersal of the pathogen they appear to be less important than seeds from late infected symptomless plants (Spring, 2001). The latter type of infection is very dependent on the weather conditions during the flowering process. Thus in dry years the number of pathogen-contaminated seeds is very low and may not exceed several in one thousand, but may be much higher after a cool and humid period in June/July. For example, Spring (2001) found that close to 10% of seeds from a field in Germany were contaminated and Döken (1989), under favourable experimental conditions, observed fungal structures in 28% of the seeds examined.
Effect on Seed Quality
Sunflower seeds produced in downy mildewed plants are either under-developed, colourless or, rarely, they look healthy. Even in the latter case, such infected seeds are of poor quality; they produce abnormal seedlings and germination rate is low (Döken, 1989).
Cohen and Sackston (1974) showed that infected seeds gave rise to symptomless (latent infection) plants. Infected seeds were also effective when used as inoculum, causing 14-89% infection in plants kept at 20°C. Most of the resulting infections were latent. Systemic infection occurred, but latent infection was more frequent in plants grown in soil containing infected plant debris. Doken (1989) also showed that seeds from infected plants rarely gave rise to plants exhibiting systemic symptoms of P. halstedii. Nevertheless, seed transmission is particularly important since sunflower plants growing from such seeds may or may not produce disease symptoms. The latent (symptomless) form of the disease quite often occurs so that one or two generations are grown before infection becomes evident (Sackston, 1981).
P. halstedii is a soil-borne pathogen. Its oospores serve as primary inoculum to underground tissues of young sunflower seedlings. It may also be windborne, causing secondary infection of leaves and/or inflorescence. Oospores develop mainly in root and lower stem tissues of mildewed plants, with or without visible symptoms and, with plant residues of the preceding sunflower crop, they come into the soil. Oospores are long-lived and are able to survive for at least 6-8 years (Sackston, 1981; Virányi, 1988).
Seed treatment with metalaxyl gave complete control of downy mildew caused by P. halstedii in trials in 1980. Plants were protected throughout the growing period (Nikolov, 1981). More recently, a study was carried out to analyse, under growth-chamber conditions and in the field, the effectiveness of 11 fungicides applied as seed treatments. Only combinations including metalaxyl or oxadixyl gave good control of P. halstedii. These fungicides gave total protection, without phytotoxicity, at 2.5 g commercial product/kg seed for 10% metalaxyl + 48% mancozeb, and at 3 g/kg seed for the combinations 10% oxadixyl + 56% propineb and 8% oxadixyl + 56% mancozeb + 3.2% cymoxanil (Achbani et al., 1999).
From the 1990s, tolerance to metalaxyl was observed in French populations of P. halstedii (Delos et al.,1997). Meanwhile field studies revealed rates of over 70% tolerant isolates in France (Albourrie et al., 1998a, b), Spain (Ruiz et al., 2000) and the USA (Gulya, 2000). On the other hand, no tolerant isolates were yet found in similar studies conducted in Hungary (Viranyi and Walcz, 2000) and Germany (Rozynek and Spring, 2000).
Treatment of sunflower seeds with 1×108cfu/ml of PGPR strain INR7 resulted in decreased disease severity and offered 51 and 54% protection under green house and field conditions, respectively (Nandeesh Kumar et al., 2008).
The effect of strobilurins was tested as seed treatment, foliar application, and seed treatment followed by foliar application. Under greenhouse conditions neither seed treatment nor foliar application of strobilurins were phytotoxic. Seed treatment with foliar application enhanced the protection of the plants as compared to only the treatment of seeds. Foliar spray treatments alone provided an intermediate control of the disease. Trifloxystrobin showed a better effect than kresoxim-methyl and azoxystrobin (Sudisha et al., 2010).
Treatment of sunflower seeds with 5% chitosan resulted in decreased disease severity and offered 46 and 52% protection under greenhouse and field conditions respectively (Nandeesh Kumar et al., 2008).
