Diabrotica virgifera virgifera (western corn rootworm)
- Summary of Invasiveness
- Taxonomic Tree
- Notes on Taxonomy and Nomenclature
- Distribution Table
- History of Introduction and Spread
- Risk of Introduction
- Habitat List
- Hosts/Species Affected
- Host Plants and Other Plants Affected
- Growth Stages
- List of Symptoms/Signs
- Species Vectored
- Biology and Ecology
- Natural enemies
- Notes on Natural Enemies
- Means of Movement and Dispersal
- Plant Trade
- Impact Summary
- Environmental Impact
- Impact: Biodiversity
- Social Impact
- 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
- Diabrotica virgifera virgifera LeConte
Preferred Common Name
- western corn rootworm
Other Scientific Names
- Diabrotica filicornis Horn
- Diabrotica virgifera LeConte
- Diabrotica virgifera var. filicornis Gillette
International Common Names
- English: Colorado corn rootworm; corn rootworm, Colorado; corn rootworm, western
- Spanish: diabrotica del maiz; gusano de la raiz del maiz; gusano del raiz
- French: chrysomèle des racines du maïs; chrysomèle des racines du maïs
Local Common Names
- Germany: Westlicher Maiswurzelbohrer
- DIABVI (Diabrotica virgifera)
Summary of InvasivenessTop of page
D. virgifera virgifera is thought to originate from Central America (Krysan and Smith, 1987). In the twentieth century, Diabrotica virgifera virgifera became a major pest of maize in North America as maize growing areas increased (Krysan and Miller, 1986; Tallamy et al., 2005). The practice of continuous maize growing was largely responsible for the expanding range of D. virgifera virgifera.
D. virgifera virgifera was accidentally introduced at least three times from North America into Europe between the late 1980s and the early 2000s (Miller et al., 2005). There is no information about the pathways of introductions, or the process of its entry, or about its period of adaptation and establishment. Larvae-induced damage was recorded for the first time near Belgrade airport in former Yugoslavia in 1992 (Baca, 1994). The species became invasive and quickly spread throughout Central Europe in the 1990s and early 2000s. Although the larvae move relatively small distances, the adults fly to maize fields and are able to actively migrate several kilometres per year (Coats et al., 1986; Isard et al., 2000, Kiss et al., 2005b). In the laboratory, adults were observed to fly up to a maximum of 24 km on flight mills (Coats et al., 1986). Under field conditions, adults may additionally be carried by weather features such as cold fronts (Grant and Seevers, 1989) and thunderstorms (Onstad et al., 1999). Conclusively, the large-scale spread of D. virgifera virgifera varies a great deal between years and regions, sometimes reaching up to 60 to 80 km per year (Edwards et al., 1999; Baufeld and Enzian, 2005). Recently, several new isolated satellite introductions were reported around Paris (France), Basel (France and Switzerland), Amsterdam (The Netherlands), in Belgium and around London (UK) (Kiss et al., 2005b). All maize-growing areas of Europe, and in the long-term also of Asia, are at risk (Baufeld and Enzian, 2005) because this pest is likely to survive and develop wherever maize is grown. Its adaptability to different climatic conditions is also reflected by its broad distribution area from Northern Mexico, throughout the USA and up to Canada (Krysan and Miller, 1986).
Taxonomic TreeTop of page
- Domain: Eukaryota
- Kingdom: Metazoa
- Phylum: Arthropoda
- Subphylum: Uniramia
- Class: Insecta
- Order: Coleoptera
- Family: Chrysomelidae
- Genus: Diabrotica
- Species: Diabrotica virgifera virgifera
Notes on Taxonomy and NomenclatureTop of page
First described as Diabrotica virgifera by LeConte (1868), the taxonomic history of this species has seen three changes: Diabrotica filicornis (Horn, 1893), Diabrotica virgifera var. filicornis (Gillette, 1910) and Diabrotica virgifera virgifera (Krysan et al., 1980). For a complete description of D. virgifera virgifera, refer to Krysan and Smith (1987).
DescriptionTop of page Eggs
Embryonic stage at diapause consists of a flattened oval egg, ca 500 µm long, consisting of undifferentiated cells representing the embryonic primordium and the amnion.
Three larval instars. Larvae are slender, white to pale yellow, with a yellowish-brown head capsule and a brownish plate on the last abdominal segment. The third instar is about 10 mm long (Chiang, 1973).
Pupae are naked and white, turning brownish before adult emergence, and are found in earthen cells in the soil near plant roots.
Adults are elongate, ca 5 mm (range 4.4-6.8 mm), greenish-yellow, with parallel yellow humeral plicae, with distinct piceous elytral vittae; lateral margin widest medially. The pronotum is shiny and yellow. The elytra are straw yellow, each with one humeral and one sutural vitta ending near the apex; humeral plicae are distinct. There are three discal sulci. The scutellum is buff. The head is shiny; the vertex is amber; the labrum and clypeus are piceous. Antennae of males are about 4.2 mm and longer than those of females. Segment three is 1.5 times longer than segment two. Segments two and three combined equal the length of segment four. The sterna are straw yellow, the episternites tawny and piceous on sutures. Legs are yellow to tawny, with the outer edge of femora piceous. The abdomen is yellow. The widths of the piceous humeral and sutural elytral vittae are highly variable (Krysan et al., 1980; Krysan and Smith, 1987).
DistributionTop of page
D. virgifera virgifera, which originates from the USA, was recorded for the first time in the former Yugoslavia in 1992 and has quickly spread to become a threat to maize in Europe (Gerginov and Tomov, 1995). The practice of growing maize continuously is largely responsible for the expanding range of D. virgifera virgifera in North America.
See also Edwards (2006), CABI/EPPO (1998, No. 63) and http://www.eppo.org/QUARANTINE/Diabrotica_virgifera/diabrotica_virgifera.htm [EPPO, 2006 #9321].
