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Helicoverpa zea
(bollworm)

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Datasheet

Helicoverpa zea (bollworm)

Pictures

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PictureTitleCaptionCopyright
Helicoverpa zea (American cotton bollworm); adult. Cuivre River State Park, Missouri USA. September 2014.
TitleAdult
CaptionHelicoverpa zea (American cotton bollworm); adult. Cuivre River State Park, Missouri USA. September 2014.
Copyright©Andy Reago & Chrissy McClarren/via wikipedia - CC BY 2.0
Helicoverpa zea (American cotton bollworm); adult. Cuivre River State Park, Missouri USA. September 2014.
AdultHelicoverpa zea (American cotton bollworm); adult. Cuivre River State Park, Missouri USA. September 2014.©Andy Reago & Chrissy McClarren/via wikipedia - CC BY 2.0
Helicoverpa zea (American cotton bollworm); a normal, 12-day-old cotton bollworm larva raised on a control diet. USA.
TitleLarva
CaptionHelicoverpa zea (American cotton bollworm); a normal, 12-day-old cotton bollworm larva raised on a control diet. USA.
Copyright©Peggy Greb/USDA Agricultural Research Service/Bugwood.org - CC BY 3.0 US
Helicoverpa zea (American cotton bollworm); a normal, 12-day-old cotton bollworm larva raised on a control diet. USA.
LarvaHelicoverpa zea (American cotton bollworm); a normal, 12-day-old cotton bollworm larva raised on a control diet. USA.©Peggy Greb/USDA Agricultural Research Service/Bugwood.org - CC BY 3.0 US
Helicoverpa zea (American cotton bollworm); larva and larval damage to tomato fruit. USA. June 2017.
TitleLarva
CaptionHelicoverpa zea (American cotton bollworm); larva and larval damage to tomato fruit. USA. June 2017.
Copyright©C.Watts/via flickr - CC BY 2.0
Helicoverpa zea (American cotton bollworm); larva and larval damage to tomato fruit. USA. June 2017.
LarvaHelicoverpa zea (American cotton bollworm); larva and larval damage to tomato fruit. USA. June 2017.©C.Watts/via flickr - CC BY 2.0
Helicoverpa zea (American cotton bollworm); late instar larva on cotton boll (Gossypium hirsutum). USA.
TitleLarva
CaptionHelicoverpa zea (American cotton bollworm); late instar larva on cotton boll (Gossypium hirsutum). USA.
Copyright©Ronald Smith/Auburn University/Bugwood.org - CC BY 3.0 US
Helicoverpa zea (American cotton bollworm); late instar larva on cotton boll (Gossypium hirsutum). USA.
LarvaHelicoverpa zea (American cotton bollworm); late instar larva on cotton boll (Gossypium hirsutum). USA.©Ronald Smith/Auburn University/Bugwood.org - CC BY 3.0 US
Helicoverpa zea (American cotton bollworm); full grown larva on cotton boll. USA.
TitleLarva
CaptionHelicoverpa zea (American cotton bollworm); full grown larva on cotton boll. USA.
Copyright©Scott Bauer/USDA Agricultural Research Service/Bugwood.org - CC BY 3.0 US
Helicoverpa zea (American cotton bollworm); full grown larva on cotton boll. USA.
LarvaHelicoverpa zea (American cotton bollworm); full grown larva on cotton boll. USA.©Scott Bauer/USDA Agricultural Research Service/Bugwood.org - CC BY 3.0 US
Helicoverpa zea (American cotton bollworm); larva in the field, on cotton (Gossypium hirsutum). USA.
TitleLarva
CaptionHelicoverpa zea (American cotton bollworm); larva in the field, on cotton (Gossypium hirsutum). USA.
Copyright©Russ Ottens/University of Georgia/Bugwood.org - CC BY 3.0 US
Helicoverpa zea (American cotton bollworm); larva in the field, on cotton (Gossypium hirsutum). USA.
LarvaHelicoverpa zea (American cotton bollworm); larva in the field, on cotton (Gossypium hirsutum). USA.©Russ Ottens/University of Georgia/Bugwood.org - CC BY 3.0 US
Helicoverpa zea (American cotton bollworm); larval damage under bloom. USA.
TitleLarval damage
CaptionHelicoverpa zea (American cotton bollworm); larval damage under bloom. USA.
Copyright©Ronald Smith/Auburn University/Bugwood.org - CC BY 3.0 US
Helicoverpa zea (American cotton bollworm); larval damage under bloom. USA.
Larval damageHelicoverpa zea (American cotton bollworm); larval damage under bloom. USA.©Ronald Smith/Auburn University/Bugwood.org - CC BY 3.0 US
Helicoverpa zea (American cotton bollworm); larva and larval damage to corn ear (Zea mays). USA. April 2017.
TitleLarval damage
CaptionHelicoverpa zea (American cotton bollworm); larva and larval damage to corn ear (Zea mays). USA. April 2017.
Copyright©Scot Nelson/via flickr - CC BY 2.0
Helicoverpa zea (American cotton bollworm); larva and larval damage to corn ear (Zea mays). USA. April 2017.
Larval damageHelicoverpa zea (American cotton bollworm); larva and larval damage to corn ear (Zea mays). USA. April 2017.©Scot Nelson/via flickr - CC BY 2.0
Helicoverpa zea (American cotton bollworm); larva and larval damage to a corn ear (Zea mays). USA.
TitleLarval damage
CaptionHelicoverpa zea (American cotton bollworm); larva and larval damage to a corn ear (Zea mays). USA.
Copyright©Eric R. Day/Virginia Polytechnic Institute & State University/Bugwood.org - CC BY 3.0 US
Helicoverpa zea (American cotton bollworm); larva and larval damage to a corn ear (Zea mays). USA.
Larval damageHelicoverpa zea (American cotton bollworm); larva and larval damage to a corn ear (Zea mays). USA.©Eric R. Day/Virginia Polytechnic Institute & State University/Bugwood.org - CC BY 3.0 US
Helicoverpa zea (American cotton bollworm); larval damage to corn ear (Zea mays). USA.
TitleLarval damage
CaptionHelicoverpa zea (American cotton bollworm); larval damage to corn ear (Zea mays). USA.
Copyright©Whitney Cranshaw/Colorado State University/Bugwood.org - CC BY 3.0 US
Helicoverpa zea (American cotton bollworm); larval damage to corn ear (Zea mays). USA.
Larval damageHelicoverpa zea (American cotton bollworm); larval damage to corn ear (Zea mays). USA.©Whitney Cranshaw/Colorado State University/Bugwood.org - CC BY 3.0 US
Helicoverpa zea (American cotton bollworm); larva in the field, feeding on a panicle of pearl millet (Pennisetum glaucum). USA.
TitleLarva
CaptionHelicoverpa zea (American cotton bollworm); larva in the field, feeding on a panicle of pearl millet (Pennisetum glaucum). USA.
Copyright©Russ Ottens/University of Georgia/Bugwood.org - CC BY 3.0 US
Helicoverpa zea (American cotton bollworm); larva in the field, feeding on a panicle of pearl millet (Pennisetum glaucum). USA.
LarvaHelicoverpa zea (American cotton bollworm); larva in the field, feeding on a panicle of pearl millet (Pennisetum glaucum). USA.©Russ Ottens/University of Georgia/Bugwood.org - CC BY 3.0 US
Helicoverpa zea (American cotton bollworm); pupa, in field, on soyabean (Glycine max). USA. September 2011.
TitlePupa
CaptionHelicoverpa zea (American cotton bollworm); pupa, in field, on soyabean (Glycine max). USA. September 2011.
Copyright©Adam Sisson/Iowa State University/Bugwood.org - CC BY 3.0 US
Helicoverpa zea (American cotton bollworm); pupa, in field, on soyabean (Glycine max). USA. September 2011.
PupaHelicoverpa zea (American cotton bollworm); pupa, in field, on soyabean (Glycine max). USA. September 2011.©Adam Sisson/Iowa State University/Bugwood.org - CC BY 3.0 US
Helicoverpa zea (American cotton bollworm); adult.
TitleAdult
CaptionHelicoverpa zea (American cotton bollworm); adult.
CopyrightPublic Domain - Released by the USGS Bee Inventory & Monitoring Lab.
Helicoverpa zea (American cotton bollworm); adult.
AdultHelicoverpa zea (American cotton bollworm); adult.Public Domain - Released by the USGS Bee Inventory & Monitoring Lab.

