Phytophthora cambivora (root rot of forest trees)
- Summary of Invasiveness
- Taxonomic Tree
- Notes on Taxonomy and Nomenclature
- Distribution Table
- History of Introduction and Spread
- Risk of Introduction
- Habitat List
- Hosts/Species Affected
- Host Plants and Other Plants Affected
- Growth Stages
- List of Symptoms/Signs
- Biology and Ecology
- Natural enemies
- Means of Movement and Dispersal
- Seedborne Aspects
- Pathway Vectors
- Plant Trade
- Wood Packaging
- Impact Summary
- Economic Impact
- Environmental Impact
- Detection and Inspection
- Similarities to Other Species/Conditions
- Prevention and Control
- Links to Websites
- Distribution Maps
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PicturesTop of page
IdentityTop of page
Preferred Scientific Name
- Phytophthora cambivora (Petri) Buisman
Preferred Common Name
- root rot of forest trees
Other Scientific Names
- Blepharospora cambivora Petri
International Common Names
- English: ink disease; ink disease of chestnut; root rot of fruit disease
- Spanish: enfermidad de la tinta; tinta del castaño
- French: encre du chataignier; encre du noyer; maladie de l'encre du chataigner; maladie de l'encre du noyer
Local Common Names
- Germany: Tintenkrankheit: Esskastanie; Wurzelfaeule: Kastanie
- Italy: mal dell'inchiostro
- PHYTCM (Phytophthora cambivora)
Summary of InvasivenessTop of page P. cambivora is an invasive species that persists and spreads in different environments. Its invasiveness is increased by its capacity to survive as a saprophyte in the soil and to produce resting structures (oospores).
Taxonomic TreeTop of page
- Domain: Eukaryota
- Kingdom: Chromista
- Phylum: Oomycota
- Class: Oomycetes
- Order: Peronosporales
- Family: Peronosporaceae
- Genus: Phytophthora
- Species: Phytophthora cambivora
Notes on Taxonomy and NomenclatureTop of page
Petri (1917) was the first to describe this pathogen as Blepharospora cambivora. It was ascribed to the genus Phytophthora by Buisman (1927) who named it Phytophthora cambivora (Petri) Buisman. P. cambivora is classified in Phytophthora Group VI, which includes P. cinnamomi and P. cryptogea (Stamps et al., 1990).
DescriptionTop of page
Cultures of P. cambivora on PDA at 25°C are uniform, with moderate to profuse, fluffy, whitish aerial mycelium, similar to P. cambivora isolates from cherry on cornmeal agar at 21°C (Mircetich and Matheron, 1976). The mycelium is variable in size, with hyphae fairly uniform in diameter, slightly undulate and smooth, when old rather wide (7 µm) and thick-walled; some strains produce hyphal swellings with rounded irregular contours, not usually botriose, on rich media (Waterhouse and Waterston, 1966).
Sporangia, formed only in an aqueous solution, vary from broadly ovate, subglobose and obpyriform to ellipsoidal with well rounded bases, depending on the culture media used.
Sporangium size has been reported as 60-75 x 40-54 µm (Petri, 1917); usually 55-65 x 40-45 (max. 85 x 60) µm, no papillae, slight apical thickening; not shed (Waterhouse and Waterston, 1966). Another isolates obtained from chestnut had sporangia that were 48-83 x 32-61 µm (usually 59-64 x 40-45 µm) (Chitzanidis and Kouyeas, 1970). The length-breadth ratio of sporangia was reported to be 1.8 by Suzui and Hoshino (1979) and 1.4 by Oudemans and Coffey (1991).
Sporangiophores are 3-4 µm in diam., simple and unbranched, and can germinate by internal proliferation.
Oogonia are yellow to brown, spherical and produced terminally. They are normally produced abundantly only when grown with a suitable isolate of the opposite mating type of the same or a different species. Sex organs of P. cambivora were produced in about 10 days when isolates were grown together with certain strains of P. nicotianae var. parasitica (Waterhouse and Waterston, 1966), although some A2 isolates produced oogonia in single culture.
The diameter of oogonia is variable, ranging from 43 to 62 µm (Waterhouse and Waterston, 1966). The average diameter has been reported as 48, 44, 46.3, 37 and 40.5 ± 5.5 µm by Chitzanidis and Kouyeas (1970), Mircetich and Matheron (1976), Gerrettson-Cornell (1977), Suzui and Hoshino (1979) and Oudemans and Coffey (1991), respectively. The oogonial wall is 2 µm thick (Waterhouse and Waterston, 1966).
The oogonial wall is covered with irregularly distributed bullate protuberances (Waterhouse and Waterston, 1966). Antheridia are always amphigynous, one- or two-celled, with an average length of 25 µm (max. 35 µm) (Waterhouse and Waterston, 1966).
The diameter of the oospores is variously reported in the literature: 36 µm, with a wall thickness of 3 µm (Waterhouse and Waterston, 1966); 37-44 µm (Mircetich and Matheron, 1976); 35-39 µm (Gerrettson-Cornell, 1978b); and 33.8 ± 5.6 µm (Oudemans and Coffey, 1991).
DistributionTop of page P. cambivora is widely dispersed on all continents. Its distribution is listed by CMI (1984), and Erwin and Ribeiro (1996) provide a more detailed list.
Many identifications require confirmation because of the symptomatic analogy of P. cambivora to P. cinnamomi and other species of Phytophthora. Isolations must be carried out and isoenzyme and molecular characterization methods used for confirmation.
P. cambivora occurs in forest trees (chestnut, beech and oak) and in plantations and nurseries in many European countries. In the USA, recent attacks of P. cambivora have been reported from Washington State, Oregon and Michigan. In Australia, P. cambivora is active in almond plantations in the south of the country. In Asia, it has been reported from Korea and Japan.
