Mycosphaerella gibsonii
(needle blight of pine)

Pictures

Picture	Title	Caption	Copyright
Title Field symptoms Caption Mycosphaerella gibsonii (needle blight of pine) ; field symptoms on a two-year-old seedling of Pinus thunbergii (Japanese black pine). Copyright ©H. Hashimoto/Bugwood.org - CC BY-NC 3.0 US	Field symptoms	Mycosphaerella gibsonii (needle blight of pine) ; field symptoms on a two-year-old seedling of Pinus thunbergii (Japanese black pine).	©H. Hashimoto/Bugwood.org - CC BY-NC 3.0 US

Identity

Preferred Scientific Name

• Mycosphaerella gibsonii H.C. Evans 1984

Preferred Common Name

• needle blight of pine

Other Scientific Names

• Cercoseptoria pini-densiflorae (Hori & Nambu) Deighton 1976
• Cercospora pini-densiflorae Hori & Nambu 1917
• Pseudocercospora pini-densiflorae (Hori & Nambu) Deighton 1987

International Common Names

• English: brown needle blight of pine; brown pine needle disease; brown-needle disease; Cercospora blight of pines; Cercospora needle-blight; Cercospora pine blight; needle blight: pine
• Spanish: cercosporiosis de las aciculas del pino
• French: cercosporiose des aiguilles du pin
• Portuguese: quiema de aciculas por Mycosphaerella

EPPO code

• CERSPD (Mycosphaerella gibsonii)

Summary of Invasiveness

Mycosphaerella gibsonii is a fungal pathogen causing needle blight, primarily in Pinus species. It causes lesions on needles, first affecting lower needles and then spreading to the upper crown. The disease eventually causes needle necrosis and needle cast, leading to defoliation, stunted growth and host plant death; it is a major obstacle to the production of pine seedlings. M. gibsonii occurs in the tropics and subtropics of South and Central America, the Caribbean, sub-saharan Africa, India, South East Asia and East Asia; the native range is uncertain. Although natural dispersal by wind and water occur locally, international spread is largely due to movement of infected nursery stock. Phytosanitary control measures such as avoiding the planting of infected plants, removal and destruction of all infected pines in nurseries and cleaning between annual production cycles in nurseries can help to reduce the spread of the pathogen. It is listed as an A1 quarantine pest in the EPPO region, and is considered of quarantine significance in South America. 

Taxonomic Tree

• Domain: Eukaryota
•     Kingdom: Fungi
•         Phylum: Ascomycota
•             Subphylum: Pezizomycotina
•                 Class: Dothideomycetes
•                     Subclass: Dothideomycetidae
•                         Order: Capnodiales
•                             Family: Mycosphaerellaceae
•                                 Genus: Mycosphaerella
•                                     Species: Mycosphaerella gibsonii

Notes on Taxonomy and Nomenclature

This fungus was originally only known in its imperfect state (Cercospora pini-densiflorae), but the teleomorph was later found and identified as Mycosphaerella gibsonii (Evans, 1984). In 2014, the One Name One Fungi project suggested that M. gibsonii should be recognized as Pseudocercospora pini-densiflorae (Sullivan, 2010). However, the Catalogue of Life still recognizes M. gibsonii as the accepted name of this species, with P. pini-densiflorae as a synonym (Kirk, 2019). Braun et al. (2013) suggested that P. montantania, described by Mehrotra (1987) on Pinus kesiya, is morphologically indistinguishable from P. pini-densiflorae [M. gibsonii], and is barely more than a morphological variety of needle blight of pine.

Description

M. gibsonii can be cultured on V8 juice + pine needle decoction agar, in natural light at 25°C (day), 0-10°C (night). Higher night temperatures (15°C) cause abnormal conidial formation (Suto, 1971). The fungus may also be cultured on PCA-UV at 25°C (Sullivan, 2010; Braun et al., 2013).