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.Last updated: 23 Apr 2020
|Continent/Country/Region||Distribution||Last Reported||Origin||First Reported||Invasive||Reference||Notes|
|Albania||Present, Localized||EPPO (2020); CABI/EPPO (2012)|
|Austria||Present, Localized||Introduced||2002||Invasive||Kiss et al. (2005); CABI/EPPO (2012); EPPO (2020)|
|Belarus||Present, Few occurrences||CABI/EPPO (2012); EPPO (2020)|
|Belgium||Present, Transient under eradication||Introduced||2003||Invasive||Kiss et al. (2005); IPPC (2010); CABI/EPPO (2012); EPPO (2020)|
|Bosnia and Herzegovina||Present, Widespread||Introduced||1997||Invasive||Kiss et al. (2005); CABI/EPPO (2012); EPPO (2020)|
|Bulgaria||Present, Widespread||Introduced||1998||Invasive||Kiss et al. (2005); CABI/EPPO (2012); EPPO (2020)|
|Croatia||Present, Widespread||Introduced||1995||Invasive||Kiss et al. (2005); CABI/EPPO (2012); EPPO (2020)|
|Czechia||Present, Few occurrences||Introduced||2002||Invasive||Kiss et al. (2005); CABI/EPPO (2012); EPPO (2020)|
|Denmark||Absent, Confirmed absent by survey||EPPO (2020)|
|Estonia||Absent, Confirmed absent by survey||EPPO (2020)|
|France||Present, Localized||Introduced||Invasive||Kiss et al. (2005); CABI/EPPO (2012); EPPO (2020)||First reported: 2002 and 2003|
|Germany||Present, Transient under eradication||CABI/EPPO (2012); Dicke et al. (2014); EPPO (2020)|
|Greece||Present, Localized||CABI/EPPO (2012); Michaelakis et al. (2010); EPPO (2020)|
|Hungary||Present, Widespread||Introduced||1995||Invasive||Kiss et al. (2005); CABI/EPPO (2012); EPPO (2020)|
|Italy||Present, Localized||Introduced||Invasive||Kiss et al. (2005); CABI/EPPO (2012); EPPO (2020)||First reported: 1998, 2000 and 2002|
|Montenegro||Present, Localized||EPPO (2020); CABI/EPPO (2012)|
|Netherlands||Absent, Eradicated||Kiss et al. (2005); IPPC (2006); CABI/EPPO (2012); EPPO (2020); CABI (Undated)||First reported: 2003 and 2005|
|Poland||Present, Localized||Introduced||2005||Invasive||Yardim et al. (2006); CABI/EPPO (2012); EPPO (2020)|
|Romania||Present, Widespread||Introduced||1996||Invasive||Kiss et al. (2005); CABI/EPPO (2012); EPPO (2020)|
|Russia||Present, Few occurrences||EPPO (2020); CABI/EPPO (2012)|
|-Southern Russia||Present, Few occurrences||CABI/EPPO (2012); EPPO (2020)|
|Serbia||Present, Localized||CABI/EPPO (2012); EPPO (2020)|
|Serbia and Montenegro||Present, Widespread||Introduced||1992||Invasive||Bača (1994); Sivčev et al. (1994)|
|Slovakia||Present, Widespread||Introduced||2000||Invasive||Kiss et al. (2005); CABI/EPPO (2012); EPPO (2020)|
|Slovenia||Present, Few occurrences||Introduced||2003||Kiss et al. (2005); CABI/EPPO (2012); EPPO (2020)|
|Spain||Absent, Confirmed absent by survey||EPPO (2020)|
|Switzerland||Present, Transient under eradication||Introduced||Invasive||Kiss et al. (2005); CABI/EPPO (2012); EPPO (2020)||First reported: 2000 and 2003|
|Ukraine||Present, Few occurrences||Introduced||2001||Invasive||Kiss et al. (2005); CABI/EPPO (2012); EPPO (2020)|
|United Kingdom||Absent, Eradicated||2017||2003||Kiss et al. (2005); CABI/EPPO (2012); IPPC (2013); IPPC (2014); EPPO (2020); CABI (Undated)|
|-England||Absent, Eradicated||EPPO (2020)|
|Canada||Present, Localized||Introduced||Invasive||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-Ontario||Present||Introduced||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-Quebec||Present, Localized||EPPO (2020); CABI/EPPO (2012)|
|Costa Rica||Present||Native||CABI/EPPO (2012); EPPO (2020)|
|Guatemala||Present||Native||CABI/EPPO (2012); EPPO (2020)|
|Mexico||Present||Native||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|Nicaragua||Present||Native||CABI/EPPO (2012); EPPO (2020)|
|United States||Present, Widespread||Introduced||Invasive||Yardim et al. (2006); CABI/EPPO (2012); EPPO (2020)|
|-Alabama||Present||CABI/EPPO (2012); EPPO (2020)|
|-Arizona||Present||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-Colorado||Present, Widespread||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-Connecticut||Present||CABI/EPPO (2012); EPPO (2020)|
|-Delaware||Present||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-Georgia||Present||CABI/EPPO (2012); EPPO (2020)|
|-Idaho||Present||CABI/EPPO (2012); EPPO (2020)|
|-Illinois||Present, Widespread||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-Indiana||Present, Widespread||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-Iowa||Present, Widespread||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-Kansas||Present, Widespread||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-Kentucky||Present||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-Louisiana||Present||CABI/EPPO (2012); EPPO (2020)|
|-Maine||Present||CABI/EPPO (2012); EPPO (2020)|
|-Maryland||Present||CABI/EPPO (2012); EPPO (2020)|
|-Massachusetts||Present||CABI/EPPO (2012); EPPO (2020)|
|-Michigan||Present||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-Minnesota||Present, Widespread||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-Missouri||Present, Widespread||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-Montana||Present||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-Nebraska||Present, Widespread||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-New Hampshire||Present||CABI/EPPO (2012); EPPO (2020)|
|-New Jersey||Present||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-New Mexico||Present||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-New York||Present||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-North Carolina||Present||CABI/EPPO (2012); EPPO (2020)|
|-North Dakota||Present||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-Ohio||Present||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-Oklahoma||Present||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-Pennsylvania||Present||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-Rhode Island||Present||CABI/EPPO (2012); EPPO (2020)|
|-South Dakota||Present, Widespread||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-Texas||Present||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-Utah||Present||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-Virginia||Present||CABI/EPPO (2012); EPPO (2020)|
|-Washington||Present||CABI/EPPO (2012); EPPO (2020)|
|-West Virginia||Present||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-Wisconsin||Present, Widespread||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