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.Last updated: 10 Jan 2020
|Continent/Country/Region||Distribution||Last Reported||Origin||First Reported||Invasive||Reference||Notes|
|Mauritius||Present||CABI/EPPO (2013); CABI (Undated)|
|South Africa||Present||CABI/EPPO (2013)|
|India||Present||CABI/EPPO (2013); CABI (Undated)|
|Japan||Present||Suzui and Hoshino (1979); CABI/EPPO (2013)|
|-Peninsular Malaysia||Present||CABI/EPPO (2013)|
|South Korea||Present||Korean Society of Plant Protection (1972); CABI/EPPO (2013)|
|Taiwan||Present||Huang et al. (2012); CABI/EPPO (2013)|
|Turkey||Present||Uraz (1950); CABI/EPPO (2013)|
|Austria||Present||EPPO (2009); CABI/EPPO (2013)|
|Belgium||Present||EPPO (2009); CABI/EPPO (2013)|
|Czechia||Present||Černý et al. (2008); CABI/EPPO (2013)|
|Denmark||Present||Koch (1971); EPPO (2009); CABI/EPPO (2013)|
|Federal Republic of Yugoslavia||Present||Skoric (1946)|
|France||Present||1996||Blin (1922); EPPO (2009); CABI/EPPO (2013)|
|Germany||Present||Jung et al. (1996); EPPO (2009); CABI/EPPO (2013)|
|Greece||Present||CABI/EPPO (2013); Kailides (1967)|
|Hungary||Present||CABI/EPPO (2013); EPPO (2009)|
|Ireland||Present||Shafizadeh and Kavanagh (2005); CABI/EPPO (2013)|
|Italy||Present||CABI/EPPO (2013); Petri (1917)|
|Norway||Present||Talgø et al. (2006); CABI/EPPO (2013)|
|Poland||Present||2003||Orlikowski et al. (2002); EPPO (2009); CABI/EPPO (2013)|
|Portugal||Present||CABI/EPPO (2013); Petri (1917)|
|Romania||Present||Bolea et al. (1995); CABI/EPPO (2013)|
|-Russia (Europe)||Present||CABI/EPPO (2013)|
|Slovakia||Present||Kunca and Leontovyč (2005); CABI/EPPO (2013)|
|Spain||Present||CABI/EPPO (2013); CABI (Undated)|
|Sweden||Present||EPPO (2009); CABI/EPPO (2013)|
|Switzerland||Present||Faes (1941); CABI/EPPO (2013)|
|United Kingdom||Present, Localized||1994||Day (1938); EPPO (2009); CABI/EPPO (2013);|
|-British Columbia||Present||McIntosh (1964); CABI/EPPO (2013)|
|United States||Present||MILBURN and GRAVATT (1932); CABI/EPPO (2013)|
|-Alabama||Present||Grente (1961); CABI/EPPO (2013)|
|-Arizona||Present||CABI/EPPO (2013); CABI (Undated)|
|-Arkansas||Present||Grente (1961); CABI/EPPO (2013)|
|-California||Present||Mircetich and Matheron (1976); CABI/EPPO (2013)|
|-Georgia||Present||Grente (1961); CABI/EPPO (2013)|
|-Maryland||Present||Grente (1961); CABI/EPPO (2013)|
|-Michigan||Present||Bielenin and Jones (1988); CABI/EPPO (2013)|
|-Minnesota||Present||Schwingle et al. (2007); CABI/EPPO (2013)|
|-Missouri||Present||Grente (1961); CABI/EPPO (2013)|
|-Montana||Present||Grente (1961); CABI/EPPO (2013)|
|-New Jersey||Present||White (1937); CABI/EPPO (2013)|
|-New York||Present||Grente (1961); CABI/EPPO (2013)|
|-North Carolina||Present||Hwang et al. (2006); CABI/EPPO (2013)|
|-Oklahoma||Present||Grente (1961); CABI/EPPO (2013)|
|-Oregon||Present||Chastagner et al. (1995); CABI/EPPO (2013)|
|-South Carolina||Present||Grente (1961); CABI/EPPO (2013)|
|-Virginia||Present||Grente (1961); CABI/EPPO (2013)|
|-Washington||Present||Yamak et al. (2002); CABI/EPPO (2013)|
|-West Virginia||Present||CABI/EPPO (2013)|
|Australia||Present, Localized||CABI/EPPO (2013); CABI (Undated)|
|-New South Wales||Present||CABI/EPPO (2013)|
|-South Australia||Present||CABI/EPPO (2013); CABI (Undated)|
|-Western Australia||Present||CABI/EPPO (2013)|
|New Zealand||Present||CABI/EPPO (2013)|
|Papua New Guinea||Present||CABI/EPPO (2013)|
History of Introduction and SpreadTop of page P. cambivora is widespread in all continents as a result of international trade. It was imported into Europe in the 18th century and was described for the first time by Puccinelli (1859), causing 'ink disease' on chestnut trees, where the disease was most prominent. In 1876 it was observed in France, in the Pyrenees and some parts of Cévennes (Allain, 1935).
Risk of IntroductionTop of page P. cambivora is widespread in temperate areas of the world. To limit the spread of this pathogen agent, its occurrence must be carefully monitored in nurseries and nursery stocks.
HabitatTop of page
P. cambivora occurs worldwide in the soils of natural forests, agricultural fields and orchards. In Australia, stands of Eucalyptus paniculata are a favoured habitat. P. cambivora is commonly recovered from the soils of these stands, together with other species of Phytophthora, where it produces no pathogenic effects on the trees (Gerrettson-Cornell, 1978a, b). Damage has, however, been detected on almond stands in Australia. This difference in symptom expression could be due to the different susceptibility of these hosts to the pathogen.
In Europe, P. cambivora is widespread in the soil. In Italy, it infects the soils of chestnut stands and is the main agent of ink disease of chestnut (Petri, 1918; Biraghi, 1953; Turchetti, 1986; Anselmi et al., 1996).
In general, shallow soils are favourable to P. cambivora and to the disease it causes. A high concentration of plant roots is produced in shallow soils. Such roots become infected more rapidly, leading to a greater pathogen population. The effect of drought is more marked in shallow soils; both the soil and roots dry out more quickly and water stress is more likely to cause infection and death in hosts with fine roots. Shallow soils may lie on top of an impervious layer of clay or a rock base that impedes drainage, and can quickly become saturated with rainwater in which zoospores are dispersed. The roots are also predisposed to the disease because of the anaerobic conditions in the soil.
