Mycosphaerella dearnessii (brown spot needle blight)
- Summary of Invasiveness
- Taxonomic Tree
- Notes on Taxonomy and Nomenclature
- Distribution Table
- Risk of Introduction
- Hosts/Species Affected
- Host Plants and Other Plants Affected
- Growth Stages
- List of Symptoms/Signs
- Biology and Ecology
- Means of Movement and Dispersal
- Plant Trade
- Wood Packaging
- Detection and Inspection
- Similarities to Other Species/Conditions
- Prevention and Control
- Distribution Maps
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PicturesTop of page
IdentityTop of page
Preferred Scientific Name
- Mycosphaerella dearnessii M.E. Barr 1972
Preferred Common Name
- brown spot needle blight
Other Scientific Names
- Cryptosporium acicola Thüm. 1878
- Dothidea acicola (Dearn.) Morelet 1968
- Lecanosticta acicola (Thüm.) Syd. 1924
- Lecanosticta pini Syd. 1921
- Oligostroma acicola Dearn. 1926
- Scirrhia acicola (Dearn.) Sigg. 1939
- Septoria acicola (Thüm.) Sacc. 1941
- Systremma acicola (Dearn.) F.A. Wolf & Barbour 1884
International Common Names
- English: brown spot disease; brown spot needle disease; brown spot of pine; brown spot: pine; needle blight of pine; needle blight: brown spot; needle blight: pine; needle disease: brown spot
- Spanish: mancha parda de las aciculas del pino
- French: tache brune des aiguilles du pin
Local Common Names
- Germany: Lecanosticta-Nadelbräune
- SCIRAC (Mycosphaerella dearnessii)
Summary of InvasivenessTop of page Brown-spot needle blight caused by Mycosphaerella dearnessii kills foliage and retards growth of many pine species. The pathogen has a wide host and habitat range and occurs on pines in tropical to temperate zones. The brown-spot fungus mainly attacks pines from Central to North America but also in localities from South America, Asia, Africa and Europe. Long known as a serious disease in longleaf pine in the Gulf States of the USA, the pathogen gained added notoriety in recent decades because of needle browning and defoliation of landscape and Christmas tree plantings of ponderosa and Scots pines in the central plains and Great Lakes regions (Phelps et al., 1978; Sinclair et al., 1987).
The impact of brown-spot disease on longleaf pine results in the reduction of total annual growth by more than 16 million cubic feet (453 thousand cubic metres) of timber. In Christmas tree plantations attack of brown-spot needle blight on Scots pine and other pines results in the loss of thousands of dollars, because of needle drop, making the trees unmerchantable (Phelps et al., 1978). M. dearnessii has spread rapidly and has a very wide distribution. Plant disease reports from the EPPO region also suggest an increase in the prevalence in that part of the world.
Taxonomic TreeTop of page
- Domain: Eukaryota
- Kingdom: Fungi
- Phylum: Ascomycota
- Subphylum: Pezizomycotina
- Class: Dothideomycetes
- Subclass: Dothideomycetidae
- Order: Capnodiales
- Family: Mycosphaerellaceae
- Genus: Mycosphaerella
- Species: Mycosphaerella dearnessii
Notes on Taxonomy and NomenclatureTop of page The brown spot disease has been known in the southern USA since the nineteenth century (Hedgcock, 1929). The causal agent in the form of the anamorphic state was first described by De Thümen (1878) as Cryptosporium acicola. But this genus was characterised by hyaline, aseptate conidia so that Saccardo (1884) moved the species on account of its septate conidia to the genus Septoria. Sydow (Sydow and Petrak, 1922) received Pinus taeda material with needle blight and erected the new genus Lecanosticta. He named the fungus Lecanosticta pini being unaware that there were earlier descriptions. Later (Sydow and Petrak 1924), he recognised the synonymy but considered that the generic concept of Lecanosticta was still valid because of the erumpent stromata and darkly pigmented conidia. The new combination Lecanosticta acicola received the general acceptance of plant pathologists and mycologists (Sutton, 1980; Arx, 1983) for the anamorph of the brown-spot fungus.
The teleomorphic form was first described as Oligostroma acicola by Dearness (1926) in association with Lecanosticta acicola. He suggested a teleomorph-anamorph connection but it was left to Siggers (1939) to prove the connection by cultural tests. He came to the conclusion that the genus Oligostroma was not suitable for the brown-spot fungus and referred the erumpent stromata to the genus Scirrhia by making the new combination Scirrhia acicola. Because of the serious effects on young stands of Pinus palustris, Wolf and Barbour (1941) undertook a detailed study of the brown-spot fungus. They accepted the anamorph disposition in Lecanosticta but considered the teleomorph to be best accommodated in the genus Systremma. Siggers (1944) defended his earlier taxonomic decision as to the correct placement of the teleomorph by research of ascostromatal morphology in Oligostroma, Systremma and Scirrhia. He concluded that the teleomorph of the brown-spot fungus is similar in terms of stromatal development and characters with the genus Scirrhia. He also refuted Wolf and Barbours' observations concerning ascospore pigmentation. Wolf and Wolf (1947) were still critical of Sigger's evidence, believing that he made research on immature stromata and immature ascospores. Probably on the basis of nomenclatural examinations, Morelet (1968) proposed the new combination Dothidea acicola. Barr (1972) later transferred the fungus to the genus Mycosphaerella using locule and ascus development as generic concepts in contrast to the position of the ascocarp, which she considered as a highly variable character. Because the name M. acicola already existed for a different fungus, she established the new name Mycosphaerella dearnessii for the teleomorphic form of the brown-spot fungus. However the most widely used name to identify the brown-spot fungus in both mycological and phytopathological literature is currently, but incorrectly, Scirrhia acicola (Punithalingam and Gibson, 1973; Gibson, 1979; Sutton, 1980; Jewell, 1983).