Stromata are dark-brown, tuberculated, filling the stomatal openings, 60-96 µm diameter. Conidiophores dense, dark-brown, straight or slightly curved, rarely septate and not branched; 10-45 x 2.5 µm; conidiogenous loci unthickened. Conidia pale yellowish-olivaceous, long-obclavate, straight or slightly curved, 3-7 septate with a truncate or rounded unthickened base and obtuse tip; 20-68 (mostly 40-50) x 2.5-4.5 µm (Ito, 1972). Asci bitunicate, clavate to cylindrical (33-) 35-38 x 5.5-7 µm, with thickened, bluntly rounded apex, rarely saccate and 32-36 x 6-8 µm, 8-spored, obliquely biseriate. Interthecial tissue present or absent (Evans, 1984). Ascospores hyaline, 1-septate, ellipsoidal to cuneate, (7.5-) 8.5 x 11 (-12.5) x 2 x 3 µm, guttulate (Evans, 1984).

Spermagonia are formed in discrete, unilocular stromata, or as locules in upper parts of large stromata (Ivory, 1987). They consist of a thin dark-brown wall enclosing white contents. The spermatia form on conidiogenous cells lining the inner wall of the locules, are hyaline, rod-shaped and 2-3 x 1 µm. They often become exuded in tiny hyaline droplets.

The fungus forms grey to grey-green or black, compact colonies which often become pulvinate and stromatic. They are mostly non-sporulating, although Asian isolates quite often produce brown, thin-walled spermagonia containing spermatia in a pale-grey slime when grown under black light. These isolates also produce conidia very occasionally in small fertile patches on stromatic colonies when exposed to black light. The African isolates grow more slowly, are more inhibited by black light, and are generally less spreading than Asian isolates (Ivory, 1987).

Distribution

The fungus is much more widely distributed in tropical and subtropical regions than previously supposed (Ivory, 1994). It occurs across the tropics and subtropics of South and Central America, the Caribbean, sub-saharan Africa, India, South East Asia and East Asia, where it may affect native Pinus species (Quaedvlieg et al., 2012). Records from native Pinus wallichiana and P. roxburghii in Nepal suggest that this fungus could be native to the Himalayas.

See also CABI/EPPO (1998, No. 216).

Distribution Table

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.

Risk of Introduction

In the EPPO and South Cone Plant Protection Committee (COSAVE) regions it is potentially dangerous to Pinus spp. wherever they are grown. Nursery trade increases the risk of spread of M. gibsonii. It is listed as an A1 quarantine pest by EPPO (OEPP/EPPO, 1980). It is also of quarantine significance to the Andean Community (CAN) and COSAVE (Brazil, Ministério da Agricultura e do Abastecimento, 1996; COSAVE, 2018). 

Habitat List

Hosts/Species Affected

M. gibsonii affects many species of Pinus. Additionally, it has recently been reported on Abies nobilis [Abies procera] in Japan (Farr and Rossman, 2018). It is particularly associated with old seedlings in tree nurseries, but is also quite common on older trees of highly susceptible species such as P. canariensis, P. radiata and P. roxburghii. Other susceptible species include P. halepensis, P. pinaster and P. sylvestris (EPPO, 2018). It also occurs on senescing foliage and leaf litter of many species at a much lower incidence level (Ivory, 1994).

During field surveys carried out by Ivory (1994) in Africa, Asia, Central America and Oceania, the fungus was found on Pinus ayacahuite, P. canariensis, P. caribaea, P. clausa, P. densiflora, P. elliottii, P. greggii, P. halepensis, P. kesiya, P. luchuensis, P. massoniana, P. merkusii, P. muricata, P. oocarpa, P. patula, P. pinaster, P. pseudostrobus, P. radiata, P. roxburghii, P. rudis [P. hartwegii], P. sylvestris, P. taeda, P. thunbergii and P. wallichiana. Other reported host species include P. cembra, P. contorta, P. echinata, P. flexilis, P. jeffreyi, P. lambertiana, P. mugo, P. nigra, P. parviflora, P. pinea, P. ponderosa, P. resinosa, P. rigida, P. strobus, P. taiwanensis, P. tabuliformis (Braun et al., 2013), P. maximinoi, P. morrisonicola, P. tecunumanii (Chen, 1965), P. armandii, Tsuga canadensis and Abies nobilis (Farr and Rossman, 2018).