Habitat ListTop of page
|Terrestrial – Managed||Cultivated / agricultural land||Present, no further details||Harmful (pest or invasive)|
|Protected agriculture (e.g. glasshouse production)||Present, no further details||Harmful (pest or invasive)|
|Managed forests, plantations and orchards||Present, no further details||Harmful (pest or invasive)|
|Managed grasslands (grazing systems)||Present, no further details||Harmful (pest or invasive)|
|Disturbed areas||Present, no further details||Harmful (pest or invasive)|
|Rail / roadsides||Present, no further details||Harmful (pest or invasive)|
|Urban / peri-urban areas||Present, no further details||Harmful (pest or invasive)|
|Terrestrial ‑ Natural / Semi-natural||Natural forests||Present, no further details||Harmful (pest or invasive)|
|Natural grasslands||Present, no further details||Harmful (pest or invasive)|
|Riverbanks||Present, no further details||Harmful (pest or invasive)|
|Wetlands||Present, no further details||Harmful (pest or invasive)|
|Coastal areas||Present, no further details||Harmful (pest or invasive)|
Hosts/Species AffectedTop of page Damage caused by P. cambivora has been observed on apple, pear, peach and almond trees. Crandall et al. (1945) and Crandall (1950) revised the host list for P. cambivora and stated that many of the host records should be referred to P. cinnamomi, but this remains to be verified although there have been misreports in the past (Waterhouse and Waterston, 1966). Further information on these hosts can be found in Wicks et al. (1984, 1997), Wicks and Lee (1986), Wilcox (1993), Browne et al. (1995), Browne and Mircetich (1996), Jee et al. (1997, 2001).
Chastagner et al. (1995) first reported P. cambivora casing stem canker and root rot on Abies procera (Noble fir Christmas tree) in Oregon, USA. P. cambivora has been recovered on oak sites, as have other species of Phytophthora, but it is rarely a factor in oak decline (Anselmi et al., 1999; Jung et al., 2000; Osswald et al., 2001; Jonsson et al., 2003).
Damage on red raspberry was observed in Scotland, UK, during 1987-1989 (Duncan and Kennedy, 1987, 1989). There is currently a resurgence of ink disease of chestnut, caused by P. cambivora, in Italy (Turchetti, 1986; Turchetti and Parrini, 1993; Anselmi et al., 1996, 1999; Turchetti et al., 2000, 2003; Vettraino et al., 2001).
Pistachio has been identified as a host of P. cambivora in artificial inoculations tests (Kouyeas, 1973).
Host Plants and Other Plants AffectedTop of page
|Abies procera (noble fir)||Pinaceae||Other|
|Acer pensylvanicum (striped maple)||Aceraceae||Other|
|Acer platanoides (Norway maple)||Aceraceae||Other|
|Aesculus hippocastanum (horse chestnut)||Hippocastanaceae||Other|
|Castanea crenata (Japanese chestnut)||Fagaceae||Other|
|Castanea dentata (American chestnut)||Fagaceae||Other|
|Castanea pumila (Allegheny chinquapin)||Fagaceae||Other|
|Castanea sativa (chestnut)||Fagaceae||Main|
|Casuarina equisetifolia (casuarina)||Casuarinaceae||Other|
|Fagus sylvatica (common beech)||Fagaceae||Main|
|Lupinus albus (white lupine)||Fabaceae||Other|
|Malus (ornamental species apple)||Rosaceae||Main|
|Malus domestica (apple)||Rosaceae||Main|
|Persea americana (avocado)||Lauraceae||Other|
|Pistacia vera (pistachio)||Anacardiaceae||Other|
|Pisum sativum (pea)||Fabaceae||Other|
|Prunus armeniaca (apricot)||Rosaceae||Other|
|Prunus avium (sweet cherry)||Rosaceae||Other|
|Prunus campanulata (Taiwan cherry)||Rosaceae||Other|
|Prunus cerasus (sour cherry)||Rosaceae||Main|
|Prunus domestica (plum)||Rosaceae||Other|
|Prunus dulcis (almond)||Rosaceae||Main|
|Prunus persica (peach)||Rosaceae||Main|
|Prunus salicina (Japanese plum)||Rosaceae||Main|
|Pyrus communis (European pear)||Rosaceae||Other|
|Quercus cerris (European Turkey oak)||Fagaceae||Other|
|Quercus ilex (holm oak)||Fagaceae||Other|
|Quercus petraea (durmast oak)||Fagaceae||Other|
|Quercus pubescens (downy oak)||Fagaceae||Other|
|Quercus robur (common oak)||Fagaceae||Other|
|Quercus rubra (northern red oak)||Fagaceae||Other|
|Tanacetum cinerariifolium (Pyrethrum)||Other|
Growth StagesTop of page Flowering stage, Fruiting stage, Seedling stage, Vegetative growing stage
SymptomsTop of page In chestnut trees infected with P. cambivora the crown becomes transparent, unlike the crown of healthy trees or those infected by blight. Leaves lose their brilliant green and become glaucous, then yellowish; their growth is arrested, and the growth of branches decreases so that they appear shortened. Sometimes there is a precocious leaf fall (in August) leading to thinning of the canopy. Chestnut husks containing fruits are smaller than normal and are concentrated at the top of the crown, all at the same high level.
The roots become soft, spongy and brittle and exhibit deep purple to almost black areas, from which a blue-black, inky substance is exuded that stains the soil close to the roots and gives the disease its name (ink disease).
On young trees, infected or dead areas, which are slightly sunken with small cracks, appear at the base of the stem. At a later stage an elongated, sunken area with a distinct edge forms near the base of the tree. When the bark is removed from the base of the trunk or from the large roots, brown, necrotic areas are seen at cambium level; these areas, called 'flame blots' are shaped like an acute-angled triangle and spread from the roots to the stem. Sometime these blots don't appear because infection in the young roots kills the tree before they have time to develop (Biraghi, 1953). Shoots arising from the collar desiccate early. The inky fluid that often exudes from dead and dying bark at the base of trunk is another typical symptom.
At an advanced stage of disease, chestnut trees are girdled at the collar, and many branches begin to wither, resulting in the death of the tree. Pathogen development in the host is usually very fast and the death of the tree follows within a year of infection. However, the infection sometimes progresses more slowly and in these cases the trees generally die by the end of the second year. The difference in the duration of infection is due to the condition of the roots. Vigorous roots take longer to colonize than those that are weak or in poor condition.
Symptomatic trees are recognizable in mild winters and a succession of dry and wet spells favour infection. Winters that are drier and warmer than usual put the trees under water stress during the following spring when growth is resumed. Springtime is highly favourable to ink disease infection.
The disease has been observed in chestnut orchards and in abandoned stands where chestnut trees were still growing in competition with other invasive tree species (Turchetti et al., 2003).
