Apiognomonia errabunda (anthracnose)
- Summary of Invasiveness
- Taxonomic Tree
- Notes on Taxonomy and Nomenclature
- Distribution Table
- Risk of Introduction
- Habitat List
- Hosts/Species Affected
- Host Plants and Other Plants Affected
- Growth Stages
- List of Symptoms/Signs
- Biology and Ecology
- Notes on Natural Enemies
- Means of Movement and Dispersal
- Seedborne Aspects
- Pathway Vectors
- Plant Trade
- Wood Packaging
- Impact Summary
- Economic Impact
- Environmental Impact
- Detection and Inspection
- Prevention and Control
- Distribution Maps
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PicturesTop of page
IdentityTop of page
Preferred Scientific Name
- Apiognomonia errabunda (Roberge ex Desm.) Höhn. 1918
Preferred Common Name
Other Scientific Names
- Apiognomonia quercina (Kleb.) Höhn. 1920
- Apiognomonia tiliae (Rehm) Höhn. 1920
- Discula quercina (Westend.) Arx 1957
- Discula umbrinella (Berk. & Broome) M. Morelet 1973
- Gloeosporidium tiliae (Oudem.) Petr. 1922
- Gloeosporium fagi (Roberge ex Desm.) Westend.
- Gloeosporium quercinum Westend. 1854
- Gloeosporium tiliae Oudem. 1873
- Gloeosporium umbrinellum Berk. & Broome 1866
- Gnomonia errabunda (Roberge ex Desm.) Auersw. 1869
- Gnomonia quercina Kleb. 1918
- Gnomonia tiliae Rehm
International Common Names
- English: anthracnose: oak; beech leaf spot; beech shoot dieback; oak leaf spot; oak shoot dieback
- Spanish: antracnosis de la encina
Local Common Names
- Germany: Blattbraeune: Eiche; Blattfleckenkrankheit: Eiche
- GNOMEB (Gnomonia errabunda)
Summary of InvasivenessTop of page A. errabunda and A. quercina are invasive species both in Europe and the USA for the following reasons: the pathogen has a three-way survival capability, with a conidial form on the twigs, leaves and buds of the tree; a sexual form with perithecia on fallen leaves, and an active mycelium on the leaves, buds and twigs in Mediterranean countries with mild winters; it produces a great number of conidia, 49,300/m³ air, at a distance of 10 m from the inoculum source after 10 days of sporulation, and 2400 conidia/m³ air at a distance of 1000 m from the source after 40 days of sporulation; it is highly adaptable to a wide variety of different climates in Europe, from parts of Russia through the countries of north-eastern Europe to the Mediterranean, and in North America, from Canada down to the Gulf of Mexico along the Pacific coast; and it has a very extensive host range.
Taxonomic TreeTop of page
- Domain: Eukaryota
- Kingdom: Fungi
- Phylum: Ascomycota
- Subphylum: Pezizomycotina
- Class: Sordariomycetes
- Subclass: Sordariomycetidae
- Order: Diaporthales
- Family: Gnomoniaceae
- Genus: Apiognomonia
- Species: Apiognomonia errabunda
Notes on Taxonomy and NomenclatureTop of page Apiognomonia errabunda is seen as a complex of species. Formerly classed in the genus Gnomonia, it was transferred to Apiognomonia by Barr (1978). It is associated with the anamorph Discula umbrinella (synonym Gloeosporium umbrinellum) and causes leaf spot and shoot dieback on numerous broadleaves in Europe and North America.
Apiognomonia errabunda sensu stricto is essentially linked to beech (Fagus sylvatica). Oak (Quercus spp.) on the other hand is infected by another species in the same complex, Apiognomonia quercina, with its anamorph Discula quercina (synonym Gloeosporium quercinum). The Linden tree (Tilia europaea) is infected by Apiognomonia tiliae, with the anamorph Gloeosporidium tiliae, and plane (especially London plane = Platanus x acerifolia) by Apiognomonia veneta, with the anamorph Discula platani. Apiognomonia veneta is considered to be a distinct species from A. errabunda by Monod (1983) and is treated in a separate datasheet in this Compendium.
The genus Apiognomonia was created by Höhnel in 1918, and many species found on Betula, Fagus, Fraxinus, Platanus, Quercus and Tilia have over time been referred to it. Apiognomonia was the teleomorph (or sexual form) and was associated with a variety of genera representing anamorph forms (von Arx, 1970).
On beech, the following species have, at some time, been related to the teleomorph A. errabunda, with the anamorph D. umbrinella: Labrella fagi, Gloeosporium fagi, Gloeosporidium fagi, Gloeosporium exsiccans, Discula fagi, Gloeosporium fagi, Myxosporina fagi, Gloeosporium fagicola and Gloeosporium fuckelii.
On oak, the species related to the teleomorph A. quercina, with the anamorph D. quercina (syn. G. quercinum) are: Gloeosporium quercigenum Westend., Myxosporina quercina, Gloeosporium canadense, Gloeosporium cecidophilum, Gloeosporium divergens, Gloeosporium intumescens, Gloeosporium marginans, Gloeosporium nervicola, Discula quercina, Gloeosporium quercuum, Gloeosporium umbrinellum and Gloeosporium umbrinellum.
On linden, the following species have been ascribed to the teleomorph A. tiliae, with the anamorph Gloeosporidium tiliae: Gloeosporium tiliae, Myxosporina tiliae, Gloeosporidium tiliae and Gloeosporium tiliaecola.
Thirty isolates of D. umbrinella found on beech, chestnut and oak have been studied using RAPD and RFLP markers. The isolates fell into four groups depending on the host tree on which they were found, which shows the high host specialization of the various members of the A. errabunda complex (Haemmerli et al., 1992).