Seed Health Tests
Blotter/grow-out (Gulya, 1995b)
1. Seeds are surface-sterilized, rinsed, and put into layers of wet filter paper to make the seeds germinate.
2. After a few days of incubation when roots are formed, the fungus if present, will sporulate on their surface under humid conditions.
3. This sporulation may also occur with otherwise symptomless, latent infected plants
DNA (Says et al., 2001)
A molecular test was developed to determine the presence of sunflower downy mildew, caused by P. halstedii, in sunflower seed samples. Several extraction methods were compared to improve the quality and yield of the scanty DNA of the fungus. Both whole and hulled seeds of contaminated samples were analysed by PCR, using P. halstedii-specific primers. The study demonstrated that the DNA of the pathogen is always detected, notably in shell fraction. These data are important for the development of a diagnostic kit.
Notes on methods
It is difficult to detect P. halstedii from sunflower seeds. Unfortunately, there is no testing method available yet that allows economical detection of the infection rate of a seedlot. However, experimental results obtained by ELISA suggest that this might be a useful method for the future (Liese et al., 1982). Subsequent attempts were made by French scientists to use immunological tests (Bouterige et al., 2000) and PCR techniques (Roeckel-Drevet et al., 1999) for the detection of P. halstedii in sunflower, but the commercial usefulness of the tests has not yet been proved. Spring and Haas (2002) showed that fatty acid analysis might become a diagnostic feature for the contamination of sunflower seeds with downy mildew. In all such cases, however, it will be a difficult matter of tracing minute amounts of pathogenic characters against a huge background of healthy host tissue.
Spring and Haas (2004) were successful with fatty acid patterns as markers of infected vs. healthy sunflower seeds. Ioos et al. (2007) developed a method for the direct detection of the pathogen in seed samples based on the ribosomal large sub unit DNA. Though their test was able to detect a near 1:50 ratio of seed contamination, this was not practical, probably because of the problem of representative sampling. Furthermore, from the practical point of view it might be of interest for sunflower growers to know in advance, i.e. before sowing what pathotype (s) exist(s) and to what extent in a particular field. For this reason, Gulya (2004) and Tourvieille et al. (2008b) developed a bioassay using soil samples to assess downy mildew risk at the field level. Under different geographical and climatic conditions, their results revealed a good correlation between the soil infestation measured by the soil bioassay and the presence of infected plants in previous years.
Plant TradeTop of page
|Plant parts liable to carry the pest in trade/transport||Pest stages||Borne internally||Borne externally||Visibility of pest or symptoms|
|Flowers/Inflorescences/Cones/Calyx||hyphae; spores||Yes||Yes||Pest or symptoms usually visible to the naked eye|
|Fruits (inc. pods)||hyphae; spores||Yes||Yes||Pest or symptoms usually visible to the naked eye|
|Growing medium accompanying plants||spores||Yes||Pest or symptoms usually invisible|
|Leaves||hyphae; spores||Yes||Yes||Pest or symptoms usually visible to the naked eye|
|Roots||hyphae; spores||Yes||Pest or symptoms usually visible to the naked eye|
|Stems (above ground)/Shoots/Trunks/Branches||hyphae; spores||Yes||Yes||Pest or symptoms usually visible to the naked eye|
|True seeds (inc. grain)||hyphae; spores||Yes||Pest or symptoms usually invisible|
|Plant parts not known to carry the pest in trade/transport|
ImpactTop of page As the majority, if not all, of systemically infected plants either die prematurely or hardly produce viable seed, they make no contribution to yield. Furthermore, reduction in seed yield may also be due to pre- or post-emergence damping-off of severely mildewed seedlings under very favourable conditions, a symptom often overlooked and/or underestimated. Disease severity may vary considerably according to region, year, growing conditions and cultivar (pathotype combination). The incidence of downy mildewed sunflowers in a field may range from traces to nearly 50% or even up to 95% (Sackston, 1981). In Europe, after its first appearance in 1941, the disease increased rapidly and by 1977 it was rated a major disease in all sunflower-producing countries (Sackston, 1981). Similarly, a dramatic increase of disease incidence occurred in North America where the pathogen spread rapidly in the Northern Great Plains (USA and Canada) (Sackston, 1981) and even now it contributes considerably to yield losses (Gulya, 1996). As for other continents, the disease is spreading in South America, in Africa and may also be of concern for Australia.