Ink disease in beech differs slightly in symptomatology from that in chestnut (Day, 1938). In beech the fungus also infects the root, especially the small roots, but usually invades the tree at or near the collar root. It mainly colonises the cambium and inner bark, from where it spreads along the roots, and also occurs in the trunk at collar level. The rapid form of the disease, characterized by wilting of the leaves, is never observed in beech. The fluid that oozes from chestnut is blackish, whereas that exuding from beech is only discoloured, and often absent altogether. Sometimes infected beech trees recover and when this happens an actively developing callus grows beneath the dead bark and roots are sent down from the callus edge to the soil below. Adventitious roots have not been noticed on beech in response to ink disease, nor have stoolshoots, which are common in chestnut.
Other trees, such as walnut, cherry, apple, peach, plum and apricot are also susceptible to P. cambivora and other species of Phytophthora. Trees in the early stages of infection are difficult to distinguish from healthy trees. As the infection progresses, the leaves become small, chlorotic and droop, growing slowly on terminal shoots. A slow decline of infected trees is often observable; sometimes the trees die suddenly in early summer without manifesting any detectable symptoms. Both collar and root rot may occur in the same tree.
Symptoms of P. cambivora often resemble those caused by other root rot or collar rot pathogens. Decayed bark at the base of the trunk is a symptom of collar rot, which often starts at several points simultaneously. The infection progresses and the pathogen colonizes the cambium and cortical parenchyma of the host around the trunk until the lower part is entirely girdled.
Infected roots become brown, brittle and necrotic, in contrast to the soft rot typical of other root rot agents. Inhibition of the root system and necrosis of the lateral roots and taproot, also caused by other species of Phytophthora, can influence the vegetative condition of the tree. Some vigorous trees cope with this root reduction without any appreciable crown symptoms, even though their water relations and nutrition uptake are affected. As a consequence, the infection is difficult to detect in seedlings of walnut and other tree species after transplanting, and infected adult trees exposed to nutritional stress and unfavourable environmental conditions may exhibit a slow decline (Vettraino et al., 2003). The only way to identify the source of the decline is by isolating the causal agent from the roots of diseased trees.
List of Symptoms/SignsTop of page
|Growing point / dieback|
|Growing point / dieback|
|Growing point / discoloration|
|Growing point / discoloration|
|Growing point / lesions|
|Growing point / lesions|
|Growing point / wilt|
|Growing point / wilt|
|Leaves / yellowed or dead|
|Leaves / yellowed or dead|
|Roots / rot of wood|
|Roots / rot of wood|
|Stems / discoloration|
|Stems / discoloration|
|Stems / discoloration of bark|
|Stems / discoloration of bark|
|Stems / gummosis or resinosis|
|Stems / gummosis or resinosis|
|Stems / necrosis|
|Stems / necrosis|
|Stems / odour|
|Stems / odour|
|Whole plant / discoloration|
|Whole plant / discoloration|
|Whole plant / plant dead; dieback|
|Whole plant / plant dead; dieback|
Biology and EcologyTop of page The complete nucleotide sequence of internal transcribed spacer 1, 5.8 S ribosomal RNA gene and internal transcribed spacer 2 from P. cambivora (GenBank acc.no. AF266763) is given in Cook and Duncan (1997).
P. cambivora, like other Phytophthora species, reproduces asexually for a large part of its life cycle. Although mutations are possible, few mutants have been produced in experiments using mutagens and highly selective methods of mutant recovery.
Isolates of P. cambivora, like those of other heterothallic species, form oospores when mated with isolates of the opposite mating type belonging to the same species, and also when paired with compatible isolates of other species. Such interspecific matings produce hybrids. Interspecific matings rarely occur, but a hybrid, P. cambivora x P. fragariae, which is morphologically similar to P. cambivora but homothallic, was recently recovered on alder by Brasier (2001).
P. cambivora lives as a saprophyte in the soil, feeding on salts and dead organic material, often in competition with other microorganisms. Infection is favoured by moist and moderate climates. The optimum temperature for growth is 22-24°C, the maximum (cessation of growth) is >32°C and the minimum 2°C (Day, 1938; Erwin and Ribeiro, 1996).
P. cambivora survives in the soil in the form of mycelium, sporangia, zoospores and oospores. It is frequently found in soils with impeded drainage, due to a high water table or to a particular soil texture. Soils may drain poorly because they are compact (fine-textured silty or sandy) or clayey (Day, 1938). P. cambivora occurs in litters and in soils containing organic substrates for nutrients such as chestnut forest soils or oak forest soils, and like other Phytophthora species is tolerant to a wide range of pH: 3.8-7 (Jung et al., 2000).
P. cambivora mycelium is not resistant to drought, but it survives in the soil, even at low temperatures; the hyphae are killed at -8°C (Petri, 1918). It persists poorly in the soil during most of the year, being recovered only in late April-May and in late September-October, during periods of rain and mild temperatures (Vettraino et al., 2001).
The sporangia and zoospores produced by P. cambivora in humid soils are the main source of infection (Petri, 1918). Sporangia are produced abundantly by young mycelia, which become sterile when they are more than 1 month old (Petri, 1925). Not all hyphae of the mycelium produce sporangia, most are barren.
In addition to the presence of water and the interaction of water with aeration, temperature influences sporangium formation in the soil. P. cambivora forms sporangia at a wide range of temperatures: 9-30°C (Pfender et al., 1977; Sugar, 1977) but sporangia are not produced and mycelial growth is halted between 4 and 2°C (Petri, 1918). Cations such as Ca 2+, Mg 2+, Fe3+ and K+ have a stimulatory effect on the production of P. cambivora sporangia (Halsall and Forrester, 1977). Petri (1925) reported that sporangium production in daylight took four times as long as in the dark. Light in the blue region of the spectrum and UV light inhibited sporangium production, as demonstrated by Petri (1925) and Ribeiro et al. (1976) on four species of Phytophthora. Sporangia are therefore formed mainly at night, whereas zoospores form primarily by day, and their expulsion also occurs during the daytime (Petri, 1918).
