Plasmopara viticola (grapevine downy mildew)
- Taxonomic Tree
- Notes on Taxonomy and Nomenclature
- Distribution Table
- Hosts/Species Affected
- Host Plants and Other Plants Affected
- Growth Stages
- List of Symptoms/Signs
- Biology and Ecology
- Means of Movement and Dispersal
- Plant Trade
- Detection and Inspection
- Similarities to Other Species/Conditions
- Prevention and Control
- Distribution Maps
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PicturesTop of page
IdentityTop of page
Preferred Scientific Name
- Plasmopara viticola (Berk. & M.A. Curtis) Berl. & de Toni
Preferred Common Name
- grapevine downy mildew
Other Scientific Names
- Botrytis viticola Berk. & M.A. Curtis
- Peronospora viticola (Berk. & M.A. Curtis) de Bary
- Plasmopara amurensis Prots.
- Rhysotheca viticola (Berkeley & Curtis) G.W. Wilson
International Common Names
- English: brown rot; downy mildew of grapevine; grey rot
- Spanish: mildiu de la vid; mildiu velloso de la vid
- French: mildiou de la grappe; mildiou de la vigne; rot-brun de la vigne; rot-gris de la vigne
Local Common Names
- Germany: Falscher Mehltau: Weinrebe; Lederbeerenkrankheit: Weinrebe; Peronospora: Weinrebe
- Italy: Peronospora della vite
- PLASVI (Plasmopara viticola)
Taxonomic TreeTop of page
- Domain: Eukaryota
- Kingdom: Chromista
- Phylum: Oomycota
- Class: Oomycetes
- Order: Peronosporales
- Family: Peronosporaceae
- Genus: Plasmopara
- Species: Plasmopara viticola
Notes on Taxonomy and NomenclatureTop of page As reviewed by Viala (1893) and Gregory (1915), P. viticola was first collected and studied in 1834 by Schweinitz, who named it Botrytis caca (Schweinitz, 1837). Berkeley and Curtis (1848) later described the organism as Botrytis viticola. De Bary (1863) transferred the pathogen to a new genus and described it as Peronospora viticola. Berlese and de Toni (1888) redescribed the pathogen as Plasmopara viticola after Schröter (1886) separated Peronospora into two genera, Peronospora and Plasmopara.
According to Grunzel (1960) and Rafaila et al. (1968), several authors (Procenko, 1946; Savulescu and Savulescu, 1952; Golovina, 1955) have suggested distinguishing different varieties and special forms of P. viticola based on morphology or host range. Grunzel (1960) and Rafaila et al. (1968) concluded there was no evidence for such divisions.
DescriptionTop of page After Hall (1989).
Mycelium composed of intercellular, colourless, aseptate hyphae 7-12 µm diameter, often irregularly shaped and swollen, bearing small, rounded vesicular haustoria, 4-10 µm diameter, formed predominantly in leaf tissues.
Sporangiophores hypophyllous, arborescent, 130-250(-700) x 11-14 µm, branching monopodially in the upper third at right angles to the main axis, and with a base tapering to a conical point; branches in a whorl of 4-5, 35-45 µm long, often with two opposite secondary branches 15-20 µm long, all having 3-4 conical tips 10 µm long 6 µm wide at base, diverging at right angles and tapering to a terminal swelling.
Sporangia ovoid, colourless, (17-)20(-25) x (10-)14(-16) µm, sometimes with a short pedicel, each producing 1-6 zoospores.
Zoospores reniform, laterally biflagellate, 6-8 x 4-5 µm, emerging from opposite the insertion point of the sporangium, via a papilla, or by direct penetration of the sporangium wall.
Oospores formed in leaf tissues, spherical, 28-40 µm diameter, containing 14-16 chromosomes, covered by two inner oospore membranes and an outer wrinkled oospore wall, germinating via a tube, 2-3 µm diameter, to give a pyriform sporangium, ca 25 x 35-40 µm, producing 8-20 zoospores.
