Neodiprion sertifer (European pine sawfly)
- Summary of Invasiveness
- Taxonomic Tree
- Notes on Taxonomy and Nomenclature
- Distribution Table
- Habitat List
- Hosts/Species Affected
- Host Plants and Other Plants Affected
- Growth Stages
- List of Symptoms/Signs
- Biology and Ecology
- Natural enemies
- Notes on Natural Enemies
- Means of Movement and Dispersal
- Pathway Vectors
- Plant Trade
- Wood Packaging
- Detection and Inspection
- Similarities to Other Species/Conditions
- Prevention and Control
- Links to Websites
- Distribution Maps
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PicturesTop of page
IdentityTop of page
Preferred Scientific Name
- Neodiprion sertifer (Geoffroy in Fourcroy, 1785)
Preferred Common Name
- European pine sawfly
Other Scientific Names
- Diprion basalis Sakisaka
- Diprion rufum Kl.
- Diprion rufus
- Diprion sertifer Geoffr.
- Lophyrus basalis Matsumura
- Lophyrus piceae Lepeletier
- Lophyrus rufus Latreille; Klug; Panz; Ratz.; Retz.
- Lophyrus sertifer Enslin
- Neodiprion rufus
- Neodiprion sertifer Fourcroy
- Neodiprion sertifera Yano
- Pteronus sertifer Geoffr.
- Tenthredo juniperi Christ
- Tenthredo pectinata rufa Retzius
- Tenthredo pini rufa Villers
- Tenthredo rufa Latreille
- Tenthredo sertifera Geoffroy
International Common Names
- English: fox-coloured sawfly; pine, sawfly, European; pine, sawfly, lesser; sawfly, fox-coloured
- French: diprion du pin; diprion du pin sylvestre; lophyre roux; tenthrede du pin d'ecosse
- Russian: ryzij sosnovyj pililschik
Local Common Names
- Czech Republic: hrebenule rysava; pilatky rysave
- Denmark: rød fyrrehveps
- Estonia: punakas männivaablane
- Finland: ruskea mäntypistiäinen; ruskomäntypistiäinen
- Germany: Buschhornblattwespe, Rote Kiefern-; Kiefernbuschhornblattwespe, Rotgelbe; Rote Kiefernbuschhornblattwespe; Rotgelbe Kiefern-Buschhornblattwespe
- Italy: tenthredine nerastra del pino
- Japan: Matu-no-ki-habati
- Netherlands: Dennebladwesp, rode; rod dennenbladwesp
- Norway: rød furubarveps
- Poland: borecznik rudy
- Sweden: röd tallstekel
- NEODSE (Neodiprion sertifer)
Summary of InvasivenessTop of page N. sertifer is an invasive pest species, of which a good example is its introduction and spread in North America.
Taxonomic TreeTop of page
- Domain: Eukaryota
- Kingdom: Metazoa
- Phylum: Arthropoda
- Subphylum: Uniramia
- Class: Insecta
- Order: Hymenoptera
- Family: Diprionidae
- Genus: Neodiprion
- Species: Neodiprion sertifer
Notes on Taxonomy and NomenclatureTop of page Members of the Diprionidae are commonly called conifer sawflies or diprionids. This is a small family with about 125 species in 11 genera (Smith, 1993; Viitasaari, 2002). According to Ross (1951), N. sertifer was first named by Geoffroy in 1785 as Tenthredo sertifera (Fourcroy, 1785). It has subsequently been included in the genera, Lophyrus, Pteronus and Diprion. The generic designation of Neodiprion dates from 1918 (Rohwer, 1918). Ross (1955) sketched the evolution of the genus based on morphological evidence. Taxonomically, N. sertifer is a distinct species. For a long time it was the only species of the genus in Eurasia, then in the 1980s many new Neodiprion species were described from China (Xiao et al., 1984, 1985). In North America, approximately 37 Neodiprion species are known, some of which form taxonomically difficult species groups and geographic races (Knerer and Atwood, 1973; Smith, 1979; McMillin and Wagner, 1993). According to Ross (1955), the origin of N. sertifer was in North America from where it (or its ancestral form) migrated via the Bering land bridge to northern Asia and then over the whole of northern Eurasia.
DescriptionTop of page Eggs
The eggs are elongate-oval, 1.7-1.8 mm long and 0.3 mm wide. They are white or pale yellow to nearly white (Wilson, 1971). The eggs are laid into slits or egg pockets, which the female cuts with her saw-like ovipositor into the edge of the needle (Ghent, 1959). One egg is laid per pocket and one to 19 (average six to eight) per needle. They are embedded in the needles and are not visible. The distance between the egg pockets is 1-1.75 mm, and they can be seen as a row of semi-lunar yellow spots in the edge of the needle. However, in the spring the pockets open and the swelled eggs become more exposed. The chorion of the parasitized eggs turns blackish or brownish depending on the parasitoid species (Pschorn-Walcher and Eichhorn, 1973).
The newly hatched larva is 3.5 mm long. The male larvae have four feeding instars with shiny black head capsules and the female larvae have five. The mature larva moults to the final non-feeding prepupal or pre-spinning larva, which spins the cocoon. Sometimes both sexes may pass through an extra feeding instar (Lyons, 1964; Juutinen, 1967). The body of L1-L2 instars is a uniform green-grey and longitudinal stripes appear in L3. A mature larva is 18 to 25 mm long and striped. The skin is densely spined. The mid-dorsal surface has a light grey to whitish narrow line, flanked by a broader green-grey line on either side. Each side of the body has two very dark lines that are separated by a thin, whitish spiracular line. The dark lines may be nearly black and tend to break up into spots. A dark spot is present in the upper area on each side of the last abdominal segment. The ventral side is light grey-green. The top of the head of the pre-spinning larva is grey-brown and paler below the eyes. The body has a dark, broken double mid-dorsal line and dark quadrangular pleural markings on a greyish background (Scheidter, 1934; Wilson, 1971).