The taxonomy of the genus Mycosphaerella is complicated, and several competing classification systems have been proposed (Arx, 1983; Sivanesan, 1984; Barr, 1996). Due to the large number of associated anamorphs, Crous and Wingfield (1996) concluded that Mycosphaerella was a polyphyletic assemblage of presumably monophyletic anamorph genera. Barr (1996) agreed and separated species with Dothistroma and Lecanosticta anamorphs into the new genus Eruptio. She decided that these two anamorphs are closely related and differ from other species within Mycosphaerella. But phylogenetic analysis of ITS (internal transcribed sequence) data contradicted these assumptions and suggested that the genus Mycosphaerella is monophyletic (Goodwin et al., 2001).
DescriptionTop of page In the life cycle of the brown-spot fungus only the anamorphic state regularly develops on diseased needles, in contrast to the sexual state, the development of which has only been confirmed in some countries.
The asexual fruiting bodies are dark green conidiomata (acervuli; see Pictures) showing subepidermal development, becoming erumpent with further growth and opening of the needle epidermis and hypodermis by one or two longitudinal slits (see Pictures). Conidiomata are elliptical to elongate, 150-800 µm long and 100-200 µm wide, arranged parallel to the long axis of the needle. However, the range of morphological variability appears to be controlled by host and habitat (Evans, 1984). The basal stroma is composed of thick-walled pseudoparenchymatic cells (textura angularis) and shows an extension from shallow to deep. Sometimes excessive stromatal development results in loculate 'acervuli', often misinterpreted as 'pycnidia'. Conidiophores are hyaline to pale brown, septate, branching below septate, cylindrical and up to 90 µm long by 2.5 µm in diameter, arising in a dense, palisade-like layer from the stroma. Conidia are formed successively from annellidic conidiogenous cells at the apex of the conidiophores (Laut et al., 1966). They are extremely variable in form, subhyaline to dark brown, straight to curved, fusiform to cylindrical, echinulate to verrucose to tuberculate, rounded apex and truncate base, thick walled, 14-50 (av. 32) x 3-5 µm and (1-)2-3(-5) septate (see Pictures). Evans (1984) described significant environmentally related changes in the anamorphic stage based on investigations of different collections. Conidia collected from pines growing in habitats exposed to a high light intensity were larger, more pigmented and ornamented compared with those from upland or cloud forest regions. These variations in fungal morphology could be best regarded as morphotypes or ecotypes (Evans, 1984). Conidia are exuded in an olive mucilaginous mass, sometimes in the form of a wedge-shaped cirrus.
The sexual fruiting bodies (ascomata pseudothecia, ascolucular development) are irregularly dispersed on dead needle tissue. Ascomata scattered, linear, innate, subepidermal development, becoming strongly erumpent and splitting the epidermis when mature, black, rarely uniloculate, most multiloculate (up to 18 locules), composed of thick-walled pseudoparenchyma, 400-1200 x 120-250 µm. Locules usually in rows, globose to flask-shaped, ostiolate, periphysate, 50-70 x 50-80 µm. Asci saccate to cylindrical, bitunicate, 8-spored, hyaline, with a round apex, 25-55 x 6.5-10.5 µm. Ascospores uniseriate or obliquely biseriate, hyaline, slightly unequally oblong to cuneate, typically 4-guttulate, 7.5-13.5 x (2-)3-3.5 µm.
Spermogonia Asteromella-like. Synanamorph are present in uni- or multiloculate stromata. Spermatia subhyaline to pale green, rod-shaped, 2.5 x 1-1.5 µm.
A full description of both anamorphic and teleomorphic states is given by Evans (1984).
In culture, on malt extract agar (2% malt extract, 2% agar agar) colonies slow growing (2.5-3 mm growth/week) at 20°C, stromatic, green-olive to olive-black, producing dark olive conidial slime, agar coloured light yellow by diffusates (see Pictures) (Pehl and Wulf, 2001).
DistributionTop of page Mycosphaerella dearnessii has a wide host and habitat range. Potentially all pine species are hosts. The anamorph is recorded throughout the entire wide global distribution. Compared to this global distribution, the teleomorph is reported less frequently and appears not to play an important role in the disease epidemiology.
See also CABI/EPPO (1998, No. 215).