DistributionTop of page P. viticola is native to the north-eastern USA. From the USA, P. viticola spread to Europe and worldwide. Today, P. viticola occurs in nearly all grape-growing regions worldwide. A few exceptionally dry grape-growing climates exist which support only minimal levels of the pathogen, such as parts of Argentina, Chile, California and Egypt (Weltzien, 1981; Lafon and Clerjeau, 1988; Emmett et al., 1992).
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|
|-Liaoning||Present||Zhang et al., 2004|
|-Ningxia||Present||Sha et al., 2007|
|-Xinjiang||Present||Chen et al., 2007|
|-Jammu and Kashmir||Present||Shahzad et al., 2006|
|-Maharashtra||Present||Khilare et al., 2003; Sawant et al., 2016|
|-Tamil Nadu||Present||Prakash et al., 2007|
|Korea, Republic of||Present||CMI, 1988; Choi et al., 2017|
|Saudi Arabia||Present||CMI, 1988|
|Sri Lanka||Present||CMI, 1988|
|Congo Democratic Republic||Present||CMI, 1988|
|Sierra Leone||Present||CMI, 1988|
|South Africa||Present||CMI, 1988|
|-British Columbia||Present||CMI, 1988|
|-Nova Scotia||Present||CMI, 1988|
|-Maryland||Present||Baudoin et al., 2008|
|-Michigan||Present||Schilder et al., 2002|
|-North Carolina||Present||Baudoin et al., 2008|
|-Pennsylvania||Present||Baudoin et al., 2008|
|-Virginia||Present||Baudoin et al., 2008|
Central America and Caribbean
|Costa Rica||Present||CMI, 1988|
|Dominican Republic||Present||CMI, 1988|
|El Salvador||Present||CMI, 1988|
|Puerto Rico||Present||CMI, 1988|
|-Minas Gerais||Present||Pereira et al., 2010|
|-Rio Grande do Sul||Present||Chavarria et al., 2007|
|-Santa Catarina||Present||Peruch and Bruna, 2008|
|-Sao Paulo||Present||Ferrari et al., 2003|
|Czech Republic||Present||Hrubý, 2003|
|Czechoslovakia (former)||Present||CMI, 1988|
|Former USSR||Present||CMI, 1988|
|Moldova||Present||Voinyak et al., 2009|
|Montenegro||Present||Latinovic and Maras, 2005|
|Russian Federation||Present||Present based on regional distribution.|
|-Russian Far East||Present||CMI, 1988|
|Slovenia||Present||Luskar et al., 2005|
|Ukraine||Present||Kilimnik and Samoilov, 2000|
|Yugoslavia (Serbia and Montenegro)||Present||CMI, 1988|
|-New South Wales||Present||CMI, 1988|
|-South Australia||Present||CMI, 1988|
|-Western Australia||Present||McKirdy et al., 1999|
|New Caledonia||Present||CMI, 1988|
|New Zealand||Present||CMI, 1988|
|Papua New Guinea||Present||CMI, 1988|
Hosts/Species AffectedTop of page P. viticola is most important as a pathogen of Vitis vinifera, V. labrusca and V. vinifera hybrids. P. viticola also attacks a number of Vitis species and related genera in the family Vitaceae, but wild hosts are not significant to disease development on the cultivated varieties (Barrett, 1939; Renfro and Bhat, 1981).
Wild Vitis host species include V. aestivalis,V. arizonica, V. berlandieri [V. cinerea var. helleri], V. californica, V. coignetiae, V. cordifolia, V. girdiana, V. monticola, V. pagnucci [V. piasezkii var. pagnuccii], V. riparia, V. romaneti, V. rotundifolia, V. rupestris and V. treleasei (Lafon and Bulit, 1981; Hall, 1989). Additionally, under laboratory conditions, Staudt and Kassemeyer (1995) observed downy mildew on V. acerifolia, V. amurensis, V. candicans [V. mustangensis], V. champinii, V. cinerea, V. doaniana, V. lanata, V. palmata, V. piasezkii, V. rubra, V. shuttleworthii, V. solonis, V. tiliifolia and V. vulpina.