The production of zoospores from sporangia is an important part of the Phytophthora life cycle because it allows the population to increase rapidly and disperse widely when films of free water are available. Zoospores are formed in the sporangia at temperatures between 8 and 12°C in the dark; they also form readily at 14°C in daylight. Under laboratory conditions zoospore production continued throughout simulated winter at 5-9°C (Petri, 1918). Humidity and temperature are the main physical factors governing the germination of sporangia. Sporangia can germinate directly, but in P. cambivora indirect germination is more common and occurs at 9-27°C (Sugar, 1977) with an optimum at 25°C. Pfender et al. (1977) and Sugar (1977) have investigated soil moisture requirements for the release of zoospores by sporangia.
Under favourable conditions, flagellate zoospores are expelled from the sporangia by osmosis. As the water enters the sporangium, the turgor pressure within the sporangium wall increases and causes the contents to be pushed out through the apical opening. Details of this process in Phytophthora are described and discussed by Gisi (1983).
Zoospores have a short life due to the lack of a cell wall. They normally swim in a helical path for several hours by means of two flagella of unequal length. Research by Newhook et al. (1981) indicated that although autonomous movement did occur, passive dispersal was the more important mode of transport. Under suitable temperature and moisture conditions, zoospores are widely dispersed by swimming toward nutrients exuded by host roots in aqueous solution and swimming away from micro-environmental hazards. Exudates overcome soil fungistasis and exert a chemotactic effect on the zoospores which gives to the motile zoospores a survival advantage over competitive micro-organisms.
The duration of zoospore motility in aqueous solutions varies under different conditions. Zoospores can swim for hours, but when they come in contact with a root, they appear to recognize certain substances in the root slime because they differentiate from the swimming to the cyst stage and germinate quickly. Zoospores in aqueous solutions migrate to the zone of elongation on the growing roots, where maximum exudation of plant materials occurs; the zoospores are positively attracted to this zone.
After germination, a germ tube is produced, which is initially single and then branched, but unicellular (Petri, 1918). It grows in a chemotactic and directional or tropic response towards the roots. Observations of the root surface suggest that the germ tubes penetrate the host directly under waterlogged soil conditions.
Chlamydospores are not known to be formed by P. cambivora, but oospores are formed and serve as resting structures. Oospores are the most resistant structures produced by P. cambivora, maintaining a source of inoculum when the pathogen has almost ceased to grow because of adverse environmental factors. Oospores can survive for long periods (more than 10 years according to Biraghi, 1953) and may be a source of genetic variation.
P. cambivora is self-sterile, and oospore formation occurs when the A1 and A2 (opposite) mating types grow together (heterothallic species). P. cambivora may form oospores as a result of selfing when stimulated by ubiquitous soil fungi, such as Trichoderma (Brasier, 1971). In laboratory tests, some A2 type colonies are homothallic as well as heterothallic, producing sex organs on solid media within 10 days of inoculation, but A1-types are sterile (Gerrettson-Cornell, 1977).
P. cambivora is of the amphigynous (around the male) sexual type: the antheridium, or male gametangial tip, is penetrated by the oogonium, the female gametangial incept (Petri, 1918). When the oogonial tip and antheridial incepts make contact, the antheridium becomes fused to the oogonium and meiosis occurs. One or more haploid antheridial nuclei pass into the oogonium, which expands to a globose structure, in which a single spore forms. Oospores go through a maturation phase prior to germination. They germinate under favourable environmental conditions after the activation phase (swelling of the oospore within the oogonium and dissolution of the inner oospore wall) either by producing a single germ tube or by producing four zoospores (Petri, 1917). When the oospores germinate near a susceptible root, the root is infected and the infection process begins.
The difficulty in isolating P. cambivora from the soil may be due to the low production of the oospore resting structures (Vettraino et al., 2001). Other limiting factors are the self-sterility of this species and the necessary presence of mating types. In addition, the percentage of oospore germination is low at 1-5% (Boccas, 1981). These considerations suggest that P. cambivora occupies the infected root before it dies and is decomposed, then spreads and establishes new infections during a limited period of the year, when environmental and especially weather conditions are favourable to the production and survival of zoospores.
Natural enemiesTop of page
Means of Movement and DispersalTop of page The natural dispersal of P. cambivora is by means of motile zoospores. Standing water is an efficient dispersal medium. The spread of P. cambivora is both directly and indirectly favoured by man.
Seedborne AspectsTop of page P. cambivora is not seedborne, but can be spread by infected seedlings.
Pathway VectorsTop of page
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|
|Bark||hyphae||Yes||Yes||Pest or symptoms usually visible to the naked eye|
|Growing medium accompanying plants||hyphae; spores||Pest or symptoms usually invisible|
|Roots||hyphae||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Seedlings/Micropropagated plants||hyphae; spores||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Stems (above ground)/Shoots/Trunks/Branches||hyphae||Yes||Pest or symptoms usually visible to the naked eye|
|Wood||hyphae||Yes||Yes||Pest or symptoms usually visible to the naked eye|
|Plant parts not known to carry the pest in trade/transport|
|Fruits (inc. pods)|
|True seeds (inc. grain)|
Wood PackagingTop of page
|Wood Packaging liable to carry the pest in trade/transport||Timber type||Used as packing|
|Solid wood packing material with bark||No|
|Solid wood packing material without bark||No|
Impact SummaryTop of page
|Fisheries / aquaculture||None|
|Fisheries / aquaculture||None|
Economic ImpactTop of page It is hard to assess the damage caused by P. cambivora because the host range varies between continents. In Australia, P. cambivora severely damages fruit trees such as apple, cherry and almond (Wicks et al., 1984; Wicks and Lee, 1985). Apple trees are affected in Japan and Korea (Nakazawa et al., 1981; Jee et al., 1997). In North America, P. cambivora infects orchards of apple and cherry (Bielenin and Jones, 1988; Browne and Mircetich, 1988) and Chastagner et al. (1995) reported that some plantations of Noble fir (Abies procera) Christmas trees were also susceptible to the fungus, with a net loss of production.
P. cambivora has been detected on apple grafts and cherry together with other species of Phytophthora in Switzerland (Bolay, 1992) and causes severe damage to red raspberry in Scotland, UK, negatively influencing the economy of these crops (Duncan et a1., 1987). The production of nuts and timber was reduced on walnut plantations in Italy as a result of P. cambivora and other species of Phytophthora (Belisario et al., 1997, 2001; Vettraino et al., 2003). Rhodendron and horse chestnut are damaged by the fungus, negatively influencing the economy of nursery growers. Beech stands are damaged in the UK and Germany (Day, 1938; Hartmann and Blank, 1998) and chestnut trees are damaged in Italy, France, Spain, Portugal and Greece. The economic repercussions on the timber and fruit trade are evident. Edible mushrooms are reported to decrease in number in stands severely infected with P. cambivora.