|-Wyoming||Present||Krysan and Smith (1987); CABI/EPPO (2012); EPPO (2020)|
History of Introduction and SpreadTop of page
D. virgifera virgifera is thought to originate from Central America (Krysan and Smith, 1987).
In the twentieth century, D. virgifera virgifera became a major pest of maize in North America as maize-growing areas increased (Krysan and Miller, 1986; Tallamy et al., 2005). The practice of continuous maize growing is largely responsible for the expanding range of D. virgifera virgifera. D. virgifera virgifera was accidentally introduced several times into Europe from North America between the late 1980s and the early 2000s (Miller et al., 2005). Larvae-induced damage was recorded for the first time near Belgrade airport in former Yugoslavia in 1992 (Baca, 1994). The species became invasive and quickly spread throughout Central Europe, becoming a threat to maize production all over Europe (Gerginov and Tomov, 1995; Kiss et al., 2005b).
See also Kiss et al. (2005b), Miller et al. (2005) and CABI/EPPO (1998, No. 63).
Risk of IntroductionTop of page
D. virgifera virgifera is on the quarantine list of EPPO and of the European Union member countries (Commission decision 2003/766/EC) (Toepfer and Kuhlmann, 2004b). It spread throughout Central Europe in the 1990s and early 2000s. Multiple introductions also occurred in Western Europe. All maize-growing areas of Europe, and in the long-term also of Asia, are at risk (Baufeld and Enzian, 2005) because it is likely to survive and develop wherever maize is grown. The fact that D. virgifera virgifera spreads relatively slowly by natural means, except when carried by storms, offers some potential for its containment (Furlan et al., 2002). However, the rate of spread still poses a serious threat to the whole of Europe.
Habitat ListTop of page
Hosts/Species AffectedTop of page
D. virgifera virgifera has a larval host range restricted to certain Poaceae. The original host range of its larvae in Central America is uncertain (Moeser and Hibbard, 2005).
For lists of secondary hosts, see Branson and Ortman (1967, 1970, 1971), Breitenbach et al. (2005), Moeser and Hibbard (2005).
Host Plants and Other Plants AffectedTop of page
|Cucurbita pepo (marrow)||Cucurbitaceae||Other|
|Fabaceae (leguminous plants)||Fabaceae||Other|
|Glycine max (soyabean)||Fabaceae||Other|
|Helianthus annuus (sunflower)||Asteraceae||Other|
|Tripsacum dactyloides (eastern gamagrass)||Poaceae||Other|
|Zea mays (maize)||Poaceae||Main|
Growth StagesTop of page Flowering stage, Vegetative growing stage
SymptomsTop of page Newly-hatched larvae feed primarily on root hairs. As the larvae grow and their food requirements increase, they burrow into roots. Larval damage is usually most severe after the secondary root system is well established and brace roots are developing. Root tips appear brown and often contain tunnels. In many cases, they are chewed back to the base of the plant. Larvae may be found tunnelling in larger roots and occasionally in the plant crown. Larvae may burrow through plants near the base, causing stunting or death of the growing point and frequently causing tillering. Root feeding commences shortly after plant emergence and early symptoms are expressed as drought or nutrient deficiencies. Plant lodging occurs later in plant development. Sites of larval damage are often pathways for infection by disease pathogens, resulting in root rots.
Adult beetles cause damage by feeding principally on pollen, silk and young kernels. Silk clipping near the husk during anthesis can cause reduced seed set in maize, which may only be observed at the time of harvest.
List of Symptoms/SignsTop of page
|Fruit / external feeding|
|Growing point / dwarfing; stunting|
|Growing point / wilt|
|Inflorescence / external feeding|
|Leaves / external feeding|
|Roots / external feeding|
|Roots / internal feeding|
Biology and EcologyTop of page
The bionomics of D. virgifera virgifera have been reviewed by Chiang (1973). D. virgifera virgifera is univoltine. Adults emerge in the summer and are prevalent in maize fields until the autumn. Eggs are the overwintering stage and are generally concentrated in the top 5-20 cm of soil, although they are situated deeper in dry soils. Eggs require a cold-induced diapause period before hatching, although a small proportion of the population may hatch during a warm, prolonged autumn. The developmental rates of post-diapause eggs of D. virgifera virgifera were investigated and the lower development threshold (10.5°C) was determined by linear regression; completion of post-diapause egg development required 258 day-degrees C above this temperature (Schaafsma et al., 1991). For the emergence of 50% of first-instar larvae, about 265 degree days are needed above a temperature of 11°C in Ontario, Canada (Schaafsma et al., 1993), whereas 292 degree-days above 11°C are needed in South Dakota, USA (Fisher, 1989), and 354 degree-days above 11.2°C in Illinois, USA (Levine et al., 1992 b). Maize has a developmental threshold of 10°C and is planted as early as the soil is free of frost (Aldrich, 1978). In Central Europe, larval emergence usually starts at the beginning of May.
Three larval instars develop in and on the roots. As the temperature increases, development time decreases, up to 33°C when no larvae survive the second instar. As the temperature increases, the proportion of time spent in the third instar increases and the proportion for the first instar decreases (Jackson and Elliot, 1988). The proportions of development time spent in different instars were similar for males and females over a range of temperatures (15-31.5°C) (Jackson and Elliot, 1988).