Other wild host genera in the Vitaceae include Ampelopsis, Cissus, Cordifolia, Cinerea, Parthenocissus and Solonis (Renfro and Bhat, 1981; Hall, 1989).
Host Plants and Other Plants AffectedTop of page
Growth StagesTop of page Flowering stage, Fruiting stage, Seedling stage, Vegetative growing stage
SymptomsTop of page P. viticola infects all green parts of the host plant that bear stomata. It generally causes yellow discoloration, necrosis and distortion.
On young leaves, lesions appear as yellow, translucent 'oilspots' with a chocolate-brown halo (see Pictures). On cultivar Ruby Cabernet, oilspots are reddish instead of yellow (Nicholas et al., 1994). Multiple oilspots can coalesce to cover much of the leaf surface. Oilspots become dry and necrotic as they age, first in the centre and later throughout the entire lesion. On older leaves, the lesions are restricted by veins to form small, angular, yellow to reddish-brown spots which combine to form a patchwork or mosaic-like pattern.
Sporulation only occurs on the lower leaf surface, where the stomata reside. The sporangiophores and sporangia appear as a white, downy, cottony growth. Under highly favourable conditions, sporulation may appear on the undersides of leaves before the yellow oilspot becomes visible on the upper leaf surface. On older oilspots, sporulation occurs primarily on the margins of the lesion.
Infected shoot tips and rachises of young inflorescences distort into a curl or corkscrew.
Infected inflorescences and young berries appear yellow or grey and may be covered with cottony spores under favourable conditions. Sporulation occurs on pedicels and berries. Clusters infected at an early stage can result in individual berries, sections of the cluster, or even entire clusters turning brown, drying and falling off the vine.
Berries infected later in the season (after 2-3 weeks post-bloom) become discoloured and shrivel but do not support sporulation. This stage is sometimes referred to as the 'brown rot' phase. Berry stems continue to sporulate after sporulation ceases on the berries.
List of Symptoms/SignsTop of page
|Fruit / discoloration|
|Fruit / extensive mould|
|Fruit / mummification|
|Fruit / premature drop|
|Growing point / discoloration|
|Growing point / distortion|
|Growing point / mycelium present|
|Inflorescence / blight; necrosis|
|Inflorescence / discoloration (non-graminaceous plants)|
|Inflorescence / distortion (non-graminaceous plants)|
|Inflorescence / lesions; flecking; streaks (not Poaceae)|
|Leaves / abnormal colours|
|Leaves / abnormal forms|
|Leaves / abnormal leaf fall|
|Leaves / fungal growth|
|Leaves / necrotic areas|
|Stems / distortion|
|Stems / mould growth on lesion|
|Stems / mycelium present|
Biology and EcologyTop of page P. viticola is an obligately biotrophic plant pathogen with a sexual overwintering phase and asexual multiplication cycles during the growing season.
The pathogen usually overwinters as oospores in fallen leaves infected in the previous season. P. viticola is heterothallic with two mating types (Wong et al., 2001). An antheridium fertilizes an oogonium to form the sexual oospore (Kortekamp et al., 1998). In mild climates such as in California the pathogen has been observed overwintering as mycelium in buds and canes of wild grape species (Barrett, 1939).
Oospores germinate in the spring when temperatures reach about 10°C and vineyard soils are wet. Oospores germinate and form a germ tube which terminates in a macrosporangium. Germination and formation of the macrosporangia can occur in as little as 24 hours of soil wetness (Gregory, 1915; Ronzon-Tran Manh Sung and Clerjeau, 1988). When wet, the macrosporangia release an average of 8-20 (Hall, 1989) and up to 60 (Ronzon-Tran Manh Sung and Clerjeau, 1988) zoospores. At 20°C zoospore release occurs after 30 or more minutes (Ronzon-Tran Manh Sung and Clerjeau, 1988). Zoospores are dispersed onto host tissue by rainsplash.