Distribution TableTop of page
The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.
|Continent/Country/Region||Distribution||Last Reported||Origin||First Reported||Invasive||Reference||Notes|
|China||Restricted distribution||CABI/EPPO, 2010; EPPO, 2014|
|-Fujian||Widespread||CABI/EPPO, 2010; EPPO, 2014|
|-Guangdong||Present||CABI/EPPO, 2010; EPPO, 2014|
|-Guangxi||Present||CABI/EPPO, 2010; EPPO, 2014|
|-Jiangsu||Present||CABI/EPPO, 2010; EPPO, 2014|
|-Jiangxi||Present||CABI/EPPO, 2010; EPPO, 2014|
|-Zhejiang||Present||CABI/EPPO, 2010; EPPO, 2014|
|Georgia (Republic of)||Present||CABI/EPPO, 2010; EPPO, 2014|
|Japan||Restricted distribution||CABI/EPPO, 2010; EPPO, 2014|
|-Honshu||Present||CABI/EPPO, 2010; EPPO, 2014|
|Korea, Republic of||Present||CABI/EPPO, 2010; EPPO, 2014|
|South Africa||Absent, invalid record||CABI/EPPO, 2010; EPPO, 2014|
|Canada||Restricted distribution||Evans, 1984; CABI/EPPO, 2010; EPPO, 2014|
|-Manitoba||Present||Evans, 1984; CABI/EPPO, 2010; EPPO, 2014|
|Mexico||Present||Evans, 1984; CABI/EPPO, 2010; EPPO, 2014|
|USA||Widespread||CABI/EPPO, 2010; EPPO, 2014|
|-Alabama||Present||Invasive||CABI/EPPO, 2010; EPPO, 2014|
|-Arkansas||Present||CABI/EPPO, 2010; EPPO, 2014|
|-Florida||Present||Invasive||CABI/EPPO, 2010; EPPO, 2014|
|-Georgia||Present||Invasive||CABI/EPPO, 2010; EPPO, 2014|
|-Idaho||Present||CABI/EPPO, 2010; EPPO, 2014|
|-Illinois||Present||CABI/EPPO, 2010; EPPO, 2014|
|-Iowa||Present||CABI/EPPO, 2010; EPPO, 2014|
|-Kansas||Present||CABI/EPPO, 2010; EPPO, 2014|
|-Kentucky||Present||CABI/EPPO, 2010; EPPO, 2014|
|-Louisiana||Present||Invasive||CABI/EPPO, 2010; EPPO, 2014|
|-Minnesota||Present||Invasive||CABI/EPPO, 2010; EPPO, 2014|
|-Mississippi||Present||Invasive||CABI/EPPO, 2010; EPPO, 2014|
|-Missouri||Present||CABI/EPPO, 2010; EPPO, 2014|
|-Montana||Present||CABI/EPPO, 2010; EPPO, 2014|
|-New York||Present||CABI/EPPO, 2010; EPPO, 2014|
|-North Carolina||Present||CABI/EPPO, 2010; EPPO, 2014|
|-Ohio||Present||CABI/EPPO, 2010; EPPO, 2014|
|-Oregon||Present||CABI/EPPO, 2010; EPPO, 2014|
|-South Carolina||Present||CABI/EPPO, 2010; EPPO, 2014|
|-Tennessee||Present||CABI/EPPO, 2010; EPPO, 2014|
|-Texas||Present||Invasive||CABI/EPPO, 2010; EPPO, 2014|
|-Virginia||Present||CABI/EPPO, 2010; EPPO, 2014|
|-Wisconsin||Present||Invasive||CABI/EPPO, 2010; EPPO, 2014|
Central America and Caribbean
|Belize||Present||Evans, 1984; CABI/EPPO, 2010; EPPO, 2014|
|Costa Rica||Present||Evans, 1984; CABI/EPPO, 2010; EPPO, 2014|
|Cuba||Present||CABI/EPPO, 2010; EPPO, 2014|
|Guatemala||Present||Evans, 1984; CABI/EPPO, 2010; EPPO, 2014|
|Honduras||Present||Evans, 1984; CABI/EPPO, 2010; EPPO, 2014|
|Jamaica||Present||CABI/EPPO, 2010; EPPO, 2014|
|Nicaragua||Present||Evans, 1984; CABI/EPPO, 2010; EPPO, 2014|
|Colombia||Present||CABI/EPPO, 2010; EPPO, 2014|
|Austria||Restricted distribution||Brandstetter and Cech, 1999; CABI/EPPO, 2010; EPPO, 2014|
|Bulgaria||Absent, unreliable record||EPPO, 2014|
|Croatia||Restricted distribution||1976||CABI/EPPO, 2010; EPPO, 2014|
|Czech Republic||Present, few occurrences||CABI/EPPO, 2010; EPPO, 2014|
|Estonia||Absent, intercepted only||CABI/EPPO, 2010|
|France||Present, few occurrences||1993||Chandelier et al., 1994; CABI/EPPO, 2010; EPPO, 2014|
|-France (mainland)||Present, few occurrences||CABI/EPPO, 2010|
|Germany||Present, few occurrences||Pehl, 1995; CABI/EPPO, 2010; EPPO, 2014|
|Greece||Present||CABI/EPPO, 2010; EPPO, 2014|
|Italy||Restricted distribution||La Porta & Capretti, 2000; CABI/EPPO, 2010; EPPO, 2014|
|-Italy (mainland)||Restricted distribution||CABI/EPPO, 2010|
|Latvia||Present, few occurrences||EPPO, 2014|
|Lithuania||Restricted distribution||EPPO, 2014|
|Macedonia||Present||Evans, 1984; CABI/EPPO, 2010|
|Netherlands||Absent, confirmed by survey||NPPO of the Netherlands, 2013; EPPO, 2014||Based on long-term annual surveys.|
|Russian Federation||Absent, unreliable record||EPPO, 2014|
|-Russian Far East||Absent, unreliable record||EPPO, 2014|
|-Southern Russia||Absent, unreliable record||EPPO, 2014|
|Serbia||Absent, unreliable record||EPPO, 2014|
|Slovenia||Transient: actionable, under eradication||CABI/EPPO, 2010; EPPO, 2014|
|Spain||Absent, unreliable record||EPPO, 2014|
|Switzerland||Present||Holdenrieder and Sieber, 1995; CABI/EPPO, 2010; EPPO, 2014|
|UK||Absent, intercepted only||EPPO, 2014|
|-England and Wales||Absent, intercepted only||EPPO, 2014|
Risk of IntroductionTop of page M. dearnessii is listed as an A2 quarantine pest by EPPO (Smith et al., 1997). The limited distribution in the region and the highly adaptable character of the pathogen with a wide ecological tolerance and host range suggests that it presents a considerable risk for the countries in the region. The fungus is believed to be of American origin. In Europe, the brown-spot fungus has been reported in Bulgaria on Pinus nigra (Kovacevski, 1938; Petrak, 1961) in Spain on P. radiata (Martinez, 1942), in Yugoslavia on P. halepensis (Evans, 1984) in Georgia (ex-USSR) on Pinus sp. (Schischkina and Tzanava, 1967), and recently in France on P. attenuata x radiata, P. radiata, P. taeda, and P. attenuata (Chandelier et al., 1994), in Switzerland on P. mugo and P. uncinata (Holdenrieder and Sieber, 1995), in Southern Germany on P. mugo (Pehl, 1995) and in Austria on P. mugo (Brandstetter and Cech, 1999).