It must also be noted that the following conifer species were found to be susceptible to the pathogen under artificial inoculation experiments: Abies veitchii, A. sachalinensis, Cedrus deodara, Picea glehnii, P. jezoensis, Pseudotsuga menziesii (Suto, 1979; Diekmann et al., 2002) and Larix kaempferi (Kobayashi et al., 1979; Suto, 1979; Diekmann et al., 2002).

Host Plants and Other Plants Affected

Growth Stages

Symptoms

Lesions can occur at any point along infected primary and secondary needles (Ivory and Wingfield, 1986). However, they often appear towards the distal part of the needles, especially on 1- to 2-year-old seedlings (Suto, 1979; Ivory, 1987). Ivory and Wingfield (1986) suggest that foliage of the lower crown is usually the most affected due to the occurrence of more favourable conditions for the infection. Observations in the Philippines by Koboyashi et al. (1978) indicated that the disease tends to start in the lower needles and spread to the upper crown later on.

Lesions are usually 5-10 mm long (Diekmann et al., 2002); they tend to progress initially from pale-green spots or bands, to a yellowish to yellowish-brown colour, followed by greyish to blackish-brown colour; eventually they coalesce, resulting in complete needle necrosis and eventual needle cast (Ivory and Wingfield, 1986; Ivory, 1987; Braun et al., 2013). Lesions on needles often lead to defoliation and can be especially damaging on young saplings; defoliation often leads to stunted growth and host plant death (Smith et al., 1997). Resulting necrotic needles are always without the reddish tint that is often characteristic of other infections of needles in Pinus species (CABI/EPPO, undated).

Dark-brown stromata fill the stomatal cavities, and numerous fruiting bodies appear as sooty spots on the lesions and, depending on their abundance, give a grey or black discolouration to the bands on the needles (Ivory and Wingfield, 1986). In warm, damp weather, small, grey brush-like tufts of elongate conidia may be just visible on the erumpent stromata (Ivory and Wingfield, 1986). Spermatia may also be extruded in tiny, clear droplets from spermagonia (Ivory, 1987). The distal portions of affected needles die rapidly and become colonized by various saprophytic fungi, whereas the proximal portions remain alive for some time (Ivory, 1987). Ivory and Wingfield (1986) also note that dead foliage may remain on the tree for many months but can be shed during high wind or heavy rain.

List of Symptoms/Signs

Biology and Ecology

Physiology and Phenology

Isolates of M. gibsonii from Asia differ distinctly from African and Jamaican isolates. A third type, which has many similarities with cultures of M. dearnessii, was found on Pinus caribaea in the Philippines (Ivory, 1994). Due to differences in conidial morphology, there are probably three ecotypes: Asia, Africa-Central America and Philippines (Ivory, 1994). M. gibsonii has been found in one or more of its three spore forms (teleomorph, conidia or spermatia) on old seedlings and small trees associated with a more or less severe needle blight of the lower crown, on senescing foliage of older trees and on hanging and dry surface litter (Ivory, 1994). Old foliage is usually infected first, but in severe cases all foliage can be infected (Crous et al., 1990). However, Singh et al. (1988) observed that primary needles appeared to be more susceptible to attack by the pathogen than mature needles.

Conidia germinate between 10 and 35°C (25°C being optimal) and the incubation period varies with environment, but is thought to be 4-6 weeks (Gibson, 1987). According to Suto (1979), heavily infested plants may have a shorter incubation period than those that are moderately infected. In Japan, Ivory (1972) indicated that the disease requires 2-3 days of moist humid conditions for dispersal and infection. Ivory (1987) indicated that under ideal conditions, 3-7 days is sufficient for the production of spores, their dispersal and needle infection by the pathogen. Following infection, stromata form in the stomatal cavities, and bear dense conidiophores. M. gibsonii overwinters as mycelial masses or immature stromata in the tissues of diseased needles (Ivory and Wingfield, 1986). M. gibsonii may also overwinter as latent infections in asymptomatic needles in instances where the needles were infected later in the year (Ivory and Wingfield, 1986; Diekmann et al., 2002; Sullivan, 2010). Symptoms associated with latent infections are often observed the following spring (Ivory and Wingfield, 1986). See also Chupp (1953), Ito (1972), Mulder and Gibson (1972) and Deighton (1987). 