The cocoon is cylindrical with bluntly rounded ends. It is light to dark golden-brown, tough and finely textured. The male cocoons are smaller (7.5-8.5 mm) than those of the females (8.5-11.0 mm). The cocoons can be sexed by size in low-density populations, but during outbreaks the male and female cocoons may overlap in size due to them starving.
The males are 6-9.5 mm long and black. The ventral part of the abdomen is reddish-brown in total or in part and sometimes the dorsal part is reddish. The legs are yellow-brown and the coxae are black. The antennae are black, bipectinate and have 24-31 segments.
The females are 7-10.5 mm long. The head and body are brown-reddish-yellow and the base of the tibia is whitish, sometimes with dark marks on the mesonotum. The eyes and pronotum are dark. The antennae have 18-28 segments, a dark flagellum and are serrate. The common names of N. sertifer in many languages refer to the colour of the female. The female of the Chinese Neodiprion dailingensis is very similar to that of N. sertifer, but differs from the latter in having the lancet with only eight annuli instead of nine (Xiao et al., 1985).
Some gynandromorphic adults of N. sertifer, exhibiting characteristics of both sexes, have been described (Watson, 1955; Heliövaara et al., 1992; Martini et al., 1999).
DistributionTop of page N. sertifer is the most widely distributed of the diprionids. It is native to Europe and Asia and has been introduced in north-eastern North America. In Europe, it occurs from the Mediterranean to the northern parts of Fennoscandia, from lowlands to mountains up to 2100 m. In Asia, it occurs in Siberia, Korea and Japan, even at a height of 3000 m (Pschorn-Walcher, 1965, 1982).
N. sertifer was probably introduced from Europe to North America. It was first recorded in New Jersey, USA in 1925 and in Ontario, Canada in 1939. It rapidly dispersed and was established as an important pest in south-eastern Canada and in north-eastern USA (Lyons, 1964; Griffiths et al., 1984).
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|
|Armenia||Present||Native||Not invasive||Kolomiets et al., 1972|
|Georgia (Republic of)||Present, few occurrences||Native||Not invasive||Kolomiets et al., 1972|
|Japan||Present||Present based on regional distribution.|
|-Honshu||Widespread||Native||Not invasive||Takeuchi, 1940; Nakamura, 1982; Morimoto and Nakamura, 1989|
|Kazakhstan||Widespread||Native||Not invasive||Gninenko, 1995; Kolomiets et al., 1972|
|Korea, DPR||Present||Native||Not invasive||Takeuchi, 1940|
|Korea, Republic of||Present||Native||Not invasive||Takeuchi, 1940|
|Turkey||Restricted distribution||Native||Not invasive||cuhadar et al., 2000; Yaman et al., 2001|
|Canada||Restricted distribution||Introduced||1939||Invasive||Lyons, 1964; Griffiths et al., 1984|
|-Newfoundland and Labrador||Restricted distribution||Introduced||1974||Invasive||Clark and Singh, 1975; Griffiths et al., 1984; Hall, 1996|
|-Nova Scotia||Restricted distribution||Introduced||1980||Invasive||Magasi, 1981; Griffiths et al., 1984; Hall, 1996|
|-Ontario||Widespread||Introduced||1939||Invasive||Brown, 1940; Raizenne, 1957; Lyons, 1964; Griffiths et al., 1984|
|-Quebec||Widespread||Introduced||1974||Invasive||Martineau & Lavallée, 1975; Griffiths et al., 1984|
|USA||Restricted distribution||Introduced||1925||Invasive||Schaffner, 1939; Lyons, 1964; Coppel and Benjamin, 1965; Wilson, 1971|
|-Connecticut||Restricted distribution||Introduced||Invasive||Lyons, 1964|
|-Delaware||Restricted distribution||Introduced||1979||Invasive||USDA, 1980|
|-Illinois||Restricted distribution||Introduced||Invasive||Benjamin et al., 1955; Lyons, 1964|
|-Indiana||Restricted distribution||Introduced||Invasive||Lyons, 1964|
|-Iowa||Restricted distribution||Introduced||Invasive||Lyons, 1964; Smith, 1979|
|-Kentucky||Present||Rieske et al., 2001|
|-Maine||Restricted distribution||Introduced||Invasive||Smith, 1979|
|-Michigan||Restricted distribution||Introduced||Invasive||Lyons, 1964|
|-Missouri||Restricted distribution||Introduced||Invasive||Lyons, 1964|
|-Montana||Restricted distribution||Introduced||Invasive||Smith, 1979|
|-New Jersey||Widespread||Introduced||1925||Invasive||Schaffner, 1939; Lyons, 1964; Smith, 1979|
|-New York||Restricted distribution||Introduced||Invasive||Lyons, 1964|
|-Ohio||Restricted distribution||Introduced||Invasive||Anon., 1959|
|-Pennsylvania||Restricted distribution||Introduced||Invasive||Lyons, 1964|