For the emergence of adults, about 415 degree-days above 9°C are needed in South Dakota, USA (Jackson and Elliot, 1988). In regions with warm, dry summers, numbers of D. virgifera virgifera beetles decline rapidly in mid-August. In climates experiencing cooler summers, adult beetles may be found as long as green maize plants are available. For adults of D. virgifera virgifera maintained at five temperatures (16, 19.5, 23, 26.5 and 30°C), the mean number of eggs laid per female was greatest (602) at 26.5°C and least (295) at 16.0°C. The median lifespan decreased with increasing temperature from 13.8 weeks at 19.5°C to 7.9 weeks at 30.0°C (Elliott et al., 1990a). A reduction in the quality of food associated with maize plant maturity significantly reduced the oviposition period, fecundity and lifespan of D. virgifera virgifera (Elliott et al., 1990b). Greater longevity and increased fecundity has also been observed in early-emerging females of D. virgifera virgifera; beetles emerging during the first part of the growing season appeared to be fitter than their late-emerging counterparts (Boetel and Fuller, 1997). However, Naranjo and Sawyer (1987) report that adults emerging relatively early and late in the field differed in size but did not significantly differ in reproductive potential. The natural mortalities of D. virgifera virgifera life stages were reviewed by Toepfer and Kuhlmann (2005).
Populations of D. virgifera virgifera usually show high levels of genetic diversity; and a mean allelic diversity ranging from 7.3 to 8.6, and expected heterozygosity ranging from 0.600 to 0.670 was reported (Kim and Sappington, 2005). D. virgifera virgifera populations exhibit little genetic differentiation as a whole across the geographic range of nine US states (western Texas and Kansas to New York and Delaware). There was no evidence for a genetic bottleneck in any D. virgifera virgifera population sampled in North America (Kim and Sappington, 2005); however, several bottlenecks were reported for the introduction areas in Europe (Miller et al., 2005).
Natural enemiesTop of page
|Natural enemy||Type||Life stages||Specificity||References||Biological control in||Biological control on|
Notes on Natural EnemiesTop of page
Natural enemies of Diabrotica spp. are listed by Kuhlmann and van der Burgt (1998) in relation to their possibilities for biological control in Europe.
Means of Movement and DispersalTop of page
Although the larvae move relatively small distances, the adults fly to maize fields and are able to migrate over short and longer distances. In addition, adults may be carried by weather features such as cold fronts (Grant and Seevers, 1989) and thunderstorms (Onstad et al., 1999).
D. virgifera virgifera can transmit Maize chlorotic mottle virus (Jensen, 1985).
Maize fields with early flowering maize hybrids may become donor fields when flowering in these fields finishes. Conversely, later flowering maize fields are attractive to beetles and thus may become trap crops.
Movement in trade
There are no obvious means of intercontinental dispersal by trade, as it is unlikely that the insects would be carried by consignments of seeds or grain. However, its multiple introductions into Europe between the late 1980s and early 2000s (Miller et al., 2005) suggest a new but unknown pathway via aeroplanes that carried D. virgifera virgifera but not other Diabrotica pest species. It is possible that D. virgifera virgifera is spread within Europe by consignments of green maize.
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|
|Bulbs/Tubers/Corms/Rhizomes||larvae||Yes||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Flowers/Inflorescences/Cones/Calyx||adults||Yes||Pest or symptoms usually visible to the naked eye|
|Fruits (inc. pods)||adults||Yes||Yes||Pest or symptoms usually visible to the naked eye|
|Growing medium accompanying plants||eggs||Yes||Pest or symptoms usually invisible|
|Leaves||adults||Yes||Pest or symptoms usually visible to the naked eye|
|Roots||larvae||Yes||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Stems (above ground)/Shoots/Trunks/Branches||adults||Yes||Pest or symptoms usually visible to the naked eye|
|Plant parts not known to carry the pest in trade/transport|
|True seeds (inc. grain)|
Impact SummaryTop of page
|Fisheries / aquaculture||None|
ImpactTop of page
D. virgifera virgifera together with D. barberi are the most serious insect pests of maize in the major maize-producing states of the north-central USA (Levine and Oloumi-Sadeghi, 1991). The larvae damage maize roots, which reduces the ability of the plant to absorb water and nutrients from the soil and causes harvesting difficulties due to plant lodging. Adult feeding on silks interferes with pollination. The quantification of yield loss varies according to cultivation practices and location of the field. Generally yield losses have been estimated at around 15% for every node (i.e. circle of roots) damaged from larval feeding (Tinsley et al., 2013).
The costs of soil insecticides to control larval damage to roots, and of aerial spray to reduce adult damage to silks, when combined with crop losses approached US$1000 million annually in the 1980s (Metcalf, 1986; Krysan and Miller, 1986). Since the introduction of the beetle into Europe at the beginning of the 1990s, economic losses have been recorded from Serbia, Hungary, Croatia, Romania, Italy and Austria. The damage potential in Europe is estimated at 472 million Euros annually, when no control measures are implemented (Wesseler and Fall, 2010).
Environmental ImpactTop of page
In the USA, granular insecticides and insecticide-treated maize seeds were applied on a maize-growing area of over 5 million hectares in 2000 (Ward et al., 2005). Transgenic maize varieties were adopted quickly after their release in the USA (e.g., Cry3Bb1 Bt maize increased from 0.2 million ha in 2003 to 12.8 million ha in 2009) (Devos et al., 2013). This initially led to a reduction in insecticide use from 25% of maize treated in 2005 to 9% in 2010. Since the emergence of resistance issues with transgenic maize in 2011, the use of soil insecticides increased again as farmers apply it as an insurance against a possible D. virgifera virgifera problem in Bt maize.
The use of neonicotinoids (a.i. clothianidin) as a preventive measure for D. virgifera virgifera eradication in Germany led to a high number of bee poisonings in 2008 (more than 11,500 honey bee colonies were affected). The abrasion of dust from maize seeds during sowing resulted in a contamination of pollen and nectar in nearby surroundings. Additionally a higher rate of clothianidin was used for the eradication programme than for other pest control measures (e.g. wireworms) (Pistorius et al., 2009). This scenario initiated a temporary ban on neonicotinoids in most EU countries.
Impact: BiodiversityTop of page
D. virgifera virgifera has a low impact on biodiversity because its host plant, maize, is exotic in large areas of North America and Europe and maize fields are therefore relatively poor in arthropod diversity.
Social ImpactTop of page
The social impact of D. virgifera virgifera has not been quantified.