P. cambivora and other species of Phytophthora particularly affect ornamental plants and the nursery industry. As the symptomatology of P. cambivora is similar to that of other species of Phytophthora, it is difficult to value losses attributable specifically to P. cambivora, particularly those from root and collar rot of apple, pear and other fruit trees.
Environmental ImpactTop of page P. cambivora, like other species of Phytophthora, can cause genetic erosion of different species and cultivars of fruit trees, but this risk can be overcome by using resistant rootstocks.
The decline of eucalyptus in Australia and oak forests in Europe due to P. cambivora is fortuitous, and its importance marginal. The impact of P. cambivora on beech and chestnut forests is significant because it puts the stability and evolution of these ecosystems at risk. Trees killed as a result of P. cambivora on mountain slopes or crests can compromise the stability of soils, leaving them exposed to erosion from runoff rainwater. In some stands in Italy, P. cambivora infection has led to the natural replacement of chestnut trees. Dead chestnut stands have gradually been invaded by more resistant oaks (Quercus pubescens), leading to the formation of new oak woods (Turchetti, 1986). Chestnut is also the main ectomycorrhizal host in most of these stands, and when it disappears the mushroom population suffers. P. cambivora, like other species of Phytophthora, colonizes anaerobic soils that are unfavourable to other fungi including Phytophthora antagonists, which suffer a great decrease in such soils.
DiagnosisTop of page
P. cambivora grows on the media used for species of Phytophthora such as oatmeal agar, cornmeal agar, potato dextrose agar (PDA) and V8 juice. Carrot agar (CA) has also been used (Brasier, 1969). It grows well at 25°C producing an abundance of fluffy whitish mycelium.
Selective media for the isolation of P. cambivora from soil have been devised, containing the antimicrobial agents penicillin G, streptomycin, neomycin, pimaricin, tetracycline, nystatin, benomyl and pentachloronitrobenzene (PCNB) (Moreira and Ferraz, 1993).
P. cambivora is moderately resistant to hymexazol (50mg/ml) (Tay et al., 1983). Another selective medium, PARBhy, contains pymaricin (10 mg), ampicillin (250 mg sodium salt), rifampicin (10 mg), hymexazol (50 mg), benomyl (15 mg), malt extract (15 g), agar (20 g) and water (1000 ml) (Robin, 1991). PARPNH was reported to be a selective medium by Jung et al. (1996). It contains multivitamin juice, agar, CaCO, pimaricin, ampicillin, rifampicin, nystatin, hymexazol and pentachloronitrobenzene. All antibiotics must be added aseptically to the medium after sterilization in an autoclave.
A simple way to isolate P. cambivora and other species of Phytophthora is by using a bait in the soil. Baits are often apple or pear fruits buried in the upper 20-30 cm of moist soil around diseased trees. After 5-7 days the apple or pears are excavated. Fruits showing symptoms of rot are transferred to the laboratory, washed with running demineralised water and surface sterilized with 95% ethanol. Tissue fragments are cut from the advancing margin of the rot, plated on PARPNH and incubated at 20°C in the dark (Jung et al., 1996).
Another method uses 'Cunningham White' rhododendron leaves (Themann and Werres, 1998) or young oak leaves (Quercus robur) as bait (Jung et al., 1996). Q. robur sprouts and leaflets that are a few days old can be used as baits by floating them on sample volumes of 30 cm³ soil flooded to 3 cm depth in 12-cm Petri dishes.
Sprouts and leaflets that develop black spots or turning brownish or blackish are washed in running demineralised water and dipped in 95% ethanol. Small segments of the discoloured tissue are placed on PARPNH and incubated at 20°C in the dark (Jung et al., 1996; Vettraino et al., 2003).
Developing colonies of Phytophthora are transferred to, and maintained on, malt agar (MA) and multivitamin agar (MVA) (Jung et al., 1996).
Isolations can be made directly from host tissues. Successful isolation of P. cambivora depends on the season, presumably influenced by the weather. The best time for isolation is late winter and spring; isolation becomes less effective as the summer progresses although autumn is also favourable for P. cambivora and other species of Phytophthora.
P. cambivora was isolated from chestnut by placing fragments of infected tissues on Petri dishes containing V8 juice media or PDA (Turchetti and Parrini, 1993). Infected tissues samples were collected from the collar region at the margin between the necrotic and healthy tissue, as suggested by Petri (1917). Sampling from this specific area is an important factor for isolation success. P. cambivora is most active in this margin, whereas other antagonistic or saprophytic fungi predominate elsewhere in the necrotic area.
Robin et al. (2001) reported a selective medium for the isolation P. cambivora from host tissues: MA (malt agar) + PARBhy (10 p.p.m. pimaricin, 250 p.p.m. ampicillin, 10 p.p.m. rifampicin, 15 p.p.m. benomyl, 50 p.p.m. hymexazol).
Serology has advantages over conventional methods for detection of species of Phytophthora: it is less time-consuming and more accurate (Erwin and Ribeiro, 1996). Cytoplasmic and cell wall antigens specific to P. cambivora and to each of three species of Phytophthora have been recovered so P. cambivora can be distinguished serologically from the other species (Halsall, 1976). Improvements have been made and the DAS-ELISA (double antibody sandwich enzyme-linked immunosorbent assay) technique is suitable for mass testing plant material and excellent for discriminating between samples from diseased and healthy plants (Olsson, 1999). DAS-ELISA polyclonal antisera were used to detect species of Phytophthora in recycling water in nurseries (Themann et al., 2002). Commercial kits are available for the serological detection of Phytophthora species. However, the sensitivity and specificity of the methods have to be improved for serological diagnosis to be used to classify Phytophthora species.
Starch gel electrophoresis could also be useful for detection of Phytophthora species. P. cambivora, P. cinnamomi and P. cactorum were compared using 18 isenozyme loci, and this analysis clearly separated the three species (Oudemans and Coffey, 1991).