|-South Dakota||Restricted distribution||Introduced||Invasive||Lyons and, 1977; Smith, 1979|
|-Wisconsin||Restricted distribution||Introduced||Invasive||Lyons, 1964|
|Austria||Widespread||Native||Not invasive||Niklas and Franz, 1957; Jahn and Sinreich, 1964; Pschorn-Walcher, 1965; Liston, 1995; Krehan, 1999|
|Belarus||Widespread||Native||Not invasive||Kharitonova & Zukhov, 1989; Rywkin, 1957|
|Belgium||Restricted distribution||Native||Not invasive||Breny, 1957; Niklas and Franz, 1957|
|Bulgaria||Widespread||Native||Not invasive||Daskalova, 1970; Tsankov et al., 1980|
|Croatia||Restricted distribution||Native||Not invasive||Opalicki, 1980; Glavas et al., 1999|
|Czech Republic||Widespread||Native||Not invasive||Simandl, 1993; Simandl and Anderbrant, 1995|
|Czechoslovakia (former)||Widespread||Native||Not invasive||Martinek, 1972; Martinek, 1974; Simandl, 1989|
|Denmark||Restricted distribution||Native||Not invasive||Borries, 1895; Bejer, 1989|
|Estonia||Widespread||Native||Not invasive||Kuusik and Kopvillem, 1970; Mihkelson, 1977; Mihkelson, 1980; Voolma, 2000|
|Finland||Widespread||Native||Not invasive||Kangas, 1941; Kangas, 1963; Juutinen, 1967; Viitasaari and Varama, 1987|
|France||Widespread||Native||Not invasive||Joly, 1953; Delplanque et al., 1987|
|Germany||Widespread||Native||Not invasive||Escherich, 1942; Niklas and Franz, 1957|
|Greece||Restricted distribution||Native||Not invasive||Avtzis, 1989; Johansson et al., 2001|
|Hungary||Restricted distribution||Native||Not invasive||Csóka, 1998; Kolonits, 1965; Dulinafka et al., 1983; Liston, 1995|
|Ireland||Present||Native||Not invasive||Pschorn-Walcher, 1965|
|Italy||Restricted distribution||Native||Not invasive||Pollini, 1979; Baronio et al., 1989; Baronio et al., 1997|
|Latvia||Restricted distribution||Native||Not invasive||Eglite and Zarins, 1993; Smits et al., 1996|
|Lithuania||Restricted distribution||Native||Not invasive||Niklas and Franz, 1957; Ziogas and Zolubas, 1998; Zolubas, 1999|
|Luxembourg||Indigenous, localized||Native||Not invasive||Liston, 1995|
|Macedonia||Restricted distribution||Native||Not invasive||Kusevska, 1973|
|Netherlands||Restricted distribution||Native||Not invasive||Hein, 1956|
|Norway||Widespread||Native||Not invasive||AustarÕ et al., 1987; AustarÕ, 1969; AustarÕ, 1971|
|Poland||Widespread||Native||Not invasive||Sitowski, 1925; Gornas, 1968; Kolk and Sierpinski, 1999|
|Portugal||Present||Native||Not invasive||Cardoso Cabral & Morais Figo, 1965|
|Romania||Restricted distribution||Native||Not invasive||Constantineanu, 1983; Mihalache et al., 1998|
|Russian Federation||Widespread||Native||Not invasive||Kolomiets et al., 1972|
|-Central Russia||Widespread||Native||Not invasive||Kolomiets et al., 1972|
|-Eastern Siberia||Present||Native||Not invasive||Kolomiets et al., 1972|
|-Northern Russia||Widespread||Native||Not invasive||Kolomiets et al., 1972|
|-Russian Far East||Present||Native||Not invasive||Kolomiets et al., 1972|
|-Southern Russia||Widespread||Native||Not invasive||Kolomiets et al., 1972|
|-Western Siberia||Widespread||Native||Not invasive||Kolomiets et al., 1972|
|Slovakia||Restricted distribution||Native||Not invasive||Leontovyc et al., 1999|
|Slovenia||Restricted distribution||Native||Not invasive||Jurc, 2001|
|Spain||Restricted distribution||Native||Not invasive||Schönwiese, 1935; Garcia-Viedma and Robredo-Junco, 1962|
|Sweden||Widespread||Native||Not invasive||Forsslund, 1945; Olofsson, 1987|
|Switzerland||Restricted distribution||Native||Not invasive||Niklas and Franz, 1957; Pschorn-Walcher, 1965|
|UK||Restricted distribution||Native||Not invasive||Crooke, 1957; Rivers and Crooke, 1962; Britton, 1985; Liston, 1995|
|-Scotland||Present||Trewhella et al., 2000|
|Ukraine||Widespread||Native||Not invasive||Meshkova, 1998; Meshkova, 1999|
|Yugoslavia (former)||Restricted distribution||Native||Not invasive||Zivojinovic, 1967; Zivojinovic and Sidor, 1975|
Habitat ListTop of page
Hosts/Species AffectedTop of page N. sertifer attacks most species of the two-needled pines, but also some soft pines. In Europe, the main host trees are Pinus sylvestris, Pinus nigra, Pinus montana [Pinus mugo] and Pinus cembra. Heavy feeding has also been found on exotic pines such as Pinus strobus, Pinus banksiana, Pinus resinosa, Pinus radiata and some Japanese pines (Pschorn-Walcher, 1965, 1982). The Canadian Pinus contorta is sometimes more heavily attacked than P. sylvestris (Rivers and Crooke, 1962; Delplanque et al., 1987; Olofsson, 1988b, 1989).