Penetration and colonization of the host
Zoospores require surface wetness to infect the host. Infection takes place only through the stomata (Gregory, 1914). Zoospores swim on the tissue surface, encyst near stomata in groups of 2 to 5, and each spore forms a single germ tube which penetrates the stomata. In the substomatal cavity the germ tube swells, forming a substomatal vesicle from which arises a single hypha. Hyphae grow intercellularly, filling the space between the host mesophyll cells and taking on an irregular shape. In as little as 3.5 hours the first haustorium forms where the pathogen contacts the host cells. Additional haustoria form as the pathogen ramifies through host tissue, parasitizing the mesophyll cells (Langcake and Lovell, 1980; Kortekamp et al., 1998). The pathogen can initiate sporulation upon reaching a non-parasitized substomatal cavity (Langcake and Lovell, 1980; Kortekamp et al., 1998). Vascular tissue restricts pathogen growth to interveinal areas in older leaves (Langcake and Lovell, 1980).
The incubation time, the period between infection and the first appearance of symptoms, depends on temperature and ranges from 4 to 21 days, with an average of 7-10 days. The incubation period is shortest on young leaves and at temperatures of 19-24°C (Muller and Sleumer, 1934; Zachos, 1959; Rafaila et al., 1968).
The pathogen sporulates through stomata during warm, humid nights. To sporulate, P. viticola requires at least 95-98% RH, temperatures between 10 and 30°C (peak production occurs at 20°C) and at least 4 hours of darkness (Blaeser and Weltzien, 1978; Lalancette et al., 1988b; Hill, 1989). Light inhibits sporulation of P. viticola (Brooks, 1979; Magarey and Butler, 1998). The sporangia remain viable for 4-8 days at high relative humidity and temperatures below about 22°C, and 1-2 days in hotter, drier conditions (Zachos, 1959; Blaeser and Weltzien, 1978; Kast and Stark-Urnau, 1999).
Individual lesions resporulate a number of times under favourable conditions, and can retain the potential to sporulate for 2-3 months (Zachos, 1959; Hill, 1989).
Secondary cycles of infection occur repeatedly throughout the growing season if weather conditions are favourable. To infect the plant, P. viticola requires surface wetness and temperatures of between 5-6°C and 30°C (Blaeser and Weltzien, 1977; Lalancette et al., 1988a). At optimal temperatures (approximately 20°C) infection can occur in 2 hours of surface wetness, with more hours of wetness required at non-optimal temperatures (Blaeser and Weltzien, 1977; Lalancette et al., 1988a).
When wet, the sporangia detach from their sporangiophores and germinate, releasing zoospores which are spread by windblown rain to new host tissue (Blaeser and Weltzien, 1978). Zoospores are released 30 to 180 minutes after the sporangia become wet (Langcake and Lovell, 1980; Kast and Stark-Urnau, 1999). Zoospores aggregate near stomata in groups of up to 10, possibly due to chemotactic signals (Royle and Thomas, 1973).
The penetration and colonization process is the same as described for the primary infection.
Means of Movement and DispersalTop of page Natural dispersal (non-biotic):
Blaeser and Weltzien (1978) observed sporangial dispersion only in wind-blown rain, not in the air. Zachos (1959) noted that high mortality of sporangia in dry conditions would prevent air dispersal. However, there is some evidence of long-distance (500-600 km) spore dispersal in regional air currents (Szoke et al., 1998).
Historically, agricultural practices have led to catastrophic dispersal of P. viticola. P. viticola is native to the north-eastern USA, and it is generally accepted that the pathogen spread to Europe on American grapes imported for use as rootstocks resistant to phylloxera (Daktulosphaira vitifolii [Viteus vitifoliae]) (Viennot-Bourgin, 1981). Downy mildew was first observed in France in September 1878 (Viala, 1893).
According to Viala (1893), the dangers of importing infected American
plants had already been asserted by Cornu several years earlier.
Strategies to prevent the spread of P. viticola on plant material include heat treatment of cuttings, maintaining disease-free tissue culture plantlets, and avoiding the spread of soil and leaf debris which may bear oospores (Emmett et al., 1992).