Identity

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Preferred Scientific Name

  • Helicoverpa zea Boddie (1850)

Preferred Common Name

  • bollworm

Other Scientific Names

  • Bombyx obsoleta Fabricius
  • Chloridea obsoleta Fabricius
  • Heliothis armigera auct.nec Huebner Hübner
  • Heliothis ochracea Cockerell
  • Heliothis umbrosa Grote
  • Heliothis zea Boddie
  • Phalaena zea Boddie

International Common Names

  • English: bollworm; bollworm, American; corn earworm; tomato fruitworm
  • Spanish: bellotero; elotero; gusano bellotero del algodon; gusano de la bellota del algodón; gusano de la mazorca; gusano de la mazorca del maiz; gusano de las cápsulas; gusano del elote del maíz; gusano del fruto del tomate; gusano elotero; noctua del tomate; oruga de la mazorca
  • French: chenille des epis du mais; noctuelle de la tomate; noctuelle des tomates; ver de la capsule; ver de l'épi du maïs

Local Common Names

  • Argentina: isoca del maiz
  • Brazil: lagarta da espiga do milho; lagarta das espicas
  • Denmark: amerikansk bomuldsugle
  • Germany: Amerikanischer Baumwollkapselwurm; Wurm, Amerikanischer Baumwollkapsel-
  • Italy: elotide del cotone; elotide del granturco; elotide del pomodoro; elotide del tomato; nottua del granturco; nottua gialla del granturco
  • Netherlands: mimosa-rups
  • Turkey: yesil kurt