In Siberia, N. sertifer occurs on Pinus sibirica (Kolomiets et al., 1972). In Japan, the host plants in the lowlands are Pinus densiflora, Pinus thunbergii and P. resinosa, and in the mountains, Pinus pumila (Suzuki, 1964; Nakamura, 1982; Morimoto and Nakamura, 1989).
In North America, the favoured hosts are the exotic P. sylvestris, P. densiflora and P. mugo, and the native P. resinosa, P. banksiana, Pinus ponderosa, P. radiata, Pinus echinata, Pinus rigida, P. strobus and many others (Craighead, 1950; Lyons, 1964). Because the range of hosts that are accepted in laboratory feeding tests is often greater than that observed in the field, it is assumed that the female oviposits on a narrower range of species than are potentially suitable for larval growth (Heitland and Pschorn-Walcher, 1993). Pinus peuce is unsuitable, and P. strobus is unfavourable for oviposition. The eggs are lost on Pinus palustris and on P. rigida when the needles die and drop (Benjamin et al., 1955; Wilson, 1971). According to Lyons (1964), no oviposition has been seen in the field on soft pines in Europe or North America. The susceptibility of a host tree to pest attack may also be dependent on the provenance or variety (Wright et al., 1967; Olofsson, 1989; Eliason and McCullough, 1997; Trewhella et al., 1997, 2000). Occasional feeding on spruces (Picea spp.) has been recorded where these grow in close proximity to pines (Gäbler, 1940; Forsslund, 1945; Crooke, 1957; Rivers and Crooke, 1962).
Host Plants and Other Plants AffectedTop of page
|Picea abies (common spruce)||Pinaceae||Other|
|Pinus banksiana (jack pine)||Pinaceae||Main|
|Pinus cembra (arolla pine)||Pinaceae||Main|
|Pinus contorta (lodgepole pine)||Pinaceae||Main|
|Pinus densiflora (Japanese umbrella pine)||Pinaceae||Main|
|Pinus echinata (shortleaf pine)||Pinaceae||Other|
|Pinus mugo (mountain pine)||Pinaceae||Main|
|Pinus nigra (black pine)||Pinaceae||Main|
|Pinus nigra austriaca||Pinaceae||Main|
|Pinus ponderosa (ponderosa pine)||Pinaceae||Main|
|Pinus pumila (Dwarf Siberian pine)||Pinaceae||Main|
|Pinus pungens (tabel Mountain pine)||Pinaceae||Other|
|Pinus radiata (radiata pine)||Pinaceae||Main|
|Pinus resinosa (red pine)||Pinaceae||Main|
|Pinus rigida (pitch pine)||Pinaceae||Other|
|Pinus sibirica (Siberian stone pine)||Pinaceae||Main|
|Pinus strobus (eastern white pine)||Pinaceae||Main|
|Pinus sylvestris (Scots pine)||Pinaceae||Main|
|Pinus thunbergii (Japanese black pine)||Pinaceae||Main|
Growth StagesTop of page Vegetative growing stage
SymptomsTop of page Large larval colonies feed on needles from spring to mid-summer. The older needles are fed on down to the needle sheath and the newly developing needles are mainly untouched. The older larvae may also consume bark from the older, thin branches. Heavy outbreaks may result in the complete removal of the old foliage. Normally the trees recover later in the summer when the new shoots and needles reach their full size. Lyons (1964) has published illustrative photos of the stages in the defoliation of one tree. Heavy defoliations in subsequent years may kill buds and twigs. Although extensive mortality seldom occurs, repeated defoliation weakens the trees and increases their susceptibility to attack from secondary pests (e.g. bark beetles).
The frass of diprionid larvae can be easily distinguished from lepidopteran frass (Escherich, 1942). The shape of the diprionid excrement pellets is rhomboid. They are initially green and later turn brownish. During heavy larval infestations, a light patter caused by the dropping frass can be heard in quiet forests and a thin layer of frass can be detected on the ground.
List of Symptoms/SignsTop of page
|Leaves / external feeding|
|Leaves / frass visible|
|Stems / external feeding|
|Stems / visible frass|
|Whole plant / plant dead; dieback|
Biology and EcologyTop of page The vast amount of literature from around the world on the biology and ecology of N. sertifer has been reviewed by Escherich (1942), Niklas and Franz (1957), Lyons (1964), Kolomiets et al. (1972), Martinek (1972, 1974), Pschorn-Walcher (1965, 1982) and Baronio et al. (1997). Wagner and Raffa (1993) have dealt with many aspects of the sawfly's life history.
N. sertifer is univoltine. The life cycle is fairly uniform throughout the whole distribution area, although temporal differences exist depending on latitude and altitude. The adults emerge from cocoons in the autumn, from August to November. The sex ratio is mostly female-biased (Craig and Mopper, 1993). The female deposits her eggs in the needles of a current year shoot and an egg cluster is formed. The number of eggs per female ranges from 30 to 140 eggs. N. sertifer is arrhenotokous i.e. unfertilized eggs produce haploid males.
Embryonic development begins in the autumn but is suspended during the winter. According to Niklas and Franz (1957), development continues throughout the winter and is retarded only by low temperatures. Breny (1956) considers the cessation as a pseudo-diapause resulting from dehydration and low availability of water from the needle tissues. According to Wallace and Sullivan (1973), a true diapause is involved. The overwintering eggs are cold-hardy and average freezing points ranging from -30°C to -37°C have been recorded (Sullivan, 1965; Juutinen, 1967; Kuusik and Kopvillem, 1970, Austarå, 1971).