M. gibsonii can be grown in pure culture quite easily, but spores other than spermatia are rarely produced (Ivory, 1994). The spermatial anamorph is known as Asteromella sp.

Environmental Requirements

There is some evidence of edaphic factors (soil pH, organic matter and nutrient content of soil) affecting the incidence of needle blight (Cruz et al., 1984). However, Singh et al. (1988) suggested that low incidence of the disease during winter and dry summer months may be due to low and high temperature respectively. Warm, damp weather provides ideal conditions for conidia production, germination and infection while dry weather often inhibits conidia production (Ivory and Wingfield, 1986). Although conidia are not produced during dry weather, the fungal pathogen may remain dormant in host tissues (Gibson, 1979). It is possible that inocula can survive under unfavourable conditions as mycelium in plant and plant residues (Gibson, 1987). Conidia can remain viable for about one month, but under moist conditions will germinate on needle surfaces within 24-40 hours and penetrate via stomata within a further 2-3 days (Ivory, 1987).

Climate

Means of Movement and Dispersal

Natural Dispersal

The primary infection source of M. gibsonii consists of airborne conidia produced in the spring from needles, and spread in the wind or by rain-splash. It is unlikely that M. gibsonii could spread from Africa and Asia via wind-borne spores. In India, Singh et al. (1988) indicated that there appeared to be a positive relationship between the spread of the disease and high rainfall. Jeger et al. (2017) suggest that due to the major role played by rain water rather than wind in dispersal, the pathogen spreads efficiently only locally, for instance through closely spaced seedlings in nursery beds.

Accidental Introduction

Silvicultural Practices

Infected needle pieces in uncleaned seed or mycorrhizal soil inocula probably enabled the disease to spread to various tropical countries (Ivory, 1987). Diekmann et al. (2002) suggests that the pathogen spreads to new areas on infected nursery stock. Additionally, Ivory (1987) reports dispersal of the pathogen to be less efficient between trees in plantations when compared to nursery settings. This may be due to closer plant spacings in nurseries than in plantations (Singh et al., 1988). Sullivan (2010) also indicated that overhead irrigation may help to spread spores in the field. 

Movement in Trade

The pathogen could enter the EU via plants for planting and other means (uncleaned seeds, cut branches of pine trees, isolated bark, growing media accompanying plants and mycorrhizal soil inocula) (Jeger et al., 2017). Additionally, latent infections of M. gibsonii may present a hazard in view of the long incubation period.

Pathway Causes

Pathway Vectors

Plant Trade

Wood Packaging

Economic Impact

M. gibsonii causes a serious needle blight of both exotic and native pines, particularly at the later nursery stage, and has become a major obstacle to production of pine seedlings (especially P. pinaster, P. thunbergii and P. densiflora) in Japan and Taiwan. The disease causes severe defoliation of susceptible seedlings (Ivory, 1972; Gibson, 1979), slow growth of seedlings, a high cull rate, and in some cases mortality up to 85% (Ivory, 1987). Ito (1972) also reported that during epidemics, there may be up to 100% of seedlings infected. Su-See (1999) has also reported slow growth and a high cull rate for trees up to 5 years old in plantations and forests. Severe defoliation in young plantations of P. radiata occurs in Tanzania, resulting in reduced growth and sometimes even death of the trees (Mulder and Gibson, 1972). The disease is important on P. merkusii and P. caribaea nurseries in Peninsula Malaysia (Ivory, 1975).

