Cyclaneusma minus
(Cyclaneusma needle-cast)

Pictures

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Identity

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Preferred Scientific Name

• Cyclaneusma minus (Butin) DiCosmo, Pered & Minter 1983

Preferred Common Name

• Cyclaneusma needle-cast

Other Scientific Names

• Naemacyclus minor Butin 1973
• Naemacyclus niveus (pro parte) (Pers.) Sacc.

International Common Names

• English: Naemacyclus needle cast

EPPO code

• NAEMMI (Naemacyclus minor)

Summary of Invasiveness

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C. minus is spread by airborne ascospores. Its host range is restricted to some species in a single genus, Pinus.

Taxonomic Tree

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• Domain: Eukaryota
•     Kingdom: Fungi
•         Phylum: Ascomycota
•             Subphylum: Pezizomycotina
•                 Class: Leotiomycetes
•                     Genus: Cyclaneusma
•                         Species: Cyclaneusma minus

Notes on Taxonomy and Nomenclature

Top of page This fungus has long been known as Naemacyclus niveus in the forest pathology literature. Unfortunately, this name was based on a long-standing error as to the identity of the type species of Naemacyclus. Korf (1962) pointed out that Fuckel had based the genus Naemacyclus on Propolis pinastri (=Naemacyclus pinastri) and not, as later authors had supposed, on Propolis nivea (Pers.) Fr. (=Naemacyclus niveus). Examination of the type material of the original species (N. pinastri) showed that it was not conspecific with N. niveus but rather with Stictis fimbriatus Schwein., the type of the genus Lasiostictis (Sacc. & Berl.) Sacc. This meant that Naemacyclus must be applied to Lasiostictis and the former genus, as commonly understood, must be given a new name. A new generic name, Cyclaneusma, was introduced by DiCosmo et al. (1983) to accommodate N. niveus (and the similar N. minor). Another complication in the literature arises from the fact that Butin (1973) showed that N. niveus comprised two similar but distinct species, N. niveus and N. minor. Naemacyclus minor (=Cyclaneusma minus) is the more economically important species but in older (pre-1973) records it cannot be distinguished from N. niveus.

Description

Top of page Ascomata are apothecial, scattered, subepidermal, somewhat rectangular in appearance when partially open, elliptical when fully open, waxy, reddish-brown when young, later becoming concolorous with the needle surface, 0.1-0.7 (mostly 0.3-0.4) mm long x 0.2-0.25 mm wide. As they develop, ascomata push through the needle epidermis, opening by a single longitudinal split (usually along a row of stomata) and raising the needle tissue on either side in one or usually two flaps which remain hinged at the sides. Mature ascomata swell when moist and the hinged flaps, composed of hyaline hyphae and the needle cuticle, epidermis and hypodermis, are pushed back, exposing a slightly convex, straw-coloured hymenial layer. Asci are unitunicate, subcylindrical, 90-120 x 8-12 µm, containing eight ascospores fasciculately arranged and sometimes twisted in a helix. Ascospores are filiform, (0-)2-septate, 65-95 x 2-3 µm, hyaline, smooth, usually slightly bent. Paraphyses filiform, branched towards the apex, aseptate, straight, hyaline, smooth. Conidiomata are pycnidial, scattered, deeply immersed, globose to subglobose, 0.1-0.2 mm diam, with walls composed of hyaline, pseudoparenchymatous cells, 2-3 µm in diam. Conidia are bacilliform, non septate, 6-10 x 1 µm, hyaline, smooth.

Distribution Table

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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.

History of Introduction and Spread

Top of page Nothing is known about the origin of C. minus. It is probably indigenous throughout the geographical range of Pinus species (i.e. practically the whole of the temperate and sub-temperate Northern Hemisphere) as it is found on indigenous Pinus species in Europe, Asia and North America. It has been accidentally introduced into New Zealand, Australia, Africa and South America where it is found only on introduced exotic Pinus species. There is no information on when the fungus was introduced as there has been much confusion about the identity of the organism.

Risk of Introduction

Top of page C. minus is not a quarantine pest.

Habitat

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C. minus is found in both natural forests and plantations.