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||Pest or symptoms usually invisible|
|Leaves||hyphae; spores||Yes||Yes||Pest or symptoms usually visible to the naked eye|
|Seedlings/Micropropagated plants||hyphae; spores||Yes||Yes||Pest or symptoms usually visible to the naked eye|
|Plant parts not known to carry the pest in trade/transport|
|Stems (above ground)/Shoots/Trunks/Branches|
|True seeds (inc. grain)|
ImpactTop of page P. viticola has caused significant impacts on grape production since the 1800s. During the early culture of European varieties in the USA, yield losses were commonly 75% (Viala, 1893). Catastrophic losses arose in Europe in the late 1800s when P. viticola was first introduced on American rootstocks. The disease caused severe losses in favourable seasons and growers abandoned the use of several highly susceptible varieties (Viala, 1893).
Potential yield losses remain high, ranging from 50 to 100% under favourable conditions. The pathogen directly attacks the young inflorescences and fruit. Indirect damage occurs when severe foliar infections cause early defoliation, exposing the fruit to sunburn and reducing winter hardiness (Emmett et al., 1992). The estimated annual crop loss in an average year in Australia is $22.5 million (Australian dollars) with an additional $10 million spent on control measures. In wet years, direct crop losses may be as high as $64 million (Magarey and Butler, 1998).
DiagnosisTop of page Place leaves displaying oilspots and clusters displaying epinasty or oilspots in a lightly moistened plastic box or plastic bag and incubate overnight in the dark at 15-25°C. Fresh spores will be visible on infected tissue in the morning. Mature berries will not produce spores but their infected pedicels will sporulate.
P. viticola is an obligate biotroph and thus cannot be grown in culture.
Detection and InspectionTop of page Oilspots are readily visible in the field and will be noticed on inspection of the leaves (Seem et al., 1985). Monitoring procedures have been described to detect primary infection at levels as low as 0.01% (Seem et al., 1985). The upper leaf surfaces should be inspected for oilspots by gently moving the shoots and leaves, spending about 30 seconds per vine for 200-300 vines. If a primary infection is observed, that spot should be monitored for the first secondary infections, which are usually within 1 m of the original primary oilspots (Nicholas et al., 1994). Vines should be inspected every 1-2 weeks, or as necessary based on weather conditions (Nicholas et al., 1994). Computer training programs have been developed for improved accuracy of disease assessments (Nutter et al., 1998).
Similarities to Other Species/ConditionsTop of page Uncinula necator, causal agent of grape powdery mildew, also infects grape leaves and fruit clusters. Symptoms of early powdery mildew can resemble those of early downy mildew. Both diseases cause light-yellow patches on the upper leaf surface. However, powdery mildew is characterized by a flat, powdery, whitish-grey appearance on upper or lower leaf surfaces and on the fruit (Pearson, 1988; Nicholas et al., 1994). In contrast, downy mildew colonies are fluffy, a brighter white, and only found on the lower leaf surface.
Botrytis cinerea, the cause of Botrytis bunch rot or grey mould, produces fluffy colonies on grape clusters. However, Botrytis sporulation is brownish-grey or olive, unlike the bright white appearance of P. viticola (Bulit and Dubos, 1988; Nicholas et al., 1994).
Herbicides such as paraquat can cause yellowing of leaf tissue but these spots produce no spores (Nicholas et al., 1994).
Erinose mites cause a white growth on the undersides of leaves which can resemble downy mildew infections. The growths are only in blister-like galls and are not accompanied by oilspots (Nicholas et al., 1994).
Prevention and ControlTop of page
Disease pressure varies significantly with weather conditions. Management must be rigorous in wet climates such as eastern North America and parts of Europe, and during unusually wet seasons in dry locations such as California or Australia.
Chemical control has been an important control measure since the late 1800s, after the classic discovery of Bordeaux mix (copper sulfate plus lime) by Millardet in 1885 (Viala, 1893). Fungicides remain the most widely used management tool against P. viticola today (Lafon and Bulit, 1981; Lafon and Clerjeau, 1988; Emmett et al., 1992).