Detection and InspectionTop of page
To detect low population levels of adult D. virgifera virgifera in July and August or to detect early infestations in June or early July, pheromone baited traps are most effective and are commercially available in North America and Europe (Hesler and Sutter, 1993). The sex pheromone racemic 8-methyl-2-decane2-ol-propanoate serves as bait on, for example, a transparent sticky trap to capture males with high sensitivity (Tóth, 2003). These traps are of simple design and are an inexpensive means of D. virgifera virgifera detection. One trap per field <5 ha) must be placed at the ear level of the maize plant and changed every 3 to 4 weeks. With the same trap type, the increase in emerging adult populations can be monitored until their number exceeds 100 per trap per week. Field infestations of D. virgifera virgifera can also be detected by surveying flowering maize plants for adults. 'Goose-necking' of the maize stalk prior to flowering indicates heavy root damage, which is often caused by rootworm larval feeding (Krysan and Miller, 1986).
The Diabrotica ID can be used to identify species of Diabrotica.
Similarities to Other Species/ConditionsTop of page
D. virgifera virgifera is closely related to the subspecies D. virgifera zea; for details refer to Krysan et al. (1980) and Krysan and Smith (1987). D. virgifera virgifera has yellowish elytra with black vittae, whereas D. virgifera zeae is largely green and has entirely pale elytra or with a narrow piceous vitta. D. virgifera virgifera is readily distinguished from D. barberi by the presence of piceous markings on the femora or amber coloration on their dorsal edge. The femora of D. longicornis are pale and uniform in colour (Krysan and Smith, 1987). Other North American species have more distinct spots or bands on the elytra. D. virgifera virgifera and D. barberi occur in the same area for a large part of their range. Their biology and behaviour is very similar (Levine and Chan, 1990). At least under laboratory conditions, they are also interfertile.
Injury caused by D. virgifera virgifera can be confused with that caused by Diatraea spp., sugarcane beetles (Euetheola humilis rugiceps) or wireworms (Elateridae).
Prevention and ControlTop of page
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.
See Levine and Oloumi-Sadeghi (1991), Toepfer et al. (2005b), Gray et al. (2009), Ivezic et al. (2009), Narva et al. (2013) and Van Rozen and Ester (2010) for a review of management strategies for D. virgifera virgifera. Further information can also be found in a series of reviews in a special issue of Agricultural and Forest Entomology on D. virgifera virgifera research (Agricultural and Forest Entomology (2009), 11(1):3-60).
The capacity for natural spread of D. virgifera virgifera is such that it is difficult to propose measures for its prevention. European countries have put in place a monitoring network using pheromone traps to follow spread. In cases of new introductions, immediate insecticide treatments have to be carried out, no maize is allowed to be grown in a focus zone around the introduction point and crop rotation is obligatory in a safety zone around the focus zone (Byrne, 2003).
The main aim at present is to contain D. virgifera virgifera in Europe and to delay its impact as much as possible, mainly by applying crop rotation.
See European Commission decision 2003/766/EC (Byrne, 2003).
In 2014 the European Commission decided to withdraw the recognition of D. virgifera virgifera as a regulated harmful organism with quarantine status by deleting it from Annex 1 to Council Directive 2000/29/EC. The containment measures were also not regarded as successful and Decision 2003/766/EC was therefore repealed. EU Member states are now responsible for providing guidelines for D. virgifera virgifera management, especially taking the principles of IPM into account (Borg, 2014).
Crop rotation is an effective control method for D. virgifera virgifera as the eggs are mainly laid in maize, and the larvae must feed on maize roots to complete their development (Ostlie and Noetzel, 1987; Levine and Oloumi-Sadeghi, 1991). However, a 100% crop rotation would be too strict as an 80% rotation of maize can keep D. virgifera virgifera below the economic threshold. In some cases (e.g. Serbia) even a 60% rotation was sufficient for control. The economic threshold is usually reached when less than 40% of maize fields are rotated. Rotation of each field within 3 years can also lead to about 20% fewer fields that need to be rotated (Szalai et al., 2013).
In principle, all possible crops, fallows or vegetables can be rotated with maize for D. virgifera virgifera management (Kiss et al., 2005a). However, certain crops, such as soyabean or monocotyledonous crops, might be less promising in long-term rotation with Diabrotica-infested maize fields. Many Poaceae are known to serve to some degree as secondary food plants for D. virgifera virgifera larvae (Branson and Ortman, 1967, 1970; Moeser, 2003; Moeser and Hibbard, 2005) and adults feed on nearly every pollen source (Moeser and Hibbard, 2005). However, larval damage on cultivated poaceous plants other than maize has not yet been recorded. In the corn belt of the USA, where soyabean was rotated with maize regularly and over many years, an increased oviposition of D. virgifera virgifera into soyabean was observed and larvae developed in the maize planted in the following year ('crop rotation resistance phenotype') (Gray et al., 1998, 2009). This rotation resistance developed from a decreased fidelity to maize by D. virgifera virgifera females as a consequence of a limited defined crop rotation scheme (Levine et al., 2002). Damage to rotated maize was first reported in an east-central region of Illinois and continued to spread throughout the corn belt (Gray et al., 2009) but at a slow expansion rate (Dunbar and Gassmann, 2013). Furthermore, crop rotation may not be completely effective in 1-year rotations due to the presence of volunteer maize. D. virgifera virgifera has low survivorship <0.1%) over two winters of diapause, but a 1-year rotation may select for rootworms with extended diapause (Krysan et al., 1984).
Most studies have found no significant differences in D. virgifera virgifera oviposition among various tillage practices. However, no-till appears to have the lowest egg mortality.
Delayed planting may result in decreased root damage as eclosed larvae can only survive a few days without feeding on suitable hosts. If planting is delayed until early June, root damage is negligible and soil insecticide usage is not warranted (Musick et al., 1980). Later flowering maize can attract D. virgifera virgifera from surrounding infested maize fields. Thus, late-planted strips of maize can be used as trap crops, although this technique is not widely practised, as its effectiveness is inconsistent.