Habitat List

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Hosts/Species Affected

Top of page Although a large number of Pinus species has been recorded as hosts of C. minus, the fungus has been experimentally shown to be pathogenic only to Pinus radiata (Gadgil, 1984) and Pinus sylvestris (Kistler and Merrill, 1977; Karadzic and Millar, 1981; Wenner and Merrill, 1986). Young plants of P. radiata <3 years old) are resistant to infection, whereas P. sylvestris plants are susceptible at all ages. In P. radiata in New Zealand, disease severity was found to be highest in 11-20-year-old stands, lower in over 25-year-old stands and lowest in 1-5-year-old stands (Bulman, 1988).

Host Plants and Other Plants Affected

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Growth Stages

Top of page Flowering stage, Fruiting stage, Seedling stage, Vegetative growing stage

Symptoms

Top of page In Pinus radiata, symptoms first appear on 1-year-old or older needles in the central and lower parts of the crown. Needles turn a mottled yellow-green at first and then a mottled yellow-brown a few weeks later. In highly susceptible trees, almost the whole crown may be affected. Transverse reddish bands are also commonly seen. In some years and some localities, needles finally become a uniform reddish-brown rather than the more usual mottled yellow-brown. Needles showing symptoms are readily detached from the tree and most are shed prematurely, generally in the spring. By early summer, the crowns of infected trees look very thin, holding only the newly flushed foliage. The susceptibility of trees to the disease is very variable and stands usually contain a mixture of susceptible trees, recognizable in spring by their yellow-brown crowns, and resistant trees with green crowns. Resistant trees, which do not develop symptoms of infection, are not necessarily immune to infection by C. minus although C. minus populations in needles of healthy trees are lower than those of diseased trees (van der Pas et al., 1984b). The severity of the disease varies considerably from year to year.

In P. sylvestris, symptoms of infection are found on needles of all ages in Europe and on 1-year-old or older needles in North America. The first symptoms appear as small, light-green spots, which coalesce turning the needle a dusty yellow with transverse brown bands. Finally, the needles become tannish brown. Infected needles are usually cast within a few months of the appearance of the symptoms.

List of Symptoms/Signs

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Biology and Ecology

Top of page In New Zealand, C. minus infects susceptible trees of P. radiata at temperatures ranging from 10 to 25°C. Current season needles are resistant to infection until they are 8-9 months old. They become infected in autumn to early winter, begin to develop symptoms of infection by mid-winter, and are usually cast in early spring when they are about 1 year old (Gadgil, 1984). The infection process is more prolonged on P. sylvestris in Pennsylvania, USA (Wenner and Merrill, 1986). Symptoms appear on needles in the summer of the second growing season even though the needles become susceptible to infection in mid-summer of the first growing season. These needles are cast at the end of the summer or in autumn when they are about 2 years old. In Europe, Karadzic and Millar (1981) reported that symptoms on P. sylvestris needles appear within 2 to 3 months of initial infection. In Germany, Rack (1981) found that C. minus could be first isolated from the current season's needles when they were about 6 months old.

Ascomata of C. minus are rarely seen on needles attached to the tree but develop readily on fallen needles. In New Zealand, ascomata develop within 2 weeks of needle fall and continue producing ascomata for 6-24 weeks. Ascomata production occurs throughout the year but the greatest number of ascomata per unit area is produced during autumn and winter (Gadgil, 1984). In Pennsylvania, USA, ascomata are produced in autumn and winter on needles cast in late summer or autumn in their second growing season. These continue development during the winter and become mature in the following spring (Wenner and Merrill, 1986). A period of rainfall (defined as 0.1 mm or more precipitation per hour) is required for ascospore release. Pawsey (1967) found that in South Australia, the numbers of ascospores trapped reached a peak within 2-3 hours of the onset of rainfall. In New Zealand, the highest number of ascospores was trapped in the fifth hour after the commencement of rainfall. Ascospores were trapped throughout the year but there was a marked seasonal variation in the frequency of occurrence of ascospores, with major peaks in autumn and winter (Gadgil, 1984). Wenner and Merrill (1986) reported that in Pennsylvania, USA, ascospore numbers peaked 2-4 hours after the onset of rain. Ascospores were released from spring to mid-winter. The major peak period of ascospore release was late spring; the numbers then declined through summer until mid to late autumn when a minor peak period of spore release was noted.

Kowalski and Millar (1981) reported that infection of P. sylvestris needles decreased with increasing concentrations of air pollutants. In a study of the influence of climate on three fungi inhabiting conifer needles, van Maanen et al. (2000) found that the presence and abundance of C. minus could not be predicted by climate alone at a regional scale.