The larvae hatch in the Spring, from April to the beginning of June. They feed gregariously on 1 year or older needles, leaving the new growing foliage mainly untouched. From June to July the non-feeding, pre-spinning larvae spin cocoons in the litter and humus layer in the ground and only occasionally in the foliage. The eonymphs in cocoons enter a Summer diapause, which is longest in the southern and lowland populations and shortest in the northern and alpine populations. Also, insects that spin cocoons early in the season undergo a longer eonymphal diapause than those spinning later (Lyons and Griffiths, 1962). The diapause may last for 1 year or more. The incidence of prolonged diapause increases with both latitude and altitude in Europe. It is 0-10% in the southern lowland areas and 50-100% in the northern countries (Juutinen, 1967; Mihkelson, 1977; Pschorn-Walcher, 1982). In the high mountains and in the north, a life cycle of 2 years is usual (Okutani and Ito, 1957; Austarå, 1969; Pschorn-Walcher, 1970; Martinek, 1972).
The population density of N. sertifer fluctuates in wide ranges. Outbreaks, resulting in the heavy defoliation of pine stands, are known throughout the whole distribution area. The outbreaks rise to a peak in 1 to 3 years, and collapse abruptly. Niklas and Franz (1957), Martinek (1972) and Pschorn-Walcher (1965, 1982) provided lists of the outbreaks recorded in Europe.
The outbreaks in Europe tend to occur synchronously over large areas at intervals of 6 to 13 years. The extent of the outbreaks varies considerably, from a few to 200 000 ha. The young pine stands on poor soils and with low water tables are most affected. Also the mature pine stands in Fennoscandia are frequently attacked.
In North America, most outbreaks have been unsynchronized, independent events initiated by the discovery of young, previously uninfested pine forests containing few natural control agents. Also the population levels have reached higher levels than in Europe (Lyons, 1964, 1977).
There seems little doubt that the inception of outbreaks is climatically regulated, either directly or indirectly, and that dry, hot weather is the critical factor. The factors responsible for the decline of an outbreak may vary in their effect from place to place. The most frequently recorded regulators are viral disease, parasitoids and predators.
There are four major hypotheses that tend to explain patterns of sawfly dynamics. The regulatory factors are microparasites (virus), parasitoids, generalist cocoon predators and plant defence mechanisms (Hanski, 1987). The dynamics of N. sertifer can be best explained by cocoon predation by shrews and small omnivorous mammals (Hanski, 1987, 1990; Larsson et al., 1993).
Natural enemiesTop of page
|Natural enemy||Type||Life stages||Specificity||References||Biological control in||Biological control on|
|Bacillus thuringiensis kurstaki||Pathogen|
|Bacillus thuringiensis thuringiensis||Pathogen|
|Dahlbominus fuscipennis||Parasite||Pupae||Canada; USA||Pinus|
|Exenterus abruptorius||Parasite||Larvae||Canada; USA||Pinus|
|Lamachus eques||Parasite||Larvae||Canada; USA||Pinus|
|Lophyroplectus luteator||Parasite||Larvae||Canada; Ontario; USA||Pinus|
|Pleolophus basizonus||Parasite||Pupae||Newfoundland; Ontario; USA||Pinus|
Notes on Natural EnemiesTop of page N. sertifer is attacked by several hymenopterous and dipterous parasitoids, and many predators including ants, bugs, beetles, lacewings, spiders, small mammals and birds. Pathogenic fungi, bacteria and a species-specific nuclear polyhedrosis virus (NsNPV) also attack it.
The diprionid larvae have special defence strategies against predators and parasitoids (Codella and Raffa, 1993) and being gregarious is one of these. The larvae also exhibit various alarm reactions and defensive displays. Prop (1960) distinguished three types of larval displays: U-bend; jerking; and stretching. The larvae possess a pair of oesophageal diverticulae with resinous contents. In response to harassment, the larvae regurgitate a droplet of the resinous liquid. This behaviour has been shown to repel ants, pentatomid bugs, spiders and birds, as well as hymenopteran and dipteran parasitoids.
There are a large number of parasitoid records from N. sertifer in the literature. However, according to Oehlke (1965) and Pschorn-Walcher (1965), a relatively high proportion of these records, particularly the older ones, are unreliable. The main sources of error have been: the mass-collection and mass-rearing of larvae and cocoons; the inadequate sample-size and faulty timing of sampling; and the false identification of the parasitoids or the hosts. The list of parasitoids presented here is based mainly on the reviews and articles of Niklas and Franz (1957), Rywkin (1957), Finlayson and Finlayson (1958), Finlayson (1960), Lyons (1964), Oehlke (1965), Kolomiets et al. (1972), Martinek (1972, 1974), and Pschorn-Walcher (1965, 1967, 1982).
Four chalcidoid egg parasitoids are known from N. sertifer in Europe (Pschorn-Walcher and Eichhorn, 1973; Pschorn-Walcher, 1982). The tetracampid Dipriocampe diprioni is strictly primary, the eulophids Closterocerus ruforum and C. formosus act as primary and hyperparasitoids, even as autoparasitoids, while the eulophid Baryscapus (Tetrastichus) oophagus is a rare hyperparasitoid. As primary egg parasitoids, D. diprioni and C. ruforum are the dominant species. In Europe, egg parasitism (including host-feeding) is generally between 5 and 50%, but during the break-down phase of the host population, much higher values have been recorded, such as from 90-95% (Juutinen, 1967; Pschorn-Walcher and Eichhorn, 1973; Juutinen and Varama, 1986). The egg parasitoids of N. sertifer do not occur in North America, and attempts to introduce and colonize them from Europe have been unsuccessful (McGugan and Coppel, 1962; Griffiths et al., 1984).