D. virgifera virgifera has few effective natural enemies in its area of origin in Central America (Eben and Barbercheck, 1996; Kuhlmann and van der Burgt, 1998). In North America, natural enemies undoubtedly help to reduce and stabilize rootworm populations but other control interventions are normally required to bring D. virgifera virgifera populations below an economic threshold. In Europe, host-specific and/or effective indigenous natural enemies do not attack any of the life stages of D. virgifera virgifera (Toepfer and Kuhlmann, 2004a).
Classical biological control
Classical biological control provides an opportunity to reconstruct the natural enemy complex of D. virgifera virgifera populations in Europe (Kuhlmann and Burgt, 1998) and partly in North America. The natural enemy complex of Diabrotica species was surveyed in their area of origin in Central America (Kuhlmann et al., 2005) and Celatoria compressa was the only parasitoid found on the target species, D. virgifera virgifera. Its host range is considered to be restricted to Diabroticite beetles, and thus C. compressa would be safe for introduction because direct and indirect impacts on other organisms would be extremely low (Kuhlmann et al., 2005).
Inundative biological control
Entomopathogenic nematodes have great potential as biological control agents of D. virgifera virgifera (Cabanillas et al., 2005). Several commercially available nematode species have proven effective at killing D. virgifera virgifera larvae; for details, see Cabanillas et al. (2005) and Toepfer et al. (2005a). The nematode species Heterorhabditis bacteriophora has been evaluated as a control agent across numerous studies on a field scale and could reach similar control efficacies to commercially available insecticide treatments (Toepfer, 2010a, b). The first large scale applications of nematodes as a commercial product for D. virgifera virgifera control was carried out in Austria in 2014 (product name: DIANEM ®).
The entomopathogenic fungi Beauveriabassiana and Metarhiziumanisopliae naturally attack D. virgifera virgifera (Toepfer and Kuhlmann, 2004a; Pilz, 2008; Rudeen et al., 2013). Diabrotica populations in the USA are not usually regulated by fungi (Maddox and Kinney, 1989). However, fungi-based products are registered against soil pests in several countries (Inglis et al., 2001) suggesting further research should be carried out towards the development of a fungal biocontrol product against D. virgifera virgifera (Toepfer et al., 2005b). The commercially available M. anisopliae strain BIPESCO5/F52 (product name: GranMet) is included in Annex 1 of Directive 91/414/EEC. Its efficacy against D. virgifera virgifera in the field is currently lower than with insecticides and nematodes (Pilz et al., 2009) but opens further options for biological control. The EU project 'InBioSoil' (www.inbiosoil.uni-goettingen.de) further explores the potential of BIPESCO5 through the use of fungal formulations for soil application and efficacy enhancing agents (e.g. semiochemicals).
Conventional breeding for resistance has resulted in germplasm with moderate levels of resistance to Diabrotica feeding (Knutson et al., 1999). There is currently no maize cultivar with native resistance available against D. virgifera virgifera, due to the limited success of conventional breeding strategies in the past (Moeser and Hibbard, 2005). The predominant mechanism for resistance is tolerance rather than antibiosis or antixenosis, for example, some maize cultivars demonstrate a tolerance to drought stress and larval feeding through the ability to regenerate roots (Branson et al., 1982). Hydroxamic acids have been identified as resistance factors to D. virgifera virgifera larvae in maize root tissue (Xie et al., 1990) and may occur in some commercial hybrids (Assabgui et al., 1993). Large screenings with a focus on root injury evaluation after artificial egg infestation identified resistance genotype but with no information on the underlying resistance mechanism. The use of more advanced genetic/genomic screening techniques (QTL, micorarrays and metabolite analysis) offers the potential to develop non transgenic maize hybrids with antibiosis against D. virgifera virgifera (Gray et al., 2009). The first steps in this process have been made (e.g. 'SUM' cultivars; El-Khishen et al., 2009) and research is ongoing, especially in Europe, to develop breeding programmes for native resistance (Ivezic et al., 2009).
Genetically modified resistance
Genetically modified maize varieties with Bt toxin expressed in their roots (Cry3Bb1, Cry34Ab1/ Cry35Ab1 and mCry3A) can avoid larval damage and have been commercialised in the USA since 2003 (Ward et al., 2005). Seven transgenic events targeting rootworms were released between 2003 and 2013 including pyramided traits with more than one Bt protein and different modes of action (first on the market SmartStax ® in 2009). More stacked Bt events were released in 2014 (eCry3.1Ab+mCry3A). The transgenic maize cultivars have been proven as highly effective (>95%) and were quickly adapted by farmers. Insect resistance management (IRM) plans were established to delay resistance to Bt maize by planting a refuge with a non Bt crop habitat.
In 2009, four populations of D. virgifera virgifera with evolved resistance to Cry3Bb1 were identified in Iowa (Gassmann et al., 2011, 2012; Devos et al., 2013). Resistance was probably triggered through the extensive use of the same Bt-maize repeatedly and exclusively (Devos et al., 2013). The ability of cross resistance to other single events (e.g., Cry34Ab1/ Cry35Ab1) is also possible (Devos et al., 2013). Caution should therefore be taken to mitigate any risks of resistance to other Bt events. Resistance models predict that resistance may be delayed with pyramided traits in Bt-maize in the absence of potential cross resistance (Onstad and Meinke, 2010). The use of these pyramided traits is increasing (Storer et al., 2012) but the Bt maize 'landscape' currently comprises a mixture of single and multiple toxins.
Growing of Bt maize in Europe is often denied by EU or national authorities (some countries have banned the use of Bt maize under the safeguard clause Articles 16 and 18) so there is currently no transformed Bt event for D. virgifera virgifera resistance authorised for cultivation (Meissle et al., 2011).
The use of RNAi (RNA interference), in which double-stranded RNA (dsRNA) molecules trigger gene silencing in a sequence specific manner, could provide a new option for D. virgifera virgifera control (Miyata et al., 2014). D. virgifera virgifera larvae are sensitive to an oral RNAi approach and transgenic plants expressing D. virgifera virgifera dsRNAs show a significant reduction in larval feeding damage (Baum et al., 2007). Further studies will evaluate its potential as a management option and the likelihood for commercial use (Burand and Hunter, 2013).