At least two morphologically distinct types of C. minus have been found in the New Zealand population but it is not yet known whether these morphological differences reflect differences in pathogenicity (Dick et al., 2001).

Cyclaneusma needle-cast is a disease of complex aetiology and the interactions between host genotype, host nutrition and the endophytic, variable fungal population are not fully understood.

Means of Movement and Dispersal

Top of page Natural dispersal of C. minus is by airborne ascospores.

Seedborne Aspects

Top of page C. minus is not seedborne.

Plant Trade

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Plant parts not known to carry the pest in trade/transport
Bark
Bulbs/Tubers/Corms/Rhizomes
Flowers/Inflorescences/Cones/Calyx
Fruits (inc. pods)
Growing medium accompanying plants
Roots
Stems (above ground)/Shoots/Trunks/Branches
True seeds (inc. grain)
Wood

Wood Packaging

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Impact Summary

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Economic Impact

Top of page In Pinus sylvestris Christmas tree plantations in Wisconsin, USA, Cyclaneusma needle-cast was estimated to have reduced the value of the crop by 26% (Ostry et al., 1990).

In Pinus radiata in New Zealand, trials carried out to explore the relationship between disease severity (percentage of green crown showing symptoms of the needle-cast) and growth, showed that an average disease severity of 60% over 6 years resulted in a 50% loss in diameter increment (Bulman and van der Pas, 2001). Projections of stand growth to age 30 for various proportions of diseased trees showed a reduction in volume of 10-14 m³/ha for each 10% increase in the proportion of diseased trees. For the country as a whole, a growth loss of 6.6% per annum for the P. radiata estate aged between 6 and 20 years was predicted with a corresponding financial loss of the order of $51 million per year (Bulman, 2001a).

Environmental Impact

Top of page The environmental impact of C. minus has not been assessed but is not likely to be important.

Diagnosis

Top of page C. minus grows readily on standard mycological media (e.g. 3% MEA) at normal laboratory temperatures (15-25°C). The standard method of isolation is to surface sterilize the needles, cut them into suitable sized segments (usually 10-30 mm) and place them on agar plates. It is usual to incubate the plates at 18-20°C as at higher temperatures C. minus colonies are often swamped by colonies of other endophytic fungi, particularly species of Lophodermium. Colonies of C. minus are reasonably easy to recognize. In culture on 3% MEA, the mycelium is white at first, later becoming pink-white to pink-beige, fluffy, collapsing in patches, margin diffuse. Ascomata and conidiomata are produced in culture, making identification simple.

Detection and Inspection

Top of page The characteristic symptoms of Cyclaneusma needle-cast, particularly the mottled yellow-brown, easily detached needles, are readily recognizable in the field. It is more difficult to detect infected trees in the summer and autumn, when the infected needles have all been cast and the symptoms of new infection have not yet appeared. The sparse crowns, bearing only the current foliage, provide a clue.

Assessment of the severity of infection (percentage of green crown showing symptoms) should be made at a time of maximum symptom expression (usually spring). Infection percentage can be estimated in 5% steps using a method developed for assessment of Dothistroma pini infection (Kershaw et al., 1988). Assessments are made either from the ground or air, depending on the purpose of the assessment. For research purposes, when a coefficient of variation of less than 10% is desirable, ground assessments made by at least three trained observers are required. For a general survey, where a coefficient of variation of up to 25% is acceptable, aerial assessments (fixed wing aircraft, flying at about 100 m above ground level at 80 knots) made by two observers provides an adequate level of accuracy (van der Pas et al., 1984a).

Similarities to Other Species/Conditions

Top of page In the field, symptoms of Cyclaneusma needle-cast may be confused with those of a condition possibly associated with magnesium deficiency known as 'upper mid-crown yellowing'. The symptoms have also occasionally been confused with those of Dothistroma needle-blight although Dothistroma-infected needles are usually redder and the infection is generally confined to the lower crown. In coastal forests, salt spray damage to tree crowns may be confused with symptoms of Cyclaneusma infection.

Microscopically, C. minus could be confused with the similar Cyclaneusma niveum. C. niveum has larger ascomata (0.3-1.0 mm long) and ascospores (75-120 x 3-4 µm) and it has sickle-shaped, rather than bacilliform, conidia.