DNA technologies have recently improved and PCR-based diagnostics can now detect Phytophthora both in plant material and in the soil and can therefore be used to identify species (Vettraino et al., 2009). Protocols to identify the pathogen and sequences are available (Cook and Duncan, 1997). The internal transcribed spacer regions (ITS1 and ITS2) of the ribosomal RNA gene repeat from Phytophthora species have been amplified using PCR and sequenced.
PCR has been used to detect species of Phytophthora that cause root rot diseases in European forests. Sequences for P. cambivora and other Phytophthora species have been detected using this diagnostic system.
The complete nucleotide sequence of ITS1, the 5.8 S ribosomal RNA gene and ITS2 of P. cambivora (GenBank acc. no. AJ007040) is given by Schubert et al. (1999).
Detection and InspectionTop of page P. cambivora is difficult to diagnose from dead trees even if the tree has died only recently. Mycelium in dead host tissues is impermanent, and the fungus is very difficult to isolate from dead stems or trunks. It is never recovered from soils around host tree with slight or no symptoms. It is best to inspect trees for P. cambivora when they are symptomatic but still green. Collar rot is a visible symptom, but can also be due to many other factors such as mechanical wounding, cracking, insect attack and other injurious agents. A pathogenic effect similar to that of ink disease from P. cambivora is, infrequently, produced by P. cactorum and P. citricola on chestnut (Biocca et al., 1993). This being so, it is difficult to identify P. cambivora positively without an isolation test. Infected areas on the stem, initiating from the collar, are visible as slight depressions in the bark. One useful way to detect the fungus is to remove the bark with a blade; if the tree is infected this will reveal the dark brown, flame-shaped lesions typical of P. cambivora underneath. Sometimes the progress of the parasite into the host roots is so rapid that the entire root system collapses before necrotic lesions have time to appear on the collar.
Many lateral roots have to be destroyed before infected trees show aboveground symptoms, and this fact must be taken into account when making a diagnosis. Another diagnostic character is the presence of husks at the top of the crown, all at the same height.
The spatial distribution of diseased trees, whether growing as single trees or as groups, in humid places or in valley bottoms, and on slopes or on crests of mountains can also be a useful diagnostic indicator.
For other hosts, inspection of the foliage and trunk for symptoms is basic for a diagnosis.
Seedlings of chestnut and other tree species can become infected with ink disease, producing brown areas on the young root or the collar.
All of these symptoms and characters are common for both P. cambivora and P. cinnamomi and so an accurate identification requires isolation of the pathogen. In Italy, P. cinnamomi is not easily recovered from forests, so any symptoms found are almost certainly those of P. cambivora. However, P. cinnamomi was recently recovered in a chestnut coppice in the Lazio region, probably introduced by an infected seedling from a nursery (Cristinzio, 1986). P. cinnamomi has also been detected in chestnut seedlings in nurseries (Turchetti and Parrini, 1993) and in walnut nurseries and plantations. It was probably introduced to these plantations with infected seedlings planted for reforestation on ex-agricultural lands, so P. cinnamomi is now starting to spread in Italy (Belisario et al., 1997, 2001, 2002; Vettraino et al., 2003).
Similarities to Other Species/ConditionsTop of page
P. cambivora is often recovered from chestnut, walnut and oak stands in association with Phytophthora citricola, P. cactorum, P. cryptogea, P. gonapodyides, P. cinnamomi and other species of Phytophthora (Jung et al., 1996, 2002; Vettraino et al., 2001, 2003). P. cambivora is classified in Group VI of the Phytophthora together with P. cinnamomi, P. cryptogea, P. drechsleri and P. gonapodyides (Stamps et al., 1990). The phylogenetic analysis of Phytophthora species on the basis of sequences of ribosomal RNA have distinguished two clades amongst species with non-papillate sporangia: one consisting of P. fragariae, P. cambivora and P. cinnamomi and the other comprising P. cryptogea, P. megasperma and P. drechsleri.
P. fragariae causes root rot on red strawberry as does P. cambivora on its hosts. A new aggressive Phytophthora pathogen on alder in Europe was reported comprising a range of heteroploid-interspecific hybrids consisting of a species resembling P. cambivora and an unknown taxon similar to P. fragariae (Brasier et al., 1999). This new Phytophthora has morphological characters similar to P. cambivora, but is homothallic. Two newly found species of Phytophthora, P. europaea and P. uliginosa, isolated from oak stands, are also similar to P. cambivora and P. fragariae in morphological characters such as having non-papillate sporangia. However, the antheridia of P. cambivora and P. fragariae are 2-1-celled, amphigynous, whereas the others are paragynous, and only P. cambivora is heterothallic (Jung et al., 2002).
Phytophthora cactorum infects chestnut, on which it causes a limited root rot (Curzi, 1933; Smith, 1937; Cristinzio and Vernau, 1954; Vettraino et al., 2001), but it has not yet been reported in the rooting zone of chestnut trees in areas where ink disease occurs. It differs from P. cambivora in that it produces chlamydospores, has paragynous antheridia and is homothallic. Extensive isoenzyme analysis with starch gel electrophoresis showed clearly that P. cambivora is quite different from P. cactorum and P. cinnamomi (Erwin and Ribeiro, 1996).
P. cambivora used to be considered similar to P. cinnamomi, but differs in the morphology of its cultures and mycelia. P. cinnamomi has coralloid hyphae, whereas the hyphae of P. cambivora are smooth. P. cinnamomi produces lightly pigmented, botryose, thin-walled hyphal swellings and chlamydospores. Some oogonia of P. cambivora are verrucose, but P. cinnamomi oogonia are always smooth walled. P. cambivora is also serologically different from P. cinnamomi and other species of Phytophthora (Halsall, 1976).
Symptoms of P. cinnamomi on plants are fairly generic and confused with those caused by other species of Phytophthora that produce root rots. P. cactorum causes a limited root rot similar to that of P. cambivora and P. cinnamomi infects chestnut trees and other hosts producing the same symptoms as P. cambivora. P. cambivora causes root rot in Italy but P. cinnamomi does not occur in this country although it has been recovered from Ericaceae, Chamaecyparis and chestnut in nurseries (Gullino and Garibaldi, 1987; Parrini et al., 1997).
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.