Most species develop effective resistance by 2-3 years of age; however, trees which are severely affected initially may remain stunted and severely blighted for many years (Su-See, 1999). Such trees often succumb to competition from healthier trees or weed growth. Resistance in mature trees appears to be dependent on the host affected. For instance, Ivory (1994) observed the pathogen as the cause of significant needle blight on older trees of P. radiata, P. roxburghii and P. canariensis, which do not appear to develop mature plant resistance by two years of age, like most other susceptible species.

Risk and Impact Factors

• Host damage
• Negatively impacts forestry

• Pathogenic

• Highly likely to be transported internationally accidentally

Similarities to Other Species/Conditions

The disease may be confused with dothistroma blight (Mycosphaerella pini), but the pathogen may be distinguished by examination of the conidia (Suto, 1971). There is never a reddish tint to the necrotic needle tissues as may occur with other infections.

Prevention and Control

Due to the variable regulations around (de)registration of pesticides, your national list of registered pesticides or relevant authority should be consulted to determine which products are legally allowed for use in your country when considering chemical control. Pesticides should always be used in a lawful manner, consistent with the product's label.

Cultural Control and Sanitary Methods

Gibson (1987) suggests that losses from the disease may be kept to a minimum by providing unfavourable conditions for spread and infection of conidiae of the pathogen. Some measures for control of M. gibsonii were listed by Ivory (1987). General nursery hygiene measures to minimize the risk of infection are listed below:

• Removal and destruction of all infected pines in and around the nursery
• Cleaning out the nursery between annual production cycles
• Physical separation of young seedlings from older plants where the nursery cycle is >12 months
• Banning transfers of plants between nurseries
• Taking care with mycorrhizal-soil introductions

Additional measures provided by Gibson (1987) for nurseries include:

• Setting up nurseries away from any plantation with diseased trees
• Proper weeding of nursery beds, pathways and nursery surroundings
• Production of robust seedlings by avoiding excessive shading, and hardening of the seedlings as soon as possible before the onset of the rainy season
• Use of temporary rather than permanent nursery sites (rotating nursery sites)
• Wide spacing of nursery plants (e.g. 15 cm)
• Minimizing needle wetting either by protecting against rain if possible or by avoiding overhead irrigation
• Avoiding the planting of infected plants
• Maintaining a proper weeding regime and pruning the lower branches to avoid direct contact with weeds

Singh et al. (1988) observed that in their experimental set-up, in nurseries where infected needles are present in the needle mulch or in mycorrhizal soil, infection appeared on needles within one month of pricking (transplanting). They recommended that:

• Existing nurseries where the pathogen is established are to be avoided in the set up of new nursery sites
• Needles or pine litter collected from infected plantations should not be used as mulch and should be collected and destroyed
• Seeds of exotic pines, when exported, should be completely freed from all seed debris before sowing in nurseries
• Planting schedules should be arranged outside of rainy months

Chemical Control

In nurseries, control can be obtained by using maneb (or mancozeb) or copper-based fungicides applied to the seedlings from the current year and 1-year-old seedlings at 2-weekly intervals during the growing season (Reddy and Pandey, 1973). It is important that all diseased seedlings are removed and burned early in the season when infection occurs. Apply foliar sprays of fungicides at 2-4 week intervals during conditions favourable for the spread of the fungus. Recommended fungicides are benomyl, chlorothalonil, mancozeb, zineb, tridemorph and thiophanate, applied with effective wetting agents (Singh et al., 1983; Ivory, 1987). Copper fungicides have been reported to be effective only in Japanese nurseries (Ivory, 1987).

Host Resistance

Resistant Pinus spp. may have a small portion of their foliage affected, but suffer little growth reduction and no mortality. Resistance has been reported on P. clausa, P. elliotii, P. kesiya, P. rigida and P. patula (Gibson, 1987; EPPO, 2018). Ivory (1987) suggests replacing highly susceptible species with a more resistant, but equally productive species, such as P. elliottii in place of P. massoniana (Ivory, 1987). 

References

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Distribution References

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Contributors

22/10/18 Updated by:

Annika Minott, Caribbean Agricultural Research and Development Institute (CARDI), Cayman Islands

Distribution Maps