Chemical control mainly focuses on the larval stage of D. virgifera virgifera as it is economically more feasible (Gerber et al., 2005). Preventive applications of granular soil insecticides or seed treatments against D. virgifera virgifera larvae are widely used as a management strategy in fields where maize is cultivated continuously. However, this strategy protects the roots within the treated band in the maize row but does not reduce populations of larvae that complete their development outside the treated band on peripheral roots (Gray et al., 1992; Boetel et al., 2003; Furlan et al., 2006). Therefore little (if any) resistance against soil insecticides has been documented so far.
Physiochemical properties, degradability, formulation and inherent toxicity are important factors influencing the efficacy of soil-applied insecticides. Tillage practices do not seem to affect the distribution of insecticides in the soil profile although reduced tillage significantly reduces surface runoff (Felsot et al., 1990). Application rates, placement and incorporation are also among the mechanical factors that influence rootworm larvae.
The history of soil-applied insecticides in the USA used to treat D. virgifera virgifera larvae includes the use of chlorinated hydrocarbons (in the 1940s and 1950s) and carbamates or organophospates (since the 1960s). Chemical control of larvae also involves seed treatments with neonicotinoids (e.g. clothianidin). Soil treatments with granular or liquid formulations are dominated by organophospates and synthetic pyrethroids or a combination of the two insecticide classes (Gassmann and Weber, 2013). The WHO has classified some of these insecticides as hazardous pesticides whose use should be restricted (http://www.who.int/ipcs/publications/pesticides_hazard_2009.pdf).
In Europe the main soil insecticides in use are neonicotinoids (clothianidin as seed treatment). Since the temporary ban of clothianidin (see Environmental Impact), the use of pyrethroids is the only option for chemical control of D. virgifera virgifera larvae. Their use in some EU countries is, however, also limited as they can be applied on an approved area (e.g. cypermethrin on 9% of cultivated maize in Austria in 2014) or over a specific time span. This subsequently leaves chemical control as a limited control option for larvae.
Control of adult populations is usually confined to seed production fields and is only occasionally applied to commercial fields. Foliar sprays are generally costly and are affected by rainfall and sprinkler irrigation; they require precise timing to target ovipositing females and must remain active against immigrating gravid females and extended beetle emergence. The most common insecticides to spray against adults in the USA are carbamates (e.g. carbaryl), organophosphates (chlorpyrifos) and synthetic pyrethroids (e.g. bifenthrin). Several cases of resistance to insecticides are known for D. virgifera virgifera, such as regionally developed resistance to organophosphates, carbamates, e.g. carbaryl or chlorinated hydrocarbons (Meinke et al., 1998; Scharf et al., 2000). Moreover, their honeybee toxicity, re-entry interval, and application difficulties due to maize height are limiting factors for application. These insecticides also endanger the biological control of the European corn borer, Ostrinia nubilalis, with Trichogramma parasitoids. The 'attract and kill' method effectively reduces the number of D. virgifera virgifera adults (Chandler, 1998). Therein, a small amount of insecticide (usually 10% of the registered amount, e.g. carbaryl) is combined with natural feeding stimulants/arrestants or attractants, e.g. cucurbitacins (Invite ®)(Borianai et al., 2006).This approach is used on a limited basis and performs best in states like Kansas with a semi-arid climate and low adult populations (Borianai et al., 2006). In Europe the same insecticidal compounds are used (except for carbamate) but are mainly confined to Eastern European countries (Van Rozen and Ester, 2010). Beetles can also be sprayed with a neonicotinoid (a.i. thiacloprid) in EU countries.
Due to the plant size at the time of adult spraying, high clearance tractors are needed for insecticide application. Most farmers in Europe cannot access such equipment, making adult management more difficult.
Aerially applied treatments against adults are not normally used unless there is an average of five beetles per ear and the maize is less than 50% silk and not yet pollinated. Control should be instigated only if an economic level of rootworm beetles is observed. Once the silks turn brown, control is no longer necessary. The most commonly used insecticides include carbaryl, fenvalerate and esfenvalerate, malathion and permethrin. Other insecticides registered for rootworm beetle control in the USA include chlorpyrifos and diazinon; these compounds are more toxic to humans. The use of aerial equipment is usually not supported by EU authorities (and not well received by the public society) due to environmental concerns such as water contamination and is therefore prohibited in certain EU countries.
As D. virgifera virgifera is a very actively moving beetle with the ability to mate throughout its entire lifespan, pheromonal control is limited. However, semiochemicals have been used to develop insecticide baits (Metcalf et al., 1987).
The pheromone for D. virgifera virgifera is (2R,8R) 8-methyl-2-decyl propanoate. The four stereoisomers of 8-methyl-2-decyl propanoate were tested for attractiveness to D. virgifera virgifera. The (2R,8R)- configuration was most attractive while (2R,8R), (2S,8R)- or (2S,8S)-isomers were neither attractive nor repellent (Guss et al., 1985).
The synthetic maize volatiles geranylacetone and alpha-terpineol attract D. virgifera virgifera and D. barberi (Hammack, 1996).
Active volatiles from Cucurbita blossoms contain 1,2,4-trimethoxygenzene, indole and (E)-cinnamaldehyde. This mixture represents an optimised Cucurbita blossom attractant volatile kairomone mixture useful in monitoring Diabrotica populations and in studying their behaviour and ecology (Metcalf et al., 1995). Other volatiles attractive to D. virgifera virgifera include 4-methoxycinnamaldehyde, 4-methosycinnamonitrile, beta-ionone and estragole. In field trapping tests, phenyl alkyl amines and phenyl alkyl alcohols with 2-carbon side chains attracted significantly more adults than did phenyl alkyl amines or phenyl alkyl alcohols with 1-, 3- or 4-carbon side chains. Cinnamaldehyde attracted the most D. virgifera virgifera of both sexes. Male D. virgifera virgifera only responded to cinnamaldehyde (Petroski and Hammack, 1998).
Early Warning Systems
Pheromone baited traps are commercially available in North America and Europe and are usually used to detect early infestations with adults of D. virgifera virgifera. Monitoring adults in one season provides a reasonably accurate forecast for damage in the following season (Levine and Gray, 1994).