Taxonomic Tree

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  • Domain: Eukaryota
  •     Kingdom: Metazoa
  •         Phylum: Arthropoda
  •             Subphylum: Uniramia
  •                 Class: Insecta
  •                     Order: Lepidoptera
  •                         Family: Noctuidae
  •                             Genus: Helicoverpa
  •                                 Species: Helicoverpa zea

Notes on Taxonomy and Nomenclature

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The taxonomic situation regarding H. zea is complicated and presents several problems. Hardwick (1965) reviewed the New World corn earworm species complex and the Old World African bollworms, most of which had previously been referred to as a single species (Heliothis armigera or Heliothis obsoleta), and pointed out that there was a complex of species and subspecies involved. Specifically, he proposed that the New World H. zea (first used in 1955) was distinct from the Old World H. armigera on the basis of male and female genitalia; he described the new genus Helicoverpa to include these important pest species. Some 80 or more species were formerly placed in Heliothis (sensu lato) and Hardwick referred 17 species (including 11 new species) to Helicoverpa on the basis of differences in both male and female genitalia. Within this new genus the zea group contains eight species, and the armigera group two species with three subspecies (Hardwick, 1970).

Because the old name of Heliothis for the pest species (four major pest species and three minor) is so well established in the literature, and since dissection of genitalia or genomics is required for identification, there has been resistance to the name change (for example, Heath and Emmet, 1983), but Hardwick's work is generally accepted and so the name change must also be accepted (Matthews, 1991). Later genomic studies confirmed that H. armigera and H. zea are sister species (Cho et al., 1995; Behere et al., 2007). Accepted common names for H. zea are bollworm, corn earworm and tomato fruitworm.

Description

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Egg

Eggs are subspherical, radially ribbed (n = 11 to 17), 0.51 mm high and 0.57 mm in diameter, attached individually to the plant substrate, white to yellowish-green when laid, developing a reddish band and finally turning dark grey before hatching. Egg maturity takes 2-3 days at 20-30°C (Neunzig, 1964; Hardwick, 1965). Eggs are preferentially laid on hosts during the flowering period and on many different tissue types, but primarily on leaves (Hardwick, 1965; Neunzig, 1969; Braswell et al., 2019c).

Larva

On hatching, the tiny caterpillars are translucent or yellowish-white, with a black head, but appear grey or brown to the naked eye; they grow through six instars usually, but five and seven instars are not uncommon, and the final body size is approximately 40 mm long. In the third instar, different colour phases can develop. Often, longitudinal lines of white, cream or yellow are present, and the spiracular band is the most distinct. As the larvae develop, the pattern becomes better defined, but in the final instar (sixth) the colouration can change abruptly into a bright pattern with extra striations. However, larvae can be green, reddish, or pink, without most of the brown or black pigmentation. Larval colour is determined by the interaction of environment (light, temperature and developmental host) and genetics. Larvae have five pairs of prolegs (Neunzig, 1964; Hardwick, 1965; Neunzig, 1969).

Pupa

Pupae are light to dark brown depending on maturity and approximately 20 mm long, with two distinct terminal cremaster spines. Pupae reside in earthen cells, ranging from 0.5-25 cm below the soil surface (Barber, 1941; Eger et al., 1983), but most commonly around 3-5 cm (Roach and Hopkins, 1979; Stadelbacher and Martin, 1980; Eger et al., 1983).

Adult

A stout-bodied (20-25 mm long) brown moth of wingspan 38-43 mm; forewing pale brown-yellow to brown-pink (female) to greenish (male) with darker transverse markings, underwings pale with a broad dark marginal band.

Distribution

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Helicoverpa zea is confined to the New World. It occurs throughout the Americas from Canada to Argentina (International Institute of Entomology, 1993).

Distribution Table

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The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.

Last updated: 12 May 2022
Continent/Country/Region Distribution Last Reported Origin First Reported Invasive Reference Notes

Europe

NetherlandsAbsent, Confirmed absent by survey
RussiaAbsent, Confirmed absent by survey
SloveniaAbsent
SwitzerlandAbsent, Intercepted only
United KingdomAbsent, Intercepted onlyOriginal citation: Seymour and (1978)

North America

Antigua and BarbudaPresent
BahamasPresent
BarbadosPresent
BermudaPresent
CanadaPresent, Localized
-British ColumbiaPresent
-ManitobaPresent
-New BrunswickPresent
-Nova ScotiaPresent
-OntarioPresent
-QuebecPresent
-SaskatchewanPresent
Costa RicaPresent
CubaPresent
DominicaPresent
Dominican RepublicPresent
El SalvadorPresent
GuadeloupePresent
GuatemalaPresent
HaitiPresent
HondurasPresent
JamaicaPresent
MartiniquePresent, Widespread
MexicoPresent, Widespread
MontserratPresent
NicaraguaPresent
PanamaPresent
Puerto RicoPresent
Saint Kitts and NevisPresent, Localized
Saint LuciaPresent
Saint Vincent and the GrenadinesPresent
Trinidad and TobagoPresent, Widespread
U.S. Virgin IslandsPresent
United StatesPresent, Widespread
-AlabamaPresent
-ArizonaPresent
-