The main phytosanitary risk is the global trade (export/import) with infected plant material. There are import restrictions on M. dearnessii in many countries, but between first infection and the first visible symptoms (latency period) attacked plants look healthy.
Hosts/Species AffectedTop of page All pine species are potential hosts. Brown spot disease affects trees of all sizes but is most damaging on small ones. The pathogen is known as one of the major obstacles to increased production of longleaf pine (Pinus palustris) in the south-eastern USA and also as a major cause of making Scots pine (P. sylvestris) Christmas trees unmarketable in the central states. Artificial infection of Picea glauca was noticed by traces of infection with heavy spore inoculum (Siggers, 1944; Punithalingam and Gibson, 1973; Evans, 1984; Skilling and Sinclair et al., 1987).
Host Plants and Other Plants AffectedTop of page
|Picea glauca (white spruce)||Pinaceae||Other|
|Pinus attenuata (knobcode pine)||Pinaceae||Main|
|Pinus ayacahuite (Mexican white pine)||Pinaceae||Other|
|Pinus banksiana (jack pine)||Pinaceae||Main|
|Pinus caribaea (Caribbean pine)||Pinaceae||Other|
|Pinus contorta (lodgepole pine)||Pinaceae||Main|
|Pinus echinata (shortleaf pine)||Pinaceae||Main|
|Pinus elliottii (slash pine)||Pinaceae||Main|
|Pinus glabra (spruce pine)||Pinaceae||Main|
|Pinus halepensis (Aleppo pine)||Pinaceae||Main|
|Pinus maximinoi (thin-leaf pine)||Pinaceae||Other|
|Pinus monticola (western white pine)||Pinaceae||Main|
|Pinus mugo (mountain pine)||Pinaceae||Main|
|Pinus muricata (bishop pine)||Pinaceae||Main|
|Pinus nigra (black pine)||Pinaceae||Main|
|Pinus oocarpa (ocote pine)||Pinaceae||Other|
|Pinus palustris (longleaf pine)||Pinaceae||Main|
|Pinus patula (Mexican weeping pine)||Pinaceae||Other|
|Pinus pinaster (maritime pine)||Pinaceae||Main|
|Pinus pinea (stone pine)||Pinaceae||Main|
|Pinus ponderosa (ponderosa pine)||Pinaceae||Other|
|Pinus radiata (radiata pine)||Pinaceae||Main|
|Pinus resinosa (red pine)||Pinaceae||Main|
|Pinus rigida (pitch pine)||Pinaceae||Main|
|Pinus serotina (pond pine)||Pinaceae||Main|
|Pinus strobus (eastern white pine)||Pinaceae||Main|
|Pinus sylvestris (Scots pine)||Pinaceae||Main|
|Pinus taeda (loblolly pine)||Pinaceae||Main|
|Pinus tecunumanii (tecun uman pine)||Pinaceae||Other|
|Pinus thunbergii (Japanese black pine)||Pinaceae||Main|
|Pinus virginiana (scrub pine)||Pinaceae||Main|
Growth StagesTop of page Flowering stage, Fruiting stage, Seedling stage, Vegetative growing stage
SymptomsTop of page The fungus produces two types of necrotic lesions on infected needles. The first type is initially straw yellow, becoming light brown with a dark border. The second type of lesion is a "bar spot" and consists of a brown spot on an amber yellow band. The yellow tissue is infiltrated with resin (Sinclair et al., 1987; Hansen and Lewis, 1997).
On needles of Pinus mugo symptoms first appear on needles as yellow to light orange, sometimes resin-soaked spots, which later become dark-brown in the centre with a yellow border (see Pictures). They usually enlarge to bands that encircle needles and cause death of parts beyond. Diseased needles typically show dead tips, central zones with spots in green tissue, and green bases (see Pictures) (Pehl and Wulf, 2001).