The most important parasitoids of N. sertifer in Europe are the larval parasitoids, Lamachus eques, Lophyroplectus luteator, Exenterus abruptorius, and the cocoon parasitoids Pleolophus basizonus (Ichneumonidae) and Dahlbominus fuscipennis (Eulophidae). Of minor importance are Exenterus amictorius, Exenterus adspersus, Synomelix scutulata (Ichneumonidae), and Drino inconspicua and Blondelia inclusa (Tachinidae) as larval parasitoids, and the ichneumonid, Agrothereutes adustus as a cocoon parasitoid (Pschorn-Walcher, 1965, 1967, 1982).
The parasitoid complex of N. sertifer in North America consists of about 30 native primary parasitoids, some native hyperparasitoids, and six introduced and established European parasitoids (Lyons 1964; Griffiths et al., 1984). The important native species are: the larval parasitoids, Exenterus nigrifrons [Exenterus canadensis], Exenterus affinis (Ichneumonidae), Spathimeigenia spinigera [Vibrissina spinigera], Diplostichus hamatus (Tachinidae); and the cocoon parasitoids, Pleolophus indistinctus, Endasys subclavatus, Delomerista diprionis, Mastrus argeae (Ichneumonidae), Tritneptis spp. (Pteromalidae), Eupelmus vesicularis [Eupelmus vesicularis] (Eupelmidae) and Hemipenthes sinuosa [Villa sinuosa] (Bombyliidae). The established European species are the ichneumonids, E. amictorius, E. abruptorius, P. basizonus, L. luteator and the eulophid, D. fuscipennis, and the tachinid, Drino bohemica (Griffiths et al., 1984). Also the introduced torymid, Monodontomerus dentipes, has been encountered a couple of times (Finlayson, 1960; Lyons 1964).
Mortality of N. sertifer due to parasitism varies greatly (10-90%) by year and locality. The parasitoids, along with the nuclear polyhedrosis virus, play an important role especially during the break-down phase of the outbreak. They often cause the abrupt collapse of the host population (Niklas and Franz, 1957; Lyons, 1964; Pschorn-Walcher, 1965, 1982, 1988).
True hyperparasitoids do not play a very significant role among the parasitoids, but multi-parasitism in the form of competition between larval and cocoon parasitoids may be increased when the total parasitism is high (Pschorn-Walcher, 1973).
The literature on predation upon N. sertifer has been reviewed by Niklas and Franz (1957), Lyons (1964), Kolomiets et al. (1972), Martinek (1972) and Pschorn-Walcher (1965, 1982). Tits (Parus spp.), lacewings (Chrysopa ventralis), and pentatomid, reduviid and nabid bugs (Picromerus bidens, Euschistus tristigmatus, Podisus spp., Rhinocoris annulatus [Rhynocoris annulatus], Zelus sp. and Nabis sp.) have been reported to occasionally prey on the eggs of N. sertifer.
Due to the effective defence mechanisms and the unpalatability of the larvae to predators, mortality of the larval stages caused by the predators is generally low. Birds, ants (Formica spp.), predatory bugs, coccinellids, neuropterans and spiders have been reported to occasionally feed on the larvae. However, Schwerdtfeger (1936) reported that ants of the Formica rufa-group were one of the principal factors causing sawfly mortality in the Prussian plain. According to Bruns and Schrader (1955), no cocoons with living N. sertifer could be found within 30 m of F. rufa nests. In Sweden, Olofsson (1992) reported a high mortality of the larvae within 40 m of a Formica polyctena nest.
Cocoon predation plays an important role in the population dynamics of N. sertifer and of the other diprionid species. The larvae of the family Elateridae (Coleoptera) destroy cocoons in the ground, and birds (mainly Parus spp.) prey effectively on the cocoons occasionally spun in the foliage. However, the most effective cocoon predators are small mammals such as shrews, mice and voles (Soricidae, Microtidae). Holling (1959), Hanski and Parviainen (1985) and Hanski (1987, 1990) studied small mammal predation of N. sertifer. According to Hanski (1990), small mammals frequently consume 80% or more of the cocoons in endemic sawfly populations when the density of cocoons in coniferous forests in Fennoscandia is less than 10 000 per ha. In contrast, during severe outbreaks when there are several million cocoons per ha, only a small fraction is destroyed by small mammals.
A lethal species-specific nuclear polyhedrosis virus disease (NsNPV) frequently infects the larvae of N. sertifer. The disease is caused by Borrelinavirus diprionis [NeseNPV], which belongs to the baculoviridae (Cunningham and Entwistle, 1981). It occurs widely in nature and is often one of the main factors causing the abrupt collapse of outbreaks (Lyons, 1964; Pschorn-Walcher, 1982). The larvae become infected either by ingesting the polyhedral inclusion bodies (PIB) with food, or via an infected parent. The parasitoids, predators and scavengers may act as transmission agents because the virus stays infectious in their faeces (Olofsson, 1988a). NsNPV has been widely used in the biological control of N. sertifer.