Prevention and Control

Top of page In New Zealand, where the proportion of Cyclaneusma-susceptible trees in Pinus radiata stands is rarely above 60%, control of the disease through modifying silvicultural practices is practicable. By adopting susceptibility to Cyclaneusma needle-cast as a primary selection criterion for thinning, it is possible to achieve an almost disease-free final crop stand. It is necessary to delay the first thinning to age 7 or 8 when symptoms of Cyclaneusma infection are easily detected. The ideal silvicultural regime for control of the disease is a heavy delayed first thinning (e.g. from 1250 stems/ha to 400-500 stems/ha at age 7) followed by a second thinning at age 9 or 10 (to a final stocking of 250 stems/ha) to remove the remaining disease-susceptible trees (Bulman, 2001b).

Chemical control of the disease is possible but not economically justifiable, except perhaps in Christmas tree plantations. In Pinus sylvestris in Pennsylvania, USA, five applications of chlorothalonil were found to give adequate control (Wenner and Merrill, 1990). In New Zealand, monthly aerial applications of dodine for 6 months gave good control of the disease in P. radiata (Hood and Bulman, 2001).

References

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Bulman LS, 1988. Incidence and severity of Cyclaneusma needle-cast in fifteen Pinus radiata plantations in New Zealand. New Zealand Journal of Forestry Science, 18(1):92-100

Bulman LS, 2001. Economic impact of the disease. In: Bulman LS, Gadgil PD, eds. Cyclaneusma needle-cast in New Zealand. Forest Research Bulletin No. 222. Rotorua, New Zealand: New Zealand Forest Research Institute, 42-48.

Bulman LS, 2001. Silvicultural control. In: Bulman LS, Gadgil PD, eds. Cyclaneusma needle-cast in New Zealand. Forest Research Bulletin No. 222. Rotorua, New Zealand: New Zealand Forest Research Institute, 55-62.

Bulman LS; van der Pas JB, 2001. Effect of Cyclaneusma needle-cast on growth. In: Bulman LS, Gadgil PD, eds. Cyclaneusma needle-cast in New Zealand. Forest Research Bulletin No. 222. Rotorua, New Zealand: New Zealand Forest Research Institute, 31-41.

Butin H, 1973. Morphologic and taxonomic studies on Naemacyclus niveus (Pers. ex Fr.) Fuck. ex Sacc. and related species. European Journal of Forest Pathology, 3(3):146-163

Cannon PF; Minter DW, 1986. The Rhytismataceae of the Indian subcontinent. Mycological Papers, No. 155. Wallingford, UK: CAB International, 123 pp.

Choi D; Simpson JA, 1991. Needle cast of Pinus radiata in New South Wales. Australian Journal of Botany, 39(2):137-152

Dick MA; Somerville JG; Gadgil PD, 2001. Variability in the fungal population. In: Bulman LS, Gadgil PD, eds. Cyclaneusma needle-cast in New Zealand. Forest Research Bulletin No. 222. Rotorua, New Zealand: New Zealand Forest Research Institute, 12-19.

DiCosmo F; Peredo H; Minter DW, 1983. Cyclaneusma gen. nov., Naemacyclus and Lasiostictis, a nomenclatural problem resolved. European Journal of Forest Pathology, 13(4):206-212

Frederick RD; Zang L; Merrill W; Tattar TA, 1980. Naemacyclus minor needlecast of Scots pine in Massachusetts. Plant Disease, 64(11):1034

Funk A, 1985. Foliar fungi of western trees. Information Report, Pacific Forest Research Centre, Canada, No. BC-X-265, 159 pp.

Gadgil PD, 1984. Cyclaneusma (Npmacyclus) needle-cast of Pinus radiata in New Zealand. 1: Biology of Cyclaneusma minus. New Zealand Journal of Forestry Science, 14(2):179-196

Gilmour JW, 1959. Pathogenic needle-cast fungi. Interim Research Release, Forest Research Institute, New Zealand Forest Service.

Giordano L; Gonthier P, 2011. An outbreak of Cyclaneusma minus needle cast on Swiss mountain pine (Pinus uncinata) in Italy. Journal of Plant Pathology, 93(4, Supplement):S4.74. http://sipav.org/main/jpp/index.php/jpp/issue/view/118

Granger CA; Duncan-Frost G, 1999. Integrated crop management schedule for the production of Christmas trees. Circular 11: Maine Department of Conservation, Maine Forest Service, USA.