DiagnosisTop of page
Due to host specialization, the downy growth on affected plants should be produced by P. halstedii, so it is not absolutely necessary to make additional microscopical observations. However, to identify the fungus from other non-crop hosts will need a thorough microscopical examination of samples and a subsequent inoculation test onto sunflower. As an obligately biotrophic fungus, P. halstedii cannot be cultured on artificial media but in vitro techniques by using cell suspension, callus or sunflower tissue cultures might be possible, at least in the short term (Gray and Sackston, 1983; Virányi and Sziráki, 1986). Efforts are being made to find molecular techniques sufficient to detect even a very small quantity of fungal biomass in the host (Tourvieille et al., 1996). A diagnostic protocol for Plasmopara halstedii is described in EPPO (2008).
Detection and InspectionTop of page Downy mildew of sunflower is easy to identify in the field by monitoring the crop for typical visible symptoms. It is primarily soilborne but can also be seedborne. Although seed infection is usually rare (less than one per thousand in seeds from systemically infected sunflower plants), such seeds are thought to be responsible for long distance disease spread (Sackston, 1981). It may therefore be required to know whether a seed lot is infected or not and, if so, to what extent. However, seedborne inoculum is difficult to detect even with time-consuming laboratory procedures. ELISA was reported as a successful tool for detecting the fungus in infected sunflower tissues (Liese et al., 1982) and has been suggested for use in seed inspection. However, no laboratory has introduced this method for the detection of P. halstedii from sunflower seed or from other plant parts.
Prevention and ControlTop of page
Due to the variable regulations around (de)registration of pesticides, your national list of registered pesticides or relevant authority should be consulted to determine which products are legally allowed for use in your country when considering chemical control. Pesticides should always be used in a lawful manner, consistent with the product's label.
P. halstedii is extremely difficult (or impossible) to eradicate once it is established in an area. Sunflower hybrids resistant to downy mildew are available, but new pathogenic forms (pathotypes) of the fungus are being formed in nature, making the use of formerly resistant cultivars in a particular area questionable (Gulya et al., 1991a). Sunflower breeders are continuously searching for, and building in, new resistance sources from wild Helianthus species and, as a result, a resistance gene-pool sufficient against the majority of currently known virulence factors of P. halstedii populations seems to be available for producing new resistant cultivars (Sackston, 1992; Skoric, 1994).
Fungicides with definite systemic and long-lasting properties (e.g. metalaxyl or related compounds) are of significance in controlling the disease (Oros and Virányi, 1987). Even with the use of resistant cultivars, fungicide seed dressings are also used to prevent underground infection of seedlings. Recently, reduced sensitivity to metalaxyl has been reported from Turkey (Delen et al., 1985), France (Lafon et al., 1996), Spain (Riuz et al., 2000) and the USA (Gulya 2000). If P. halstedii strains tolerant of metalaxyl and other phenylamides did arise in local field populations, those chemicals currently used should be replaced by others. An easy to handle and fast test for the screening of field isolates for metalaxyl/fungicide tolerance was recently developed on the base of leaf disk infections (Rozynek and Spring 2001). As future alternatives to fungicide treatments, research efforts are being made to find out new ways of control, such as chemically induced resistance or the use of biological antagonists (Sackston et al., 1992). BABA-induced resistance against P. halstedii in sunflower is mediated through enhanced expression of genes for defense related proteins (Nandeesh Kumar et al., 2009).
ReferencesTop of page
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Distribution MapsTop of page
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