Many anamorph forms are related to the Apiognomonia complex. Those of the genus Discula are listed here, only those occurring on forest trees or other tree species, without giving the synonyms by which they may have been designated in the past:
Discula acerina on Acer spp.
D. betulina on Betula spp.
D. betulina var. betulina on Betula spp.
D. betulina var. macrocarpa on Betula spp.
D. brunneotingens on Pinus sp.
D. campestris on Acer spp.
D. coloradensis on Populus grandidentata/P. angustifolia
D. cytospora on Populus sp.
D. cytosporea on Populus spp.
D. effusa on Pyrus sp. and Malus sp.
D. fagi on Fagus spp.
D. fraxinea on Fraxinus sp.
D. ochrostica on Eucalyptus spp.
D. orni on Fraxinus spp.
D. pinicola on Picea spp. and Pinus spp.
D. pinicola var. mamosa on Pinus spp.
D. pinicola var. pinicola on Pinus spp.
D. platani on Platanus spp.
D. quercicola on Quercus spp.
D. sassafras on Sassafras sp.
D. sanguisorbae on Sorbus sp.
D. terminaliae on Terminalia catappa
Saccardo (1884) also has: D. platyspora on Platanus sp., D. magnoliae on Magnoliae sp., D. citri on Citrus sp., D. macrosperma on Salix sp., D. rugosa on Acer sp., D. obscura on Acer sp. and D. packiana on Ostrya spp.
A recently identified species that deserves a special mention is Discula destructiva, the most serious disease agent on Cornus florida, C. nuttallii and C. kousa in Canada and the USA. The teleomorph of D. destructiva is currently unknown (Mielke and Daughtrey, 1989); all attempts to reproduce it in the laboratory have so far been unsuccessful. Some authors have suggested that the teleomorph is also part of the Apiognomonia errabunda complex, but such a relation has never really become accepted (Mielke and Langdon, 1986; Schneeberger and Jackson, 1989). At present D. destructiva cannot be considered conspecific with the anamorph forms of the genus Discula related to A. errabunda: its cultural characteristics and range of hosts are too different (Neely and Himelick, 1967; Monod, 1983).
DescriptionTop of page A. errabunda sensu stricto (anamorph Discula umbrinella)
On senescent or dead leaves, A. errabunda produces spherical perithecia, almost black in colour and shiny, with a diameter of 130-450 µm and containing two-celled hyaline ascospores measuring 2-4 x 12-16 µm.
Acervula of D. umbrinella are subepidermal and erupt in the form of dark, waxy discs or pads, with a diameter of ca 240-260 µm. Conidia are one-celled, hyaline, ovoid or oblong, rarely curved, 2-6 x 8-16 µm. On PDA the colony has white, woolly mycelium with concentric haloes that tend to grow darker.
A. quercina (anamorph Discula quercina)
A. quercina produces dark, almost brown perithecia, 330-450 x 580-850 µm, on senescent and dead leaves. The asci are evanescent and measure 8-15 x 40-55 µm. The ascospores are two-celled, hyaline, tending to pale yellow, and measure 3-7 x 13-22 µm. On leaves, the acervula of D. quercina are round or elliptical, with their diameter frequently exceeding 250 µm. The conidia are one-celled, hyaline, ellipsoid, 3-6 x 8-15 µm. Colonies are a very light brown; the mycelium is sunken with hyaline, septate hyphae.
The acervula of D. destructiva are dark, globose, subcuticular, with a diameter of 30-135 µm on the leaves and 90-340 µm on the twigs. The conidia are hyaline, one-celled, elliptical or spindle-shaped, and measure 2.5-3.5 x 7-12 µm on the leaves and 2.5-4 x 6-10 µm on the twigs. Colonies on PDA or malt agar are granular, white at first, turning darker with age.
DistributionTop of page Detailed information on the distribution of Discula and some other genera in Italy may be found in Anselmi et al. (2002).
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|
|Azerbaijan||Present||Guseinov, 1974; Guseinov, 1975|
|Japan||Present||Present based on regional distribution.|
|-Honshu||Present||Sahashi et al., 1999; Sahashi et al., 2000|
|Korea, Republic of||Present||Lee et al., 2013|
|Canada||Present||Present based on regional distribution.|
|-New Brunswick||Present||Canadian Forest Service, 2004|
|Mexico||Present||Alvarado et al., 2007; Alvarado et al., 2007|
|USA||Present||Present based on regional distribution.|
|-California||Present||Hecht and Poinar Parmeter, 1986; Sinclair et al., 1987; Raabe, 1990|
|-Connecticut||Present||Connecticut Agricultural Experiment Station, 2004|
|-Iowa||Present||Iowa State University Plant Disease Clinic, 2004|
|-Mississippi||Present||Parris and Byrd, 1962|
|-Ohio||Present||Wilson and Carroll, 1994|
|-Oregon||Present||Toti et al., 1992; Wilson and Carroll, 1994|
|-Washington||Present||Washington State University Cooperative Extension,|
|-Wisconsin||Present||Ambuel et al., 1977|
|Austria||Present||Phillips and Burdekin, 1992; Toti et al., 1992; Halmschlager et al., 1993|
|Belgium||Present||Antheunis & de Temmerman, 1983; Phillips and Burdekin, 1992|
|Czech Republic||Present||Prihoda, 1982; Procházková, 1995|
|Denmark||Present||Phillips and Burdekin, 1992|
|France||Widespread||Morelet, 1989; Phillips and Burdekin, 1992; Toti et al., 1992|
|Germany||Widespread||Butin & Kehr, 1997; Sutton, 1980; Phillips and Burdekin, 1992|
|Italy||Widespread||Mesturino and Mugnai, 1986; Ragazzi et al., 1995; Anselmi, 2001|
|Netherlands||Present||Phillips and Burdekin, 1992|
|Norway||Present||Fluckiger et al., 1999|
|Poland||Present||Kowalski and Kehr, 1992|
|Romania||Present||Sutton, 1980; Marcu et al., 1997|
|Russian Federation||Present||Present based on regional distribution.|
|-Central Russia||Present||Guseinov, 1975|
|-Northern Russia||Present||Treigiene, 1997|
|-Western Siberia||Present||Shirnina, 1997|
|Slovakia||Present||Ivanová and Bernadovicová, 2006|
|Slovenia||Present||Milevoj and Kravanja, 1999|
|Switzerland||Widespread||Toti et al., 1992; Toti et al., 1993; Viret and Petrini, 1994|
|UK||Present||Moore, 1959; Sutton, 1980; Phillips and Burdekin, 1992; Toti et al., 1992|
|-New South Wales||Present||Sutton, 1980|
|New Zealand||Present||Sutton, 1980|
Risk of IntroductionTop of page A. errabunda and A. quercina are not subject to quarantine regulations because they are widespread. It would nevertheless be desirable to have such regulations for countries where the two pathogens do not occur, but where there are large populations of beech and oak.