Grodnitskaya ID; Senashova VA, 2012. Disease of the common pine seedlings new for the forest nurseries of Siberia. Zashchita i Karantin Rastenii, No.2:48-50. http://www.z-i-k-r.ru

Hartman JR; Hill DB, 1996. Needle cast diseases of conifers. ID-85: Cooperative Extension Service, College of Agriculture, University of Kentucky, USA.

Hâruta O; Fodor E; Teusdea A, 2007. Complex diseases in Pinus nigra Arnold situated along Crisul Repede River Gorge. (Boli complexe la Pinus nigra Arnold în Defileul Crisului Repede.) Analele Institutul de Cercetari si Amenajari Silvice, 50:169-184.

Hood IA; Bulman LS, 2001. Chemical control. In: Bulman LS, Gadgil PD, eds. Cyclaneusma needle-cast in New Zealand. Forest Research Bulletin No. 222. Rotorua, New Zealand: New Zealand Forest Research Institute, 50-54.

Jurc M; Jurc D; Gogala N; Simoncic P, 1995. Air pollution and fungal endophytes in needles of Austrian pine. Bioindication of stress in forest trees and forest ecosystems. Papers given at the closing workshops and colloquium of the TEMPUS JEP programme, held in Ljubljana, Slovenia in August 1995 [coordinator Batic, F.]. Phyton Horn., 36:111-114.

Karadzic D; Millar CS, 1981. Infection of Pinus sylvestris by Naemacyclus minor. Current Research on Conifer Needle Diseases, 99-101.

Karadzic D; Zoric R, 1981. Contribution to the knowledge of the biology of Npmacyclus minor Butin, the pathogen of needle cast of Scots pine. Zastita Bilja, 32(1):79-90

Kershaw DJ; Gadgil PD; Ray JW; van der Pas JB; Blair RG, 1988. Assessment and control of Dothistroma needle blight. 2nd revised edition. Forest Research Institute Bulletin No. 18. Wellington, New Zealand: Ministry of Forestry.

Kistler BR; Merrill W, 1977. Naemacyclus minor, a primary pathogen of Scots pine. Proceedings of the American Phytopathological Society, 3:214-215.

Korf RP, 1962. A synopsis of the Hemiphacidiaceae, a family of the Helotiales (Discomycetes) causing needle blights of conifers. Mycologia, 54:12-33.

Kowalski T; Millar CS, 1981. Fungi infecting needles of Pinus sylvestris in Poland in relation to air pollution zone. Current Research on Conifer Needle Diseases, 93-98; 18 ref.

Maanen Avan; Debouzie D; Gourbiere F, 2000. Distribution of three fungi colonising fallen Pinus sylvestris needles along altitudinal transects. Mycological Research, 104(9):1133-1138; 31 ref.

Marks GC; Fuhrer BA; Walters NEM, 1982. Tree diseases in Victoria. Handbook, Forests Commission Victoria Melbourne Australia, No.1:xviii + 149 pp.

Merrill W; Kistler BR, 1974. Naemacyclus needlecast of Scots Pine epidemic in Pennsylvania. Plant Disease Reporter, 58(3):287-288; 8 ref.

Millar CS; Minter DW, 1980. Naemacyclus minor. CMI Descriptions of Pathogenic Fungi and Bacteria, No. 659. Wallingford, UK: CAB International.

Minter DW, 1994. The Rhytismatales on conifers from Europe. In: Capretti P, Heiniger U, Stephan R, eds. Shoot and Foliage Diseases in Forest Trees, Proceedings of a joint meeting of the Working Parties Canker and Shoot Blight of Conifers (S2.06.02) and Foliage Diseases (S2.06.04), Vallombrosa, Italy: Instituto di Patologia e Zoologia Forestale e Agraria, Universita degli Studi di Firenze, 65-84.

Myren DT(Editor), 1994. Tree Diseases of Eastern Canada. Ottawa, Canada: Canada Communication Group-Publishing, 159 pp.

Narvaez AM, 1999. Enfermedades foliares an Pinus ponderosa: momento de decisiones. Chile Forestal, 24(275):28-29.