Control of P. cambivora is difficult to achieve because the fungus can survive for several years as resting oospores in the soil or in infected host tissues (Biraghi, 1953; Turchetti, 1986, Vettraino et al. 2001). The aim of control strategies is to limit the damage caused and to prevent the natural spread of the fungus. These goals must be met in different ways:
Petri (1918) suggested pruning infected chestnut trees to stimulate the root system to produce new lateral roots, a method that has proved suitable for other hosts, especially fruit trees. Felling severely infected trees at ground level is another possibility, as the tree stump will produce new shoots indicating that the root system has recovered. Other treatments specific to chestnut have been proposed, for example, Gandolfo, which consists of stripping the collar and roots of infected trees and leaving them exposed to low winter temperatures. This kills the mycelium of P. cambivora and enables the trees to recover. A similar method was proposed by Urquijo-Landaluze (1941) in Spain, consisting of exposing and cleaning the collar and roots and treating them with copper salts. However, these methods are difficult to employ in chestnut stands and orchards.
Disease prevention is very important, especially in nurseries. Forest and fruit seedlings must be grown in pathogen-free soils and the residual roots of seedlings must be completely eliminated before they are grown. Soil must be well drained and water stagnation prevented, because excess soil moisture favours the fungus. Seedlings must not be watered with water originating from infected areas. It is advisable to use of pathogen-free planting material, potting mixes and containers.
Jeffers and Aldwinckle (1988) found P. cambivora and P. cactorum in apple rootstock from nurseries in the USA and Europe. Both soil and seedlings should be sampled regularly for diagnosis using baits, isolation from host tissues and PCR-based technologies to improve detection. Infected seedlings must be eliminated at a primary stage and any remaining healthy seedlings treated with metalaxyl or fosetyl-Al.
Mulches have been tested against some species of Phytophthora including P. cambivora (Merwin et al., 1992; Fraser et al., 2001). In Australian forest sites dominated by Acacia pulchella, P. cinamomi was significantly reduced and mulching investigations carried out by D'Souza et al. (2001) are continuing. The use of chicken manure to control P. cinnamomi was tested in Australia by Aryantha et al. (2000). In Italy, control trials, in which mulching was combined with organic amendments to treat chestnut trees over 3 years, recovered most of the trees (Turchetti et al., 2000). Mixtures of amendments (cow manure, chicken manure and biological amendment) can provide effective biological control that will preserve the environment of chestnut stands (Turchetti et al., 2003).
Mychorrhizal fungi may prove effective in controlling P. cambivora and may help in soil sanitation and the production of healthy seedlings, especially chestnut seedlings (Branzanti et al., 1999).
Soil solarization, in which the soil is heated to 8-10°C above ambient temperature by covering with transparent plastic sheeting, was studied as a means of controlling P. cambivora in naturally infected almond and cherry orchards. In field experiments, soil temperatures above 40°C were rarely reached at a depth of 10 cm in solarized soil. This treatment controlled P. cambivora in an irrigated cherry orchard but not in non-irrigated almond orchards. Although solarization did not control established P. cambivora infections, it significantly increased the growth of the trees (Wicks, 1988b).
The use of resistant rootstocks is another means of controlling P. cambivora. In chestnut, high levels of resistance have been reported for Asiatic chestnut (Castanea crenata) and the hybrids C. sativa x C. crenata. Studies on the resistance of these hybrids to P. cambivora and P. cinnamomi are continuing and some offer interesting perspectives with regard to their graft compatibility with cultivated varieties and to productivity (Cristinzio and Grassi, 1993; Salesses et al., 1993; Pereira et al., 1995; Breisch and Hennion, 2004).
Nut trees, fruit trees and red raspberry are all affected by species of Phytophthora including P. cambivora, which is not host specific, and when rootstocks are chosen for an orchard, the Phytophthora species occurring in that orchard should always be considered. Mircetich and Matheron (1976) and Wilcox and Mircetich (1985) reported that although cherry Mazzard rootstock was resistant to P. cambivora, it was as susceptible as susceptible Mahaleb rootstock to other species of Phytophthora (P. cinamomi, P. citricola, P. cryptogea, P. drechsleri and P. megasperma). Cherry rootstock CAB-6P was resistant to seven species of Phytophthora including P. cambivora, but it was susceptible to P. cactorum, P. cryptogea, P. citrophthora and P. citricola (Thomidis and Sotiropoulos, 2003). In an earlier study, Thomidis et al. (2002) found that selected apple and pear rootstocks were immune to nine species of Phytophthora, but were susceptible to infection by P. cactorum and P. citricola.
The resistance of rootstocks is influenced by environmental factors. Tests carried out in California, USA, found that Phytophthora infection on apple rootstock EMLA 106 was most severe in late spring, and less severe in August to October (Brown and Mircetich, 1996). Rootstocks are also affected by conditions that favour P. cambivora, and the virulence of isolates of different species of Phytophthora involved in the disease. In an Australian study, the cherry rootstocks Mazzard and Mahaleb were not resistant to P. cambivora when grown in infested soils (Wicks, 1989).
Metalaxyl and fosetyl-Al were tested for control of P. cambivora in chestnut trees in Greece. Two applications were used, one in the spring and one in autumn (Skoudridakis and Bourbos, 1990). Both fungicides were applied as soil drenches and fosetyl-Al was also applied as a foliar spray. When applied as a soil drench, fosetyl-Al was significantly less effective than any of the other treatments.
Metalaxyl effectively controlled species of Phytophthora on red raspberry (Duncan and Kennedy, 1987) whereas fosetyl-Al and metalaxyl controlled these pathogens on cherry (Bielenin and Jones, 1988; Matheron et al., 1988)). In Australia, these fungicides and phosponate, used as a soil drench and a foliar spray, effectively controlled species of Phytophthora infecting cherry and almond (Wicks and Lee, 1985; Wicks, 1988a; Wicks and Hall, 1988, 1990).
Phenylamide fungicides are effective against different species of Phytophthora, and sodium tetrathiocarbonate shortened the duration of zoospore motility of six species of Phytophthora including P. cambivora as well as reducing sporangium production in the soil (Matheron and Matejka; 1988; Oros and Komives, 1991; Browne and Mircetich, 1992).
Dinitroaniline herbicides significantly reduced the activity of P. cambivora and three other species of Phytophthora on Mahaleb cherry seedlings (Wilcox, 1996).
Trichoderma harzianum and T. koningii, which are antagonistic to P. cambivora, are included in a commercial product 'Promot'. Chestnut trees in Greece treated with this biological product received a significant degree of protection against P. cambivora (Bourbos and Metzidakis, 2000).
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