There are multiple pre- and post-infection chemicals available. Currently used pre-infection chemicals, or protectants, include: copper-based compounds (e.g. copper hydroxide, copper oxychloride and cuprous oxide); cyclimide (e.g. captan); dithiocarbamates (e.g. mancozeb); chlorothalonil; and dithianon.
Phenylamide compounds such as metalaxyl and benalaxyl have both pre- and post-infection activity (Nicholas et al., 1994) as do strobilurines such as azoxystrobin (Wong and Wilcox, 2001). Phosphonates provide post-infection activity (Nicholas et al., 1994).
First applications are generally advised at 3-8 inches of shoot-growth, immediate pre-bloom, and post-bloom to protect the young inflorescences and fruit. For the remainder of the season sprays may be based on a routine schedule (usually every 10-14 days) to maintain continuous protection of the vines. Alternatively, sprays may be based on disease risk as determined by weather conditions and forecasting models.
Resistance to phenylamide fungicides has been observed in Europe (Leroux and Clerjeau, 1985). The possibility of future resistance has triggered interest in monitoring and preventing this threat (Genet and Vincent, 1999; Wong and Wilcox, 2000). Gauthier and Amsden (2014) reported Qol-resistant downy mildew of grape in Kentucky.
The complex interactions of P. viticola with the environment render grape downy mildew a candidate for disease forecasting and modelling. Spray timing models have been used in Europe since the early 1900s (Muller and Sleumer, 1934; Populer, 1981). In more recent years computer-based forecasting models and decision tools have been developed in Italy (Rosa et al., 1993), France (Fouassier et al., 1997), Austria (Denzer, 1998), Germany (Huber et al., 1998), Hungary and Slovakia (Szoke et al., 1998), the USA (Park et al., 1997; Madden et al., 2000) and Australia (Magarey et al., 1991; Wachtel and Magarey, 1997). Most downy mildew models incorporate temperature, rainfall, relative humidity and leaf wetness, and more complex simulators incorporate information on host growth stage and varietal susceptibility. Models can be integrated into pre- or post-infection treatment strategies.
Resistant Crop Cultivars
Most cultivars of V. vinifera are highly susceptible to downy mildew, as are many cultivars of V. labrusca and interspecific hybrids.
Breeding for resistance to downy mildew has been ongoing since the 1930s (Matthews, 1981). Wild American Vitis species and related genera have been identified as possible sources of resistance. Putative resistance mechanisms include surface features such as leaf hairs or warty structures (Matthews, 1981; Staudt and Kassemeyer, 1995; Kortekamp and Zyprian, 1999), single-gene hypersensitive response (Matthews, 1981) and defence chemicals including peroxidase, viniferin, resveratrol and flavonoids (Langcake, 1981; Dai et al., 1995; Kortekamp et al., 1998).
Attempts to genetically engineer grapevine began in the late 1980s with Agrobacterium-mediated transformation (Baribault et al., 1989) and more recently with biolistics, or the 'gene gun' (Kikkert et al., 1996). Research continues on the introduction of genes for resistance to downy mildew and other pathogens (Reisch et al., 1996).
Cultural practices alone are unlikely to give sufficient control, especially under conditions favourable to downy mildew (Lafon and Clerjeau, 1988). However, cultural methods can influence disease development. High humidity and extended periods of leaf wetness enhance the development of downy mildew, thus techniques to promote air circulation and minimize surface wetness may reduce disease development (Palti and Rotem, 1981). Pruning and trellising methods which reduce canopy density decrease downy mildew levels (Wearing et al., 1999). Using furrow irrigation instead of overhead irrigation also reduces disease (Emmett et al., 1992). Primary infection may be prevented by ploughing to bury oospores in leaf litter, and avoidance of irrigating soil for long periods in order to prevent oospore germination (Palti and Rotem, 1981).
ReferencesTop of page
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