Field Monitoring/Economic Threshold Levels
The monitoring of high population levels of larvae is easily accomplished, whereas the monitoring of low population levels and the quantitative measurements of larval densities are more labour intensive. To detect high population levels of larvae and to determine the need for control in areas where maize is grown continuously without crop rotation, heavy larval damage can be measured. 'Goose-necking' of the maize stalk prior to flowering indicates root damage, which is often caused by rootworm larval feeding (Krysan and Miller, 1986). The larvae can be detected in June upon excavation of the plants, or the damage to the maize roots can be rated in early July. To detect larvae, 10 to 20 randomly selected maize roots with surrounding soil have to be excavated per field <5 ha) with cores or with a shovel (Bergmann et al., 1981). To achieve a rough estimate of population levels, 30 systematically chosen plant roots have to be excavated per hectare. Larvae can be counted by hand-search over black plastic sheets or in Berlese-Tullgren-funnels. In the same sampling programme, maize roots can be collected, washed, and larval damage can be rated according to root-rating scales, i.e. the Iowa traditional scale 1 to 6 (Hills and Peters, 1971) or the node-injury scale 0.00 to 3.00 (Oleson et al., 2005).
To monitor low population levels of D. virgifera virgifera adults in July and August, baited sticky traps are most effective (Gerber et al., 2005). The sex pheromone racemic 8-methyl-2-decane2-ol-propanoate serves as bait on a transparent sticky trap to capture males with high sensitivity (Toth et al., 2003). One trap per field <5 ha) can be placed at the ear level of the maize plant and the increase in adult populations can be monitored until their number exceeds 100 per trap per week (Toepfer and Kuhlmann, 2004b).
To estimate high population levels of adults and to follow their temporal occurrence, three methods are commonly implemented from late June until September or at least during the period just before silking until the end of silking (Gerber et al., 2005). For determination of economic thresholds these methods should be implemented after mid-July. Monitoring adults in one season gives a reasonably good forecast for damage in the following season (Levine and Gray, 1994; Shaw et al., 1984).
Adults can be counted visually on five whole plants in each of 10 randomly selected sites in a maize field (Baca et al., 1995; Hesler and Sutter, 1993). Samples should be taken in a 'U'-shape to ensure that samples are taken from each quarter of the field.
The adults can be counted visually by examining the ear zone of five plants at 32 random sites per field (for a total of 160 plants), accounting for approximately 50% of the adult number per plant. A sequential sampling table is available and can save time when dealing with high populations (Hein and Foster, 1988).
Less labour intensive monitoring methods involve two trap types: (a) The non-baited yellow sticky trap is a simple, inexpensive and commonly used trap to capture males and females with low sensitivity. Two traps per field of <5 ha and up to four traps in larger fields have to be placed at ear level of the maize plants (Hesler and Sutter, 1993) and changed every 3 to 4 weeks. To achieve a rough estimation of population levels, 12 traps have to be placed per field, each approximately 20 m apart. (b) The floral kairomone, 4-methoxy cinnamaldehyde (MCA) plus indole, serves as a bait on a yellow sticky trap to capture females and males at higher sensitivity than the non-baited yellow sticky trap, and males at four times lower sensitivity than the pheromone sticky trap (Zoltan et al., 2001). One trap per field <5 ha) has to be placed at the ear level of the maize plant and changed every 3 to 4 weeks.
Several models were developed to predict population growth of Diabrotica spp. (Elliott et al., 1990; Tollefson, 1990; Elliott and Hein, 1991; Schaafsma et al., 1993; Szalai et al., 2011). A rough estimate is that a D. virgifera virgifera population in an isolated, weed-free monoculture field builds up to its maximum population within 4 to 5 years with an annual growth rate of 4. In areas with large-scale maize production and neighbouring infested fields, major population increase is observed in the first and second year after infestation of a field.
Economic thresholds are expressed by (1) the number of larvae per plant in root-soil samples, (2) rating the root damage, or (3) the average and cumulative numbers of adults per plant (Gerber et al., 2005):
(1) About 8 to 10 third-instar larvae per plant root are often estimated to cause economic damage in commercial maize at 60,000 plants per hectare, depending on local requirements (Toepfer and Kuhlmann, 2004b).
(2) In the root damage rating system, the value 2.5 or 3 on the traditional IOWA scale is considered as economic damage (Turpin and Maxwell, 1976). A new linear decimal 0.00-3.00 node injury scale has lately been used to rate root damage (Oleson et al., 2005). On this scale the suggested economic threshold level in conventional grain maize is at 0.25 for the corn belt of the USA and 0.75 for irrigated conventional maize growing in northern Italy (Edwards et al., 2010).
(3) More than 0.75 to 1 adult per whole plant on average have to be recorded several times from mid-July until the beginning of August to meet the economic threshold necessary to control D. virgifera virgifera in the following year (Turpin et al., 1972; Stamm et al., 1985). The threshold for damage in the current year of commercial maize production is about 5 to 10 adults per whole plant. At 0.5 adults per whole plant, rotation of the crop is recommended for the following year. When more than 6 adults are captured per yellow sticky trap on average per day, economic damage is expected in the following year (Baca et al., 1995). Unfortunately, counting larvae or adults does not provide reliable estimations of absolute population densities, as larvae and adult distributions are clumped (Ellsbury et al., 1996). Furthermore, all recommendations depend on regional weather and soil conditions as well as on the maize variety, e.g. sweet, commercial or seed maize (Baca et al., 1995), and finally on the infestation of neighbouring fields.
The application of one control option alone will not control this pest in the long term (Onstad et al., 2001; Toepfer et al., 2005b). The main elements of D. virgifera virgifera management include monitoring pest populations (principally adults), use of economic thresholds, and integration of control tactics (including crop rotation, insecticide treatment, transgenic Bt maize and in the future, applications of entomopathogenic nematodes and/or fungi) (Nishimatsu and Jackson, 1998; Toepfer and Kuhlmann, 2004b).
General IPM guidelines for maize growing in Europe are published by IOBC (Boller et al., 1997).
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