HabitatTop of page The many associations between oaks and other trees in mixed stands have been described for central Europe by Mayer (1984) and Ellenberg (1988). In simple terms the prototype of such associations can be taken to be the mixed stand of oak and hornbeam (Carpinus betulus), in which Quercus petraea and/or Q. robur grow mixed with hornbeam in mesophile conditions on fertile hilly terrain. From these associations other important groups branch out, growing in different ecological conditions. On the more arid side there is the oak-birch association, with a high proportion of Q. petraea, though this species thrives less here. Where the soil is fresher and richer in nitrogen, the predominant association becomes one of maple and ash, with Q. petraea and/or Q. robur at a strong competitive disadvantage. On very humid soils the alder-elm combination comes to the fore and Q. robur can easily renew itself only if it is periodically flooded with alluvial silt (Perrin, 1954).
For the southern Mediterranean it is difficult to give a general picture of oak groves in a few words because of their diversity. In Italy, a reference oak stand can be taken to be one in which Q. pubescens and Q. cerris are mixed with hop hornbeam (Ostrya carpinifolia), which here replaces Carpinus betulus, though hornbeam remains the main competitor of the oaks. In more arid areas, the hop hornbeam becomes less successful and the only formations that remain are skimpy stands of Q. petraea and flowering ash (Fraxinus ornus), with an undergrowth of herbaceous species of dry brome grass (Ubaldi et al., 1984).
In more acidic soils the most successful stands are those containing Q. cerris, but also Q. petraea and chestnut (Oberdorfer and Hofmann, 1967). The latter species are replaced further south in Italy by Q. cerris with various species of heather (Erica) and Coronilla emerus, which are more thermophilous (Giordano and Schirone, 1989).
In hilly terrain in Italy it is not uncommon to see stands of Q. pubescens and Q. cerris with smaller admixtures of hop hornbeam. In the Po Valley there are plain-growing oak groves with Q. robur and Carpinus betulus, or with Q. robur, Q. cerris and Carpinus betulus (Agostini, 1965; Paiero, 1965). In coastal areas of central and southern Italy, the remaining plain-growing stands consist of Q. robur, Q. cerris, Q. frainetto (only in the south) as well as cork oak and Q. ilex (Corti, 1955; Padula, 1985).
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 The Apiognomonia complex includes numerous anamorph forms (apart from Discula umbrinella and D. quercina, which are dealt with in this datasheet), among which D. destructiva sp. nov. deserves a special mention for its wide distribution and pathogenicity. This species causes serious damage on Cornus florida and C. nuttallii in Canada and the USA (Redlin, 1991).
Information on the host range can be found in Farr et al. (1989), Kowalski and Kehr (1992), Phillips and Burdekin (1992), Toti et al. (1993), Wilson and Carroll (1994), Sahashi et al. (2000) and Anselmi et al. (2002).
An undefined species of Discula has been found in the leaves of Fagus crenata in Japan (Sahashi et al., 2000).
Host Plants and Other Plants AffectedTop of page
|Castanea sativa (chestnut)||Fagaceae||Other|
|Celtis (nettle tree)||Ulmaceae||Other|
|Fagus crenata (Japanese beech)||Fagaceae||Other|
|Fagus sylvatica (common beech)||Fagaceae||Main|
|Quercus alba (white oak)||Fagaceae||Main|
|Quercus aliena (oriental white oak)||Fagaceae||Other|
|Quercus bicolor (swamp white oak)||Fagaceae||Other|
|Quercus cerris (European Turkey oak)||Fagaceae||Main|
|Quercus coccinea (scarlet oak)||Fagaceae||Other|
|Quercus falcata (red oak)||Fagaceae||Main|
|Quercus frainetto (Hungarian oak)||Fagaceae||Other|
|Quercus garryana (Garry oak)||Fagaceae||Main|
|Quercus laurifolia (Laurel oak)||Fagaceae||Main|
|Quercus macrocarpa (mossy-cup oak)||Fagaceae||Other|
|Quercus nigra (water oak)||Fagaceae||Main|
|Quercus palustris (pin oak)||Fagaceae||Other|
|Quercus petraea (durmast oak)||Fagaceae||Main|
|Quercus phellos (Willow oak)||Fagaceae||Main|
|Quercus pubescens (downy oak)||Fagaceae||Main|
|Quercus robur (common oak)||Fagaceae||Main|
|Quercus rubra (northern red oak)||Fagaceae||Other|
|Quercus shumardii (spotted oak)||Fagaceae||Other|
|Quercus suber (cork oak)||Fagaceae||Main|
|Quercus trojana (Macedonian oak)||Fagaceae||Other|
|Quercus velutina (black oak)||Fagaceae||Other|
|Quercus virginiana (Live oak)||Fagaceae||Other|
|Tilia cordata (small-leaf lime)||Tiliaceae||Other|
Growth StagesTop of page Seedling stage, Vegetative growing stage
SymptomsTop of page A. errabunda sensu stricto (anamorph Discula umbrinella) causes leaf spot and shoot dieback on Fagus sylvatica. On the upper leaf surface, the leaf spots are brown and roundish or irregular in outline, whereas on the lower leaf they are olive-green. In summer and autumn, many roundish pustules (the acervula of the anamorph, D. umbrinella) form on the spots. The perithecia of the teleomorph A. errabunda form on senescent leaves or leaves that have fallen to the ground.
A. quercina (anamorph Discula quercina) essentially causes leaf spot and shoot dieback although symptoms vary depending on the oak species infected, the vegetative stage of the tree at the time of infection, the tree organ affected, and the climate.
In Mediterranean countries the first symptom that appears on the leaves shortly after the annual resumption of growth is small, brown spots. These spots increase in size and become reddish-brown, irregular in outline, and bounded by the leaf veins (angular spots); when more nearly round these spots measure 12-22 mm in diameter. The spots may become confluent and cover the entire leaf surface between the veins. However, a part of the leaf blade remains green. When the spots cover 70-80% of the leaf blade, the leaf withers, becomes papery and hazel-brown, and the leaf blade becomes twisted. The entire expanding foliage in the crown is often destroyed.
Older leaves are also affected, but symptoms remain limited to brown spots, which spread but do not occupy the entire leaf area.
The infection often passes through the leaf stalk to the shoot, although the shoot can also be infected directly. If the infection completely surrounds the shoot, it dies. The infection also proceeds to the twigs, on which a canker forms, 8-10 cm long and slightly sunk into the wood. Twigs so infected will die (shoot dieback). The cones are also often infected, becoming covered with small, black specks.
Repeated infections over the years, always entailing the death of leaves, shoots and twigs, gradually cause very extensive crown thinning and ultimately the death of the tree. The perithecia of the teleomorph A. quercina form on the leaves, whereas the acervula of the anamorph D. quercina differentiate on the cankers.
The symptoms as they appear in European countries are described, albeit in a general way, by Grove (1937), who states that in more than usually rainy years, especially in the spring and summer, the incidence of the disease increases as it did in England, UK, in the summer of 1980.
In North America, symptoms vary on the many susceptible oak species. On Quercus alba, which is probably the most sensitive to this infection, symptoms manifest themselves in a number of ways: rapidly spreading blight of leaves and shoots, with browning of young leaves during the growth period; a large and irregularly shaped portion of the leaf blade dies off, and the leaf blade becomes twisted, but part of the leaf area remains green; and small necrotic spots form on the adult leaves.
On Quercus phellos and Q. laurifolia the spots are often very numerous, small, brown or black, and surrounded by a halo of green tissue. On Q. nigra and Q. falcata the spots are larger and often surrounded by a yellow ring. When infection is severe the apical shoot dies and folds in upon itself. On Q. palustris and Q. shumardi the spots on the upper leaf blade are round, reddish-brown, varying in size and surrounded by a pale-green halo, whereas on the lower leaf blade they are often concave. On Q. virginiana, which is not a very sensitive species, leaf spots also form, but at a very low density, 6-10 per square inch compared with 100-150 spots per square inch on more sensitive species. On Q. durandii the density of spots is even lower than that on Q. virginiana.
Further details can be found in Grove (1937), Parris and Byrd (1962), Morelet (1989) and Ragazzi et al. (1999c).
List of Symptoms/SignsTop of page
|Fruit / abnormal shape|
|Fruit / lesions: black or brown|
|Fruit / reduced size|
|Growing point / mycelium present|
|Leaves / abnormal colours|
|Leaves / abnormal forms|
|Leaves / abnormal leaf fall|
|Leaves / leaves rolled or folded|
|Leaves / necrotic areas|
|Stems / canker on woody stem|
|Stems / dieback|
|Stems / discoloration of bark|
|Stems / distortion|
|Stems / gummosis or resinosis|
|Stems / necrosis|
|Stems / stunting or rosetting|
|Whole plant / discoloration|
|Whole plant / early senescence|
|Whole plant / plant dead; dieback|
|Whole plant / seedling blight|
Biology and EcologyTop of page The biological and ecological characters given in this datasheet refer to A. errabunda sensu stricto and to A. quercina with their respective anamorphs, Discula umbrinella and Discula quercina.
A. errabunda and A. quercina are most easily found on Fagus sylvatica and Quercus spp., respectively; however, they have a vast host range to cover their geographic range. In North America they are found from the western provinces of Canada down along the coast to the Gulf of Mexico. In Europe they occur from central and southern Russia across north-eastern Europe, all the way to the countries of the Mediterranean. Such an extensive geographic range, with so many different climates, shows the adaptability of these fungi, though it is possible that ecotypes have differentiated within the species (A. errabunda and A. quercina) that are adapted to the particular climates of the two continents.
This is also seen in Italy, where A. quercina occurs in many different parts of the country, from areas with a semi-arid climate (e.g. Sicily, 37-38° N, 13-15° E) to those with a continental climate (e.g. Tuscany, 43-44° N, 10-12° E) (Anselmi et al., 2002).
In winter A. quercina survives mainly through its anamorph form (acervula) that differentiates on cankers along the branches and twigs. The teleomorph form (perithecia on dead leaves on the ground) ensures survival in more temperate areas.
The primary infection, in all areas, appears after the winter season when the temperature rises in the spring (18-20°C is sufficient for perithecia and acervula to mature). The ascospores and conidia then begin to infect the leaves, buds, shoots and twigs. The teleomorph Apiognomonia usually colonizes the leaves, buds and shoots, whereas the anamorph Discula colonizes not only the leaves and buds but also the twigs (bark and wood) (Ragazzi et al., 1999a). The ascospores and conidia are dispersed by wind, water and seeds, and possibly by insects as well (Petrini, 1991). Under test conditions, ascospores and conidia caused high levels of infection with the following regime parameters: day/night 16/8 h, 22/18°C, 40/65% RH and 20,000/0 lux (Ragazzi et al., 1999b).
The literature reveals that, on the whole, species of the Apiognomonia complex are adapted to temperatures ranging from 3 to 30°C (Neely and Himelick, 1967; Shishkina, 1969; Sinclair et al., 1987; Smith et al., 1988; Wilson and Carroll, 1994). The percent infected leaf area increases as the inoculum concentration increases from 100 to 100,000 conidia/ml, but then decreases at 1,000,000 conidia/ml. An inoculum concentration of 100,000 is therefore optimal for producing extensive leaf necrosis (Ragazzi et al., 1999a). Under experimental conditions of day/night 12/12 h, 25/15°C, 50/75% RH and 25,000/0 lux, conidia production gradually increased until it peaked at 23,000 conidia/m³ air, 14 days after sporulation. Conidia production is greatest between 21.00 h and 06.00 h (Ragazzi et al., 1999a).
The frequency of occurrence and extent of colonization are greater at lower altitudes and diminish as the altitude increases (Toti et al., 1993). Secondary infections are caused mainly by conidia that have differentiated in the canker lesions, produced by the primary inoculum, on the branches and twigs. Infected trees produce infected cones. The inoculum (conidia + ascospores) released from a mixed stand of Quercus cerris and Q. pubescens, averaging 15 m tall and growing at an altitude of 400 m a.s.l., decreases with distance from the inoculum source, so that 10 days after the start of sporulation, the number of propagules trapped in a spore trap increased from 49,300 propagules/m³ air at 10 m from the source to 14,200 propagules/m³ at 1000 m. After 40 days, the corresponding figures were 16,200 propagules/m³ at 10 m and 2400 propagules/m³ at 1000 m (Ragazzi, Università degli Studi di Firenze, Italy, unpublished data).
Individual species of the Apiognomonia complex are invasive in stands of Quercus spp. mixed with Fraxinus spp., Alnus spp., Castanea sativa, Fagus sylvatica, Corylus spp. and Acer spp, depending on location. In the USA, it is the pure oak stands that are most seriously affected. In both Europe and the USA, infection level is related to rainfall, increasing when rainfall is higher in the early part of the growing season (March-April in Europe and March-April-May in the USA). The incidence of A. errabunda on F. sylvatica increased on permanent observation plots in Alpine sites in Switzerland after the application of nitrogen fertilizer (Fluckiger et al., 1999).
A. errabunda sensu stricto is frequently found in association with Aureobasidium pullulans, Diplodina cf. microsperma, Epicoccum purpurescens, Microsphaeropsis sp., Phialophora sp., Phoma sp., Phomopsis sp. and Sporormiella intermedia.
A. quercina is associated with Alternaria alternata, Biscogniauxia mediterranea, Cladosporium cladosporioides, Cytospora sp., Diplodia mutila, Epicoccum spp., Monochaetia sp., Phoma cava, Phomopsis quercina, Trichoderma spp., Tubakia dryina and Ulocladium sp.
Notes on Natural EnemiesTop of page There are currently no known natural enemies of A. errabunda and A. quercina. However, it should be noted that on oak in Italy, the tree organs from which A. quercina is easily isolated are those that do not harbour any mycetes known to be biological antagonists of these agents. A. quercina is not found, or is much less common, at sites where potential antagonist species such as Acremonium, Aureobasidion, Paecilomyces, Ramichloridium and Trichoderma are frequent (Ragazzi, Università degli Studi di Firenze, Italy, unpublished data).
Means of Movement and DispersalTop of page Conidia and ascospores of A. errabunda and A. quercina are dispersed by air currents. Fallen leaf fragments bearing perithecia are waterborne, being dispersed on the ground by water runoff. Infected seedlings can also spread these pathogens. It is thought, though so far without experimental proof, that many insects that are vectors of fungi associated with declining oaks (some suspected, some established; Tiberi et al., 2002) are also vectors of A. errabunda and A. quercina.
Seedborne AspectsTop of page The acorn becomes infected, but it is not known where on the fruit the fungus is located, whether it contaminates the seeds, how long the fungus remains viable on the seed, or whether it can transmit the disease in this way.
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|
|Fruits (inc. pods)||hyphae||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Leaves||fruiting bodies; spores||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Seedlings/Micropropagated plants||hyphae||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Stems (above ground)/Shoots/Trunks/Branches||fruiting bodies; spores||Yes||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Plant parts not known to carry the pest in trade/transport|
|Growing medium accompanying plants|
|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|
|Wood Packaging not known to carry the pest in trade/transport|
|Processed or treated wood|
Impact SummaryTop of page
ImpactTop of page The economic impact of A. errabunda and A. quercina is hard to assess. Many oak species that become infected have no specific uses and there is no market for them. Other species are used for timber but precise data on losses to these trees caused by infections from pathogenic agents are not accessible, if only because the infections they cause do not attack the trunk and therefore seem economically insignificant. It should nevertheless be remembered that repeated infections, with their defoliations in consecutive years, lead to a lower growth rate and, hence, a lower annual increment. For the owner of a stand so affected this certainly means an economic loss, which is aggravated by the cost of removing the dead wood.
In urban and periurban parks the removal of leaves fallen in consequence of infection by species of Apiognomonia is without doubt a burden on the city administration.
Economic ImpactTop of page The economic impact of A. errabunda and A. quercina is hard to assess. Many oak species that become infected have no specific uses and there is no market for them. Other species are used for timber but precise data on losses to these trees caused by infections from pathogenic agents are not accessible, if only because the infections they cause do not attack the trunk and therefore seem economically insignificant. It should nevertheless be remembered that repeated infections, with their defoliations in consecutive years, lead to a lower growth rate and, hence, a lower annual increment. For the owner of a stand so affected this certainly means an economic loss, which is aggravated by the cost of removing the dead wood.
In urban and periurban parks the removal of leaves fallen in consequence of infection by species of Apiognomonia is without doubt a burden on the city administration.
Environmental ImpactTop of page Fagus sylvatica has a wide range within which it grows in pure stands or in mixed stands with Abies sp., at higher altitudes, and Quercus spp., at lower altitudes, as well as with other tree species of less importance.
In all of these stands, both A. errabunda and, where there are oaks, A. quercina occur endemically. The incidence of these fungi is higher when the season is favourable and this can cause heavy defoliation and death of the apical twigs.
The environmental impact is nevertheless thought to be only modest: the tree, beech or oak, may die after repeated attacks in successive years, but during this long period natural renewal may allow the stand to retain its original forest cover and tree density. It is difficult to isolate the effect of A. quercina alone from the whole context of oak decline, though it is certainly one of a number of pathogenic agents that cause the death of oak trees. As such, the environmental impact of these two pathogens in particular cannot be assessed.
DiagnosisTop of page A. errabunda (anamorph Discula umbrinella).
D. umbrinella can be isolated from leaves of Quercus alba and Q. rubra by the following method:
The fungus was isolated from leaf discs using the method of Wilson and Carroll (1994) with modifications in the solution concentrations and in the times and number of rinses during the sterilization procedure. A 6-mm-hole punch sterilized in 95% ethanol was used to remove six leaf discs located equidistantly along the midvein of each leaf. Each disc was placed on the sterile filter paper of a large vacuum filtration apparatus. The apparatus consisted of a plastic container (27.5 x 40 x 14.6 cm), an aluminium frame with an open aluminium lattice (33.1 x 46.2 x 2.8 cm), a sheet of porous Teflon filter (39.7 x 26.9 cm) (Reed Plastics, Rockville, Maryland, USA), and a sheet of sterile filter paper (39 x 26 cm) (Whatman, 3MM). The apparatus was attached in tandem to a vacuum flask and vacuum line. The entire apparatus was assembled within a laminar flow hood to maintain sterility throughout the isolation procedure. The sterile Whatman paper was placed on the Teflon surface of the vacuum filtration unit, premoistened with sterile distilled water, which was allowed to drag by gravity. The leaf discs were sterilized on the apparatus with 95% ethanol for 50 s, 70% ethanol for 50 s, 0.5% sodium hypochlorite with Tween 80 (0.2 ml/100 ml) for 2 min, and were then rinsed three times with sterile distilled water. Sterilization fluids were applied with squeeze bottles or pasteur pipettes. Excess liquid was vacuumed away from the leaf discs, which were then transferred to 24-well tissue-culture dishes containing 1 ml per well of oak leaf agar (OLA) consisting of 1.0 g dried and ground white oak leaves, 20 g Difco agar, 1000 ml distilled water. Prior to preparation of the OLA medium, leaves were collected, dried at 100°C for 15 min, shredded in a Waring blender for 1 min, then ground in a coffee grinder for 30 s. Dishes were incubated at 24°C under 12 h fluorescent lighting per day to induce conidiomata production.
Apiognomonia quercina (anamorph Discula quercina)
The isolation of D. quercina followed Ragazzi et al. (1999c). Two bark fragments of 3 cm² each were excised from Quercus cerris twigs, one from near the tip and one from near the base of the twigs, and two wood fragments were similarly removed, one from the base and one from the tip of the same twigs. Two fragments of embryo tissue and two of bud scale were also removed from each of two basal buds and two apical buds per twig. All fragments were sterilized by immersion in 3% sodium hypochlorite for 4 min followed by immersion in 60% ethanol for 2 min. The fragments were placed in Petri dishes, 9 cm in diameter, containing 20 ml PDA, plus 30 p.p.m. streptomycin to suppress bacterial growth. Incubation was in a controlled environment chamber at 22±2°C, 70% RH, with constant lighting at 15,000 lux.
Colony differentiation started after 4-5 days of incubation. Colonies were a very light brown, almost yellow, the mycelium was immersed, with hyaline, septate and branched hyphae. Growth, on oak leaf agar (OLA) at 25°C, was slow, reaching a diameter of ca 6.5 cm after 8 days. Acervula, 150 µm in diameter, epidermic, separate or confluent, light brown, smooth-walled and with an irregular dehiscence were produced after about 5 days on impoverished PDA (6 g/l potato dextrose broth (PDB) + 20 g/l agar). Conidiophores were 13-3-5 x 21 µm, hyaline, septate, single or branched (but only at the base), straight or slightly curved, elongated towards the tip. Conidia were 3.5-5 x 9-15 µm, hyaline, oblong, ellipsoid, not septate, with a smooth, thin wall, a blunt tip and the base more or less truncated.
PDA was certainly an excellent medium for the isolation of A. quercina, but for colony growth the best medium was OLA, pH 6.6, prepared by boiling 30 g oak leaves in 350 ml water for 30 min and adding 8 g agar (Difco). The best growth conditions in a controlled environment were: 30 W fluorescent lighting, a 12-h day with day/night temperatures 24/18°C; 50/75% RH, light intensity 15,000/0 lux. Under such conditions the colony differentiated in 3 days and reached a diameter of 9 cm in 6 days (Ragazzi et al., 2002). Other authors have carried out similar experiments, the most important of which have been Neely and Himelick (1967) and Wilson and Carroll (1994).
Neely and Himelick stated that isolates of Gnomonia quercina from white oak were studied on 10 agar media (Difco) with added bean pod, cabbage infusion, corn meal, lima bean, malt extract, potato dextrose, prune and Sabourand-dextrose. Growth rates were determined at temperatures ranging from 3 to 30°C.
Conidia production was greatest on bean pod, malt extract and PDA. No isolate produced many conidia on cabbage infusion, corn meal or Sabourand-dextrose. All isolates tended to form zonation rings on many of the media. The aerial mycelium of all isolates was white tending to pale grey on all media. Zonation rings varied in colour from grey to black. On PDA all isolates produced a dark-yellow pigment that spread throughout the medium after incubation for 4-6 weeks at 21°C. Colonies reached their greatest diameter size (7-9 cm) on prune, Sabourand-dextrose, corn meal and PDA. The optimal temperature for growth was 18°C.
Wilson and Carroll (1994) isolated D. quercina from various tissues of Quercus garryana. Surface sterilization of tree tissues was carried out as described in Wilson and Carroll (1994):
- Acorn shells (60 s in 95% ethanol, 60 s in 70% ethanol, 5 min in 33% NaOCl, 5 washes in sterile, distilled water)
- Naked seeds (shelled acorns) (30 s in 70% ethanol, 10 min in 10% NaOCl with 1% SDS, 5 washes in sterile, distilled water)
- Twigs, bark, wood (60 s in 95% ethanol, 2 min in 70% ethanol, 8 min in 33% NaOCl, 5 washes in sterile, distilled water)
- Leaves (50 s in 95% ethanol, 60 s in 70% ethanol, 5 min in 33% NaOCl, 4 washes in sterile, distilled water)
- Prebud burst and young postbud burst leaves (25 s in 95% ethanol, 30 s in 70% ethanol, 2 min in 33% NaOCl, 4 washes in sterile, distilled water).
NaOCl: dilutions (by volume) from a 5% sodium hypochlorite household bleach stock solution.
SDS: sodium dodecyl sulfate is a detergent used as a wetting agent.
Isolation was attempted from three organs:
Shelled acorns: the shell was removed, leaving only the cotyledons, which were attached at the embryo region. A single piece of each acorn seed (about a quarter of the total acorn) was surface sterilized. The pieces were placed in 22-mm-diam. sterile test tubes with 2 ml sterile water, and incubated under ambient laboratory conditions for 6 weeks, then scored for D. quercina.
Twigs: one 12-cm length of 1-2- and 3-year-old twigs was cut from trees. These were put on ice, taken back to the laboratory and immediately surface sterilized. The terminal 1 cm of twig length was cut off, then the remaining 10-cm length was cut into four equal sized parts. All 40 1-year-old segments, and 10 out of the 40 segments of the 2- and 3-year age classes of twigs were placed onto PDA and left to incubate under ambient laboratory conditions, then scored for the presence of the endophyte. All the remaining 2- and 3-year-old twig pieces had the bark removed from the wood. Bark of 2- and 3-year-old twigs can be prized off the wood with a sterile razor blade. The bark and wood were both separately placed onto PDA, incubated for 3 weeks under ambient laboratory conditions, then scored for the presence of the endophyte.
Increment cores of the tree trunk: two core samples per tree were taken at breast height from five trees. As oak trees are difficult to core with increment corers, the cores only went 10 cm into the tree trunk. To prevent contamination of the core sample with bark fungi, a small section of bark was removed from the tree around the point where the increment corer was inserted. The increment corer was sterilized in the field by flaming with ethanol. The cores were immediately placed in sterile glass tubes on ice, taken to the laboratory, surface-sterilized, broken in half, placed on PDA, incubated, and scored for D. quercina.
Studies to establish the cultural characteristics of A. quercina can be found in Stoneman (1898), Edgerton (1908), Westerdijk and van Luijk (1920) and Schuldt (1955).
Detection and InspectionTop of page A. errabunda and A. quercina cause a type of leaf spot that is diagnostic of these fungi: no other known biotic or abiotic agents produce similar spots. The presence of these two mycetes is practically confirmed when the spots are accompanied by cankers on the twigs, caused by the anamorphs Discula umbrinella and D. quercina, respectively. Absolute certainty can only be attained by isolating the fungi.
Toti et al. (1993) isolated D. umbrinella from the buds, twigs and leaves of Fagus sylvatica as follows: twigs from 10 randomly selected F. sylvatica trees were collected in March 1991 in Switzerland. One branch, 1-1.5 cm in diameter and carrying at least 30-40 buds, was excised from each tree. Five buds, along with the branch segment bearing the buds, were taken from each branch and surface sterilized by sequential immersion in 75% ethanol (1 min), NaOCl with 3-5% available chlorine (5 min), and 75% ethanol (30 sec). Each bud was cut longitudinally and the outer scales removed. The young, rolled-up leaf was cut into two parts, and the first 3-cm length of each piece of contiguous twig was sectioned longitudinally into two pieces. The twig pieces directly underneath the buds were contiguous pieces, as distinct from the distal pieces, which were separated from the buds by the contiguous twig pieces. Bud scales, rolled up leaf pieces, and twig pieces were placed in that order on 2% MEA (2% agar, 1% malt extract, Difco) supplemented with 50 p.p.m. terramycin to suppress bacterial growth. All plates were incubated at 20±2°C for up to 3 weeks, depending on the growth rate of the fungi. Isolation was by transfer of mycelium to MEA slants. Near-UV light (Philips TL 40W/05) was used to induce sporulation. Other detection methods are described in 'Diagnostic Methods'.
In addition to traditional detection methods, a number of biomolecular techniques are now widely available in plant pathology. Haemmerli et al. (1992) identified and compared isolates of D. umbrinella in F. sylvatica, Castanea sativa, Quercus petraea and Q. rubra using the following protocol:
All strains of D. umbrinella were isolated from symptomless leaf tissues of F. sylvatica, C. sativa, Q. petraea and Q. rubra as described by Sieber and Hugentobler (1987); all isolates were mycelial isolates. Cultures were maintained on 2% MEA slants (2% malt extract, 2% agar, both Difco ) at 4°C.
Isolates were grown for 10 days at 20°C in shake culture containing 50 ml of V8 juice. The mycelium was removed by centrifugation (10 min at 3000 x g), lyophilized and ground to a fine powder. Mycelial DNA (30-40 mg) was extracted and purified by a total DNA CTAB mini-prep extraction method (Zolan and Pukkila, 1986).
PCR amplification conditions
Amplification reactions were carried out in volumes of 25 µl containing 10 mM Tris-HCl, pH 8.3; 50 mM KCl; 2 mM MgCl2 (for primers P1, P2, P14, OPE2, OPE4, OPE7, OPE12, OPE15); 2.5 mM MgCl2 (primers P11, OPE3, OPE5, OPE6, OPE11, OPE16, OPE20); 3 mM MgCl2 (P4, OPE1); 100 µM each of dATP, dCTP, dGTP and TTP (Boehringer Mannheim); 0.2 M primer; 25 ng of DNA; and 1 U of Taq DNA polymerase (Boehringer Mannheim). Amplification was performed in a Perkin Elmer Cetus Gene Amp PCR system 9600 programmed for two cycles of 30 sec at 94°C, 30 s at 36°C, 120 sec at 72°C, 36 cycles of 20 s at 94°C, 15 s at 36°C, 15 s at 45°C, 90 s at 72°C, followed by 10 min at 72°C. Reaction products were resolved by electrophoresis (4 V/cm) in a 1.5% agarose gel, run in 1 x TPE for 4 h and stained with ethidium bromide.
A total of 17 random primers, each with a length of 10 nucleotides and a minimum GC content of 50% were used. The primers were supplied by Operon Technologies Inc., Alameda, California, USA, and some were synthesized by MicroSynth AG, Windisch, Switzerland.
Hybridization and autoradiography
Three RAPD marker products: no. 211, a 2100-bp product of isolate 11 by primer P2; no. 214, a 1170-bp product of isolate 2 by primer P14; and no. 1814, a 1000-bp product of isolate 18 by primer P14, were transferred from the agarose gel to reaction tubes using autoclaved toothpicks and amplified under the conditions described above. Twenty-five nanograms of reamplified DNA was labelled with 50 µCi (a-32P)dCTP (3000 Ci/mmole; Amersham International; random primed DNA labelling kit, Boehringer Mannheim) and used as hybridization probes on Southern blots. EcoRI-digested total genomic DNA was separated by horizontal agarose electrophoresis for 15 h and blotted to Hybond N+ membranes (Amersham) (Southern, 1975). Prehybridization and hybridization reactions were carried out in plastic bags at 65°C with 5 x SSPE, 5 x Denhardt's solution, 1% SDS, 0.5 mg of herring sperm DNA. Hybridizations were incubated at 65°C for 14 h.
The occurrence of each amplification product in the gel was scored as 1, non-occurrence as 0. The resulting matrix was used to compute indices of between-isolate distance using the PCT option offered by the software package SYSTAT 5.1 (Wilkinson, 1989). This matrix computes the percentage of value comparisons resulting in disagreements in two column or row profiles, and is particularly useful for categorical or nominal scales as it does not apply any weighting to the matrix. The dendrogram was constructed by UPGMA-clustering (Sneath and Sokal, 1973).
Prevention and ControlTop of page
Any methods of control must be applied to trees in the nursery or to young trees in the field. On adult trees, the general decline caused by A. errabunda on beech and A. quercina on oak, involves other organisms and is therefore difficult to control.
In the nursery and arboretum it is good practice to gather up the leaves, twigs and branches that fall to the ground, to keep the trees in good condition by proper watering and fertilizing, and to ensure good ventilation.
Although it remains doubtful whether A. quercina is transmitted by seed, it is nevertheless strongly advisable to use healthy seed (checked to be free of the mycete) or seed from healthy trees. Any beech or oak trees that become infected in adult stands must be eliminated.
Chemical treatments are rare. Only in cases where anthracnose has defoliated oak trees for 3-5 years in succession is thiophanate-methyl applied to start vegetative growth, with repeated treatment after 7-10 days. Additional spraying may be carried out when the growing season is cool and humid (see http://www.entomology-umn.edu/cues/dx/CB/oanthab.htm, the site of the Center for Urban Ecology and Sustainability, The University of Minnesota, USA).
One promising means of control is to promote fungi that are biological antagonists against A. quercina and occur on the same organs on the oak tree that yield the disease agent.
ReferencesTop of page
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Distribution MapsTop of page
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