Means of Movement and DispersalTop of page Natural dispersal (non-biotic)
N. sertifer larvae normally pupate (spin cocoons) in the litter and soil, and occasionally in the foliage or branches. During outbreaks there is a risk that the larvae pupate in forests in containers, vehicles, camping equipment etc., and thus get transported into new areas.
N. sertifer is an introduced, dispersing species in North America. Some new infestations did not result from natural dispersal but from the introduction of infested nursery stock (Griffiths et al., 1971).
Pathway VectorsTop of page
Plant TradeTop of page
|Plant parts liable to carry the pest in trade/transport||Pest stages||Borne internally||Borne externally||Visibility of pest or symptoms|
|Bark||pupae||Yes||Pest or symptoms usually visible to the naked eye|
|Growing medium accompanying plants||pupae||Yes||Pest or symptoms usually visible to the naked eye|
|Leaves||eggs; larvae; pupae||Yes||Pest or symptoms usually visible to the naked eye|
|Stems (above ground)/Shoots/Trunks/Branches||larvae; pupae||Yes||Pest or symptoms usually visible to the naked eye|
|Plant parts not known to carry the pest in trade/transport|
|Fruits (inc. pods)|
|True seeds (inc. grain)|
Wood PackagingTop of page
|Wood Packaging liable to carry the pest in trade/transport||Timber type||Used as packing|
|Solid wood packing material with bark||Yes|
|Solid wood packing material without bark||Yes|
|Wood Packaging not known to carry the pest in trade/transport|
|Loose wood packing material|
|Processed or treated wood|
ImpactTop of page N. sertifer is a univoltine, early season defoliator. The larvae feed on older needles and leave the new growing needles mainly untouched. Hence tree mortality is normally modest, even after heavy infestations. In North American Christmas tree plantations, N. sertifer has been a serious problem and control applications have been widely used, because even a light defoliation renders the trees unmarketable (Lyons, 1964; Wilson, 1971).
Relatively little is known about the effect of defoliation on tree growth. In Michigan, 20, 65, 85, and 100% defoliation before new foliage developed, caused 14, 23, 37, and 63% loss in terminal shoot elongation and 18, 47, 53, and 71% reduction in radial increment, respectively. The 100% defoliation was simulated and trees survived 3 years of complete defoliation (Wilson, 1966).
In Hungary, heavy defoliation caused 30-45% reduction in annual growth (Kolonits, 1965). In Sweden, Forsslund (1945) found the height growth loss to vary between 25% and 60% in young Scots pine stands defoliated for two consecutive years. Eklund (1964) recorded a reduction of 52% in the diameter growth of old pine stands due to defoliation. The measurements were made 1-2 years after the infestation, therefore they do not cover the whole recovery period, which may be as long as 9 years (Austarå et al., 1987).
In Finland, the mortality of Scots pine after 1 years defoliation was recorded as approximately 4%, and a reduction in the volume increment for the following 5 years was 20% corresponding to one normal annual increment (Juutinen, 1967; Tiihonen, 1970). Most of the killed trees were suppressed or weakened before the infestation. In Norway, a heavy defoliation of Scots pine for 2 years in 90-120-year-old forests caused a volume loss of 33% during 9 years. This loss corresponds to three normal annual increments (Austarå et al., 1987). The economic consequences of the growth loss and mortality have been calculated by Austarå et al. (1987), and Lyytikäinen-Saarenmaa and Tomppo (2002).
Detection and InspectionTop of page The overwintering egg clusters, which appear as rows of yellow spots (egg pockets) in the edges of the needles in the current year's shoots, can be detected from the Autumn to the Spring by careful inspection. The signs of defoliation reveal the feeding larval colonies. The presence of cocoons in the soil can be checked. Pheromone traps containing the female sexual pheromone can be set to catch male insects at the time of their emergence from the cocoons in Autumn.
Similarities to Other Species/ConditionsTop of page The way of feeding and the symptoms of defoliation are very similar between many pine sawfly species. In Europe, N. sertifer is the only diprionid species that overwinters in the egg-stage and defoliates pines in the Spring or early Summer. For proper identification of the pest species, samples of the egg-bearing needles, larvae, larval skins, adults or cocoons should be taken.
Prevention and ControlTop of page
One of the best examples of the successful development of biological control methods in forest protection is the nuclear polyhedrosis virus of N. sertifer (NsNPV). The virus was discovered in Germany (Escherich, 1913). Bird (1953, 1955, 1961) developed a supply of the virus from a few dead larvae of N. sertifer from Sweden in 1949. This was used for experimental purposes and eventually for release as a control agent. Due to the intensive studies in America and Europe (e.g. Bird and Whalen, 1953; Franz and Niklas, 1954; Benjamin et al., 1955; Krieg, 1955; Rivers and Crooke, 1962; Nuorteva, 1972; Donaubauer, 1973; Cunningham and Entwistle, 1981; Juutinen, 1982; Podgwaite et al., 1984), the NsNPV became the most widely used baculovirus in several countries (Pschorn-Walcher, 1982; Morris et al., 1986). It has been registered under different names in the USA, UK, Finland and the Czech Republic. In Ontario, N. sertifer was a major pest of Christmas tree plantations but since the 1970s it has only been a minor pest due to the use of the NsNPV and parasitoid introductions (Griffiths et al., 1984; Morris et al., 1986).