Offord HR, 1964. Diseases of Monterey Pine in native stands of California and in plantations of western North America. US Forest Service Research Paper Pacific Southwest Forest and Range Experiment Station, No. PSW-14, pp. 37.

Ostry ME; Nicholls TH; Carlson JC; Adams GC, 1990. Cyclaneusma needlecast in Scots pine Christmas tree plantations in the Lake States. In: Merrill W, Ostry ME, eds. Recent Research on Foliage Diseases, Conference Proceedings, Carlisle, Pennsylvania, USA: USDA Forest Service, General Technical Report WO-56, Washington, DC, USA, 19-21.

Pas JB van der; Kimberley MO; Kershaw DJ, 1984. Evaluation of the assessment of Dothistroma needle blight in stands of Pinus radiata. New Zealand Journal of Forestry Science, 14(1):3-13

Pas JB van der; Slater-Hayes JD; Gadgil PD; Bulman L, 1984. Cyclaneusma (Npmacyclus) needle-cast of Pinus radiata in New Zealand. 2: Reduction in growth of the host, and its economic implication. New Zealand Journal of Forestry Science, 14(2):197-209

Pawsey RG, 1967. Spore discharge of Naemacyclus niveus following rainfall. Transactions of the British Mycological Society, 50(3):341-347.

Peterson GW, 1981. Pine and juniper diseases in the Great Plains. General Technical Report, Rocky Mountain Forest and Range Experiment Station, USDA Forest Service, No. RM-86, ii + 47 pp.; 28 pl. (26 col.); 4 pp. ref.

Peterson GW; Walla JA, 1986. Naemacyclus (Cyclaneusma) needle cast of pines. In: Riffle JW, Peterson GW, eds. Diseases of Trees in the Great Plains. General Technical Report RM-129: Rocky Mountain Forest and Range Experiment Station, Fort Collins, Colorado, USA, 122-123.

Podger FD; Wardlaw TJ, 1990. Spring needle-cast of Pinus radiata in Tasmania: I. Symptoms, distribution, and association with Cyclaneusma minus. New Zealand Journal of Forestry Science, 20(2):184-205

Rack K, 1981. Interactions between Naemacyclus minor and Lophodermium pinastri during 'pine needle cast'. In: Millar CS, ed. Current Research on Conifer Needle Diseases, Proceedings of the International Union of Forestry Research Organizations Working Party on Needle Diseases Conference, Sarajevo, Yugoslavia. Aberdeen, UK: Aberdeen University, 103-111.

Rajchenberg M; Cwielong P; Rovelotti J; Cuevas C, 1995. Falsa Banda Roja (Cyclaneusma minus) en plantaciones de Pináceas exóticas en la Región Andino Patagónica Argentina. Actas de las IV Jornadas Forestales Patagónicas en S. M. de Los Andes, Tomo I, 277-289.

Stahl W, 1966. Needle-cast fungi on conifers in the Australian Capital Territory. Australian Forestry, 30(1):20-32.

Szab= I, 2002. Fungi causing leaf diseases on forest trees II. Leaf and shoot pathogens of conifer trees. No^umlaut~ve^acute~nyve^acute~delem, 38(11):581-585; 17 ref.

Taylor NJ; Nameth ST, 1996. Needle diseases on 2-needle conifers in Ohio. Extension Factsheet HYG-3071-96. Ohio, USA: Ohio State University Extension.

Walkowiak J, 1999. Forests and Prairies Division 1999 Iowa Forest Health Report. Iowa, USA: Forestry Bureau, Iowa Department of Natural Resources.

Wenner NG; Merrill W, 1986. Cyclaneusma needlecast in Pennsylvania: a review. In: Peterson GW, ed. Recent Research on Conifer Needle Diseases, Proceedings of the International Union of Forestry research Organizations Working Party on Needle Diseases Conference, Gulfport, Mississippi, USA. USDA Forest Service, General Technical Report WO-50, Washington, DC, USA, 35-40.

Wenner NG; Merrill W, 1990. Control of Cyclaneusma needlecast on Scots pine in Pennsylvania. In: Merrill W, Ostry ME, eds. Recent Research on Foliage Diseases, Conference Proceedings, Carlisle, Pennsylvania, USA. USDA Forest Service, General Technical Report WO-56, Washington, DC, USA, 27-33.

Distribution Maps

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