The transition from lesions to healthy green tissue is abrupt. In the brown-coloured dead parts of the needle, black stroma of the fructification develops under the epidermis visible as round black spots (see Pictures). During further development the oval-shaped fruit bodies, arranged parallel to the long axis of the needle, break through the epidermis opening by a longitudinal slit, or two slits, raising a flap of epidermis and hypodermal tissue (see Pictures). Under moist conditions, mature conidiomata produce mucilaginous, olive green spore masses (see Pictures). After severe attack the whole needle initially turns brown, then light brown to grey (see Pictures), and abscises prematurely. Less severe damage may delay needle fall for one or two years. Heavily infected pines typically show twigs with only last year's needles, giving a paintbrush appearance as the pathogen develops; these needles may shed. Over several years this may result in branch and tree death (Pehl and Wulf, 2001).
Often needles of lower branches are attacked by the brown-spot fungus and the pathogen gradually moves up the crown.
List of Symptoms/SignsTop of page
|Leaves / abnormal colours|
|Leaves / abnormal leaf fall|
|Leaves / fungal growth|
|Leaves / necrotic areas|
Biology and EcologyTop of page Life Cycle
If the brown-spot fungus is established in an area, it spreads each spring by conidiospores. Conidia are produced under moist conditions such as rainfall or fog and are mostly passively transported in water droplets and wind-driven water onto needles. The conidia can also be spread by insects and on forestry equipment such as shearing tools (Skilling and Nicholls, 1974). After germination on the needle surface, hyphae enter the needle through the stomata (Setliff and Patton, 1974). However, under certain conditions, the brown-spot fungus may also enter through wounds (Skilling and Nicholls, 1974; Kais, 1975a). The main period for infection is from spring to late summer, but spore dispersal can vary independently of climate. After infection occurs, characteristic symptoms develop. Conidial stroma begin to form as soon as the mesophyll tissues become necrotic and the invading mycelium becomes intracellular (Wolf and Barbour, 1941). After a period of stroma development, mature conidiomata break through the epidermis. The period from inoculation to display of symptoms and mature fruiting bodies varies with temperature, time of year and species of pine; from about 1 or 2 months on young foliage to 6 months or more on old foliage. In general, succulent needle tissue is more susceptible than mature tissue.
Ascospores were also produced, but not in all countries or localities where the anamorph occurs. In the southern USA ascospores and conidia are produced whereas in central and northern areas all infections are apparently due to conidia; ascomata and ascospores are not found. Ascospores mature at any time within 2 to 3 months after infected needle tissues die (Lightle, 1960). The spores are discharged from the ascomata during rain, dew formation or fog and are dispersed by wind, within and beyond the immediate locality.
Factors Affecting Infection
Warm, wet weather favours brown-spot needle blight. Fruiting bodies must be moist to produce spores. Spores germinate and enter needles only when they are wet. Although infection can occur over a wide range of temperatures, it is most rapid in longleaf pine if day and night temperatures are about 30°C and 21°C, respectively (Sinclair et al., 1987).
Means of Movement and DispersalTop of page Natural Dispersal
Conidiospores are only produced under moist conditions and exude from acervulus in a mucilaginous, olive-green, wedge-shaped spore mass. Conidia are dispersed over short distances (from tree to tree) by rain splash. The conidia are spread over longer distances by wind-driven water.
During rain, dew formation or fog, ascospores were discharged from the fruit bodies and windblown within and beyond the immediate locality.
Conidia can also be spread by insects or on forestry equipment such as contaminated shearing tools (Skilling and Nicholls, 1974).
Brown spot needle blight has been moved with seeds contaminated with needle debris (Smith et al., 1997).
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|
|Leaves||fruiting bodies||Yes||Yes||Pest or symptoms usually visible to the naked eye|
|Seedlings/Micropropagated plants||Yes||Yes||Pest or symptoms usually visible to the naked eye|
|Stems (above ground)/Shoots/Trunks/Branches||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)|
|Growing medium accompanying plants|
|True seeds (inc. grain)|
Wood PackagingTop of page
|Wood Packaging not known to carry the pest in trade/transport|
|Loose wood packing material|
|Processed or treated wood|
|Solid wood packing material with bark|
|Solid wood packing material without bark|
ImpactTop of page In Central America, M. dearnessii is omnipresent but rarely in the form of a serious needle blight. Often secondary needles are affected, causing some premature needle cast on hosts in natural pine stands. Severe needle disease can be observed on hosts at the extremes of their altitudinal range (Evans, 1984).
First reports of the brown-spot fungus as a serious needle blight pathogen are on Pinus palustris (longleaf pine) in the Gulf States of the USA. (Hedgcock, 1929). For Evans (1984) this is additional evidence to the hypothesis that the fungus is an exotic in this region, attacking susceptible and non-adapted indigenous pine species. Brown-spot blight continues to be the most important disease of Pinus palustris in the southern USA. This pathogen seems to be the main limiting factor to the establishment of this pine species in its natural range (Henry, 1954; Jewell, 1983). On P. ponderosa in Missouri, Luttrell (1949) associated a serious decline with brown spot needle blight. In several north-central states the brown-spot fungus is a major constraint to the growing of P. sylvestris for Christmas trees (Nicholls et al., 1973; Skilling and Nicholls, 1974) making the affected trees unsaleable. There are also reports from the Altiplano of Colombia that the pathogen is the cause of a severely debilitating needle cast of P. radiata (Gibson 1980).
DiagnosisTop of page
Reliable identification of the brown-spot fungus is only possible through evidence of the characteristic conidia of the anamorphic state (Lecanosticta acicola) or by identification using molecular biological methods.
If there are no mature conidiomata present, isolation of the fungus from the needle is possible. Samples are best taken from affected needles with parts of brown, dead tissue (brown spots, bands or dead parts with black stroma spots). After sterilisation of the needle surface, needles are cut in segments of 4 to 6 mm under sterile conditions and then placed on malt extract agar medium (MEA: 2% malt extract, 2% agar agar) in 9 cm Petri dishes.
Method for surface sterilisation of pine needles:
Needles are immersed in:
70% ethanol for 30 sec.
NaOCl (commercial bleach, 2% of active chlorine) for 60 sec.
96% ethanol for 10 sec.
sterile water for cleaning.
After an incubation period of 1 to 3 weeks at room temperature and daylight, isolates can be examined. Mycelium usually appears on the two cut surfaces of the needle segment. If there are fast- and slow-growing fungi visible, the slow-growing mycelia must be subcultured on fresh malt extract agar. Because of the slow mycelial growth in culture and the presence of endophytic fungi in pine needles, isolation is not always successful. Other fungi or bacteria also present in needles that show rapid growth on culture medium often mask possible isolation of the pathogen.
Identification can be made on culture where the conidia is also produced. On MEA, a white aerial mycelium appears first, which turns greenish-olive to dark olive, forming stromatic and erumpent colonies. At 20°C the mycelium growth reaches 2.5-3 mm a week. Conidia of the anamorph are visible as olive-blackish slimy spore masses. Colonies also produce yellow diffusates on MEA (see Pictures).
Under the microscope the conidia are subhyaline to light brown-olive, thick-walled, spinulose to verrucose, straight to curved, fusiform to cylindrical, 1-5 septate, 14-32-50 x 3-5 µm, with a rounded apex and truncate base (see Pictures). The best view is achieved using differential interference contrast optics at an magnification > 400 times.
It is also possible to identify M. dearnessii by molecular biological methods. Pehl et al. (2004) used a PCR-based ITS-RFLP technique for differentiating M. pini from M. dearnessii and ten other fungi that occur frequently in Europe on pine needles. rDNA is obtained from 0.5-1 mg freeze dried mycel of fungi grown on liquid culture media or direct by 1 or 2 infected needle segments (1-3 mm length, 1.5-3 mg) showing immature fruit bodies or stroma. Mycelium or fresh needle segments are homogenized using a glass micro mortar (25-100 µl, Roth) and extraction buffer from the Qiagen DNeasy Plant Mini Kit. rDNA is isolated by using the Qiagen DNeasy Plant Mini Kit following the manufacturer's instructions. Because of the smaller quantity of testing material the amounts listed in the protocol (e.g. buffers, RNase) were reduced to ¼ of the prescribed values. rDNA concentration is determined fluorimetrically using fluorescent dye Hoe 33258 and a DyNa Quant 200 fluorometer (Pharmacia).
PCR is carried out employing 50 µl reaction volume and a Biometra Tpersonal 48 thermocycler. The reaction mixture contains 3 units Taq DNA polymerase (Strategene), 5 µl 10x reaction buffer (Stratagene), 0.5 mM MgCl2, 0.1 mM dNTP's (Boeringer Mannheim), 0.6 µM forward ITS 4-primer 5'-TCCTCCGCTTATTGATATGC-3' and 0.6 µM reverse ITS 5-primer 5'-GGAAGTAAAAGTCGTAACAAGG-3' according to White et al. (1990) and 2 ng template rDNA. The PCR program consists of an initial denaturation for 2 ½ min at 94°C, 35 cycles with 1 min denaturation at 94°C, 1 min annealing at 55°C and 2 min extension at 72°C, and a final extension for 5 min at 70°C. After completion of the PCR, aliquot samples are separated electrophoretically using a 2% agarose gel, and amplified rDNA is stained with 1 µl/ml ethidium bromide and visualized using an UV-transilluminator.
Aliquot samples containing the amplified rDNA are digested with the restriction endonucleases Hae, Hha I, Hpa II, Hinf I and NCI I (Gibco) following the manufacturer's instructions. The DNA restriction fragments obtained are separated electrophoretically using a 2.5% agarose gel and visualized as described above. Fragment sizes are estimated by comparison with a DNA size marker (100 bp ladder, Gibco).
Mycosphaerella dearnessii is identified on the basis of the species-specific DNA restriction fragments obtained by carrying out the ITS-RFLP analysis as described above. Pictures of species differentiation by ITS-RFLP restriction fragment patterns of M. dearnessi, M. pini and a further 10 pine needle fungi are shown by Pehl et al. (2004).
A diagnostic protocol for Mycosphaerella dearnessii is described in EPPO (2008).
Detection and InspectionTop of page Attack by Mycosphaerella dearnessii causes premature needle fall. Less severe damage may delay needle fall for one or two years: mostly 2- and 3-year-old needles are cast. Heavily infected pines typically show twigs with only the last year's needles, giving a paintbrush appearance. On infected needles, symptoms initially appear in late summer. A quick presumptive indication of needle attack with M. dearnessii is pine needles showing yellow, occasionally resin-soaked spots which later become brown in the centre contrasting to the prominent yellowish border (see Pictures). But final confirmation can only be reached by microscopic identification of the conidial stage, when the typical conidia are produced or by identification with molecular biological methods.
Similarities to Other Species/ConditionsTop of page Sometimes symptoms may be confused with adverse environmental conditions such as chlorofluorocarbon and sulphur dioxide pollution or needle discoloration caused by nutrient deficits such as magnesium and potassium.
The macroscopic symptoms and morphological features of Mycosphaerella dearnessii (anamorph: Lecanosticta) can easily be confused with red band needle blight (Mycosphaerella pini, anamorph: Dothistroma) especially at the beginning of the disease and later, if typically red bands are not produced yet or suppressed (Pehl and Wulf, 2001). Also some disease stages of Mycosphaerella gibsonii (anamorph: Pseudocercospora), which causes cercospora needle blight (brown needle disease), are very close to M. dearnessii. Because of the very similar morphological features the teleomorphs of these three species are difficult to distinguish without any other characteristic information such as a profuse reddish tint to the necrotic needle tissue characteristic to attack with M. pini (Evans, 1984).
The differences between the anamorphs are the best way to separate these species. Lecanosticta and Dothistroma form conidiomata varying between acervuli and pseudopycnidia, controlled by host and climate. They differ from the fruit bodies of Pseudocercospora which form sporodochia. The most important and consistent feature that distinguishes M. dearnessii from M. pini is the nature of the spore wall of the conidia. Conidia of M. dearnessi have melanin granules integrated in the outer wall of the conidia. Under the microscope, the spores are pigmented, thick-walled with a verrucose surface. The best view is achieved using differential interference contrast optics at > 400 times magnification. In contrast, the conidia of M. pini are hyaline, thin-walled and smooth. Sizes of the conidia do not differ distinctly. An illustrated comparison of M. dearnessii and M. pini needle disease is given by Pehl and Wulf (2001).
The anamorphic states of these three fungi can also be distinguished in culture. On malt extract agar colonies of M. pini are slow growing, grey-brown-black, stromatic and produce a whitish conidial slime. A reddish-brown diffusate is present in the agar. M. dearnessi on the same media grows significantly larger and show green-black stromatic colonies with an olive green conidial slime. Typically a yellow diffusate is visible. M. gibsonii is the fastest growing species and forms woolly mycelial grey-coloured colonies (Evans, 1984).
Necrotic brown spots on attacked needles caused by Brachonyx pineti can also be confused with the brown-spot state of M. dearnessii. For B. pineti, a little round hole in the centre of the necrotic spot can be seen with a hand lens. This hole enables the insect to get to the needle tissue. Spots caused by M. dearnessii never show any holes.
Prevention and ControlTop of page
Cultural Control and Sanitary Methods
In longleaf pine stands brown-spot needle blight can be controlled by prescribed ground fires. Young pines in the grass stage can survive these low-intensity fires, which are used in winter to destroy dead infected foliage that harbors the pathogen (Sinclair et al., 1987). Another practical suppression method is shelterwood regeneration. This is the most promising approach to natural regeneration of longleaf pine in which seedlings are established as advanced reproduction under overstories of medium density. The pine canopy then protects the regenerated seedlings from brown-spot infection (Phelps et al., 1978).
Dense planting supports infection so it is important to keep enough space between seedlings, in both nurseries and plantations. While the foliage is wet, shearing and other cultural operations that could spread spores should be avoided (Sinclair et al., 1987). The grower should avoid planting his land with just one pine species or variety to prevent catastrophic losses (Phelps et al., 1978).
In Scots pine, long-needle varieties are somewhat resistant to infection and should be the preferred species in Christmas tree plantations. Varieties such as Austrian Hill or German should be planted to reduce brown-spot infections (Phelps et al., 1978).
Extensive experiments on spraying to control brown-spot in nurseries were conducted by Siggers (1932, 1944). The fact that conidia may be produced in any season throughout the year; that they are distributed by rain-splash; that ascospores are formed in dead tissues most on fallen needles and are air-borne; and that several crops of new needles may be formed within a year indicate that numerous sprayings would be required to give adequate protection.
According to Phelps et al. (1978), Skilling and Nicholls (1974) and Kais (1975b), brown spot needle blight is easily suppressed by applications of Bordeaux mixture, chlorothalonil, benomyl and copper hydroxide in nurseries, seed orchards, and plantations of longleaf pine and Scots pine. Seedlings should be sprayed at 10- to 30-day intervals depending on the amount of rainfall, from the beginning of spring through late summer. It is important to initiate spraying in the spring when the newly emerging fascicled needles are 2 to 5 cm long. Usually four to six applications are sufficient (Phelps et al., 1978). It is also recommended to make a final spray just prior to planting. This will ensure protection during establishment of seedlings in the field.
ReferencesTop of page
Arx JA von, 1983. Mycosphaerella and its anamorphs. Proceedings of the Koninklijke Nederlandse Akademie van Wetenschappen, Series C, 86: 15-54.
Barr ME, 1972. Preliminary studies on the Dothideales in temperate North America. Contrib. Univ. Mich. Herb., 9:523-638.
Barr ME, 1996. Planistromellaceae, a new family in the Dothideales. Mycotaxon, 60:433-442.
Chandelier P; Lafaurie C; Maugard F, 1994. Decouverte en France de Mycosphaerella dearnessii sur Pinus attenuata x radiata. C.R. Acad. Agric. Fr., 80:103-108.
Crous PW; Wingfield MJ, 1966. Species of Mycosphaerella and their anamorphs associated with leaf blotch disease of Eucalyptus in South Africa. Mycologia, 88:441-458.
Dearness J, 1926. New and noteworthy fungi IV. Mycologia, 18:236-255.
EPPO, 2014. PQR database. Paris, France: European and Mediterranean Plant Protection Organization. http://www.eppo.int/DATABASES/pqr/pqr.htm
Gibson IAS, 1979. Diseases of forest trees widely planted as exotics in the tropics and southern hemisphere. Part II. The genus Pinus. Diseases of forest trees widely planted as exotics in the tropics and southern hemisphere. Part II. The genus Pinus. Commonwealth Forestry Institute. Kew, UK; Commonwealth Mycological Institute; Oxford UK, xi + 135 pp.
Goodwin SB; Dunkle LD; Zismann VL, 2001. Phylogenetic analysis of Cercospora and Mycosphaerella based on the internal transcribed spacer region of ribosomal DNA. Phytopathology, 91:648-658.
Hedgcock GG, 1929. Septoria acicola and the brown-spot disease of pine needles. Phytopathology, 19:993-999.
Henry BW, 1954. Sporulation by the brown spot fungus on long leaf pine needles. Phytopathology, 44:385-386.
Jewell FF, 1983. Histopathology of the brown spot fungus on longleave pine needles. Phytopathology, 73:854-858.
Kovacevski JC, 1938. Parasitic fungi new for Bulgaria. Fifth contribution. Rev. Inst. Rech. Agron. Bulg., 8:3-13.
La Porta N; Capretti P, 2000. Mycosphaerella dearnessii, a needle-cast pathogen on mountain pine (Pinus mugo) in Italy. Plant Disease, 84:922.
Laut JG; Sutton BC; Lawrence JJ, 1966. Brown spot needle blight in Canada. Plant Disease Reporter, 50:208.
Lightle PC, 1960. Brown-spot needle blight of Longleaf Pine. US Department of Agriculture, Forest Service, Forest Pest Leaflet, 44:1-7.
Luttrell ES, 1949. Scirrhia acicola, Phaeocryptopus pinastri and Lophodermium pinastri associated with the decline of Ponderosa pine in Missouri. Plant Disease Reporter, 33:397-401.
Martinez JB, 1942. Las Micosis del Pinus insignis en Guipuzcoa. Inst. For. Invest. Exp. Madr., 13.
Morelet M, 1968. De Aliquibus in Mycologia Novitatibus (3 note). Bull. Soc. Sci. Nat. Archeol. Toulon. Var., 177:9.
Nicholls TH; Skilling DD; Hudler GW, 1973. Scirrhia acicola in Scotch pine Christmas tree plantations. Plant Disease Reporter, 56:712-713.
Pehl L; Burgermeister W; Wulf A, 2004. Mycosphaerella-Nadelpilze der Kiefer - Identifikation durch ITS-RFLP-Muster. Nachrichtenbl. Deut. Pflanzenschutzd., (in press).
Pehl L; Wulf A, 2001. Mycosphaerella-needle fungi on pines - symptoms, biology and differential diagnosis. Nachrichtenblatt des Deutschen Pflanzenschutzdienstes, 53:217-222.
Petrak F, 1961. Die Lecanosticta-Krankheit der Föhren in Österreich. Sydowia, 15:252-256.
Phelps WR; Kais AG; Nicholls TH, 1978. Brown-spot needle blight of pines. U.S. Department of Agriculture, Forest Service, Forest Insekt & Disease Leaflet, 44:1-8.
Saccardo PA, 1884. Sylloge Fungorum, 3:507.
Schischkina AC; Tzanava NI, 1967. Systremma acicola (Dearn.) Wolf et Borbour - Stadium perfectum Dothistromatis acicolae (Thüm.) A. Schischk. Et N. Tzan. Nov. Sist. Niz. Rast., 276.
Siggers PV, 1932. The brown-spot needle blight of longleaf pine seedlings. Jour. Forestry, 30:579-593.
Siggers PV, 1944. The brown spot needle blight of pine seedlings. U.S.D.A. Technical Bulletin, 870:1-36.
Smith IM; McNamara DG; Scott PR; Holderness M, 1997. Quarantine pests for Europe. Second Edition. Data sheets on quarantine pests for the European Union and for the European and Mediterranean Plant Protection Organization. Quarantine pests for Europe. Second Edition. Data sheets on quarantine pests for the European Union and for the European and Mediterranean Plant Protection Organization., Ed. 2:vii + 1425 pp.; many ref.
Sydow H; Petrak F, 1922. Ein Beitrag zur Kenntnis der Pilzflora Nordamerikas, insbesondere der nordwestlichen Staaten. Annales Mycologici, 20:178-218.
Sydow H; Petrak F, 1924. Zweiter Beitrag zur Kenntnis der Pilzflora Nordamerikas, insbesondere der nordwestlichen Staaten. Annales Mycologici, 22:387-409.
Thümen F de, 1878. Fungorum americanorum triginta species novae. Flora, 61:177-184.
Wolf FA; Wolf FT, 1947. The Fungi, Volume 1. New York, USA: J. Wiley & Sons.
Distribution MapsTop of page
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