The incubation time of NsNPV is 1-2 weeks. The virus should be applied on early instars (L1, L2) to be effective and cause over 90% mortality. According to the critical review by Olofsson (1988a), the virus plays a minor role in the population dynamics of N. sertifer; often a smaller role than other biotic mortality factors. Therefore, virus treatment should be resorted to in exceptional cases only, when there are specific indications that a treatment is required. For example, when severe damage by secondary forest insects, e.g. bark beetles, can be expected. See van Frankenhuyzen (2002) for an assesssment of the successful control of N. sertifer in Canada with the NPV.
In the laboratory and field tests performed by Heimpel and Angus (1963), Bacillus cereus was pathogenic (25% mortality) to larvae of N. sertifer, but the Bacillus thuringiensis var. sotto Ishiwata toxin was ineffective (Angus, 1956). Sezen et al. (2001) investigated the insecticidal potential of the bacterium, Serratia marcescens Bn10, which was isolated from Balaninus nucum. Mortality of the N. sertifer larvae was 88% within 5 days.
The rhabditid nematode, Heterorhabditis heliothidis, caused 100% mortality of N. sertifer larvae within 120 hours at 15°C in the laboratory. The intact cocoons were not susceptible and only 11% mortality in 168 hours was observed. However, the pupae that were dissected out of the cocoons were all parasitized in 48 hours (Finney and Bennett, 1983).
In small ornamental trees or in pine plantations of small acreage, N. sertifer can be controlled simply by hand-picking and destroying the larval colonies. A mere knocking down of the larvae is useless, because the larvae climb back into the trees within a few hours (Teräs, 1982).
Different pine species, provenances and varieties have been shown to vary in their susceptibility to N. sertifer (refer to Notes on host range). Where possible, less susceptible alternatives should be used in forestry programs.
The larvae of N. sertifer normally reject the young foliage of the Scots pines. This feeding pattern is mainly caused by a deterrent compound identified as 13-keto-8(14)-podocarpen-18-oic acid (Niemelä et al., 1982). This substance was mainly found in the young foliage of pines, and the highest concentrations were found in the trees whose needles had been badly damaged by the N. sertifer larvae.
Like many other species of diprionids, the males of N. sertifer are strongly attracted by the female sex pheromones. The investigation of the diprionid sex pheromones started in the late 1950s (Coppel et al., 1960), and the first pheromone was identified 16 years later (Jewett et al., 1976). Anderbrant (1993, 1999) thoroughly reviewed the literature since then and up to 1999.
The inactive precursor of the N. sertifer pheromone is 3,7-dimethyl-2-pentadecanol (diprionol) and the active compound is (2S,3S,7S)-diprionyl acetate or propionate, which has proved to be attractive in Europe, North America and Japan (Anderbrant, 1993, 1999). Traps baited with synthetic pheromone can be used for many purposes. The flight period can be determined (Simandl, 1993; Schedl, 1994), the abiotic factors important for flight can be investigated (Jönsson and Anderbrant, 1993), and the behaviour of the sawflies can be studied (Östrand, 2001). The first attempts to use pheromone monitoring traps for determining population densities or population trends of N. sertifer showed either significant relationships (Baldassari et al., 2000), only weak relationships (Lyytikäinen-Saarenmaa et al., 1999, 2001) or no significant correlation (Herz et al., 2000) between the trap catch and the sawfly density or defoliation.
The use of sex pheromones in the mating disruption technique to control populations resulted in the suppression of the N. sertifer population in isolated pine stands in Italy (Martini et al., 2002). It was ineffective in larger areas of pine forests in Sweden (Anderbrant et al., 1995).
The diprionid sawflies are susceptible to most stomach and contact poisons, and a vast array of insecticides have been utilized in abatement programmes (Lyons, 1964; Coppel and Benjamin, 1965, Pschorn-Walcher, 1982). Early control programmes mainly utilized lead or calcium arsenate (Hamilton, 1943), a hazardous chemical that is not recommended.
Currently chlorinated hydrocarbons have been replaced with organophosphorous insecticides and pyrethroids, which have proved to be effective against the diprionid larvae (Wallner, 1968; Malinowski, 1995; Safronov, 1996). In North America, malathion has been used to control N. sertifer (Wilson, 1971).
The insect growth regulators, acylurea insecticides, have been actively developed and tested since the 1970s. They interfere with the normal formation of the cuticle and the deposition of chitin, and are environmentally safer than many early pesticides. Skatulla (1975) reported that aerial spraying of diflubenzuron caused 100% mortality of the N. sertifer larvae in 17 days, and no adverse side effects were observed. Ultra low volume aerial application of diflubenzuron resulted in 98-100% mortality of the N. sertifer larvae (Adomas, 1999). Malinowski (1995) demonstrated that flufenoxuron, novaluron and teflubenzuron were more active than diflubenzuron and triflumuron against the diprionid larvae. According to Skatulla (1975), diflubenzuron remains effective for 4 months. Schwenke (1979) discussed the possible environmental side effects of diflubenzuron applications.
Azadirachtin is a natural insecticide from the neem tree (Azadirachta indica) and strongly reduced the feeding activity and body weight of N. sertifer larvae under laboratory conditions. 100% mortality was achieved. For experimental treatment in the field, azadirachtin concentrations at levels from 0.001-0.00001% were proposed (Malinowski, 2002).
According to All and Benjamin (1976), certain insect antifeedant chemicals have the potential to reduce Neodiprion larval feeding. When larvae were subjected to field sprays, feeding, development and colony integrity were often disrupted. When colonies disbanded, the solitary larvae were more vulnerable to entomophages. Thus the antifeedants could be important components in an integrated control program to maximize the effectiveness of the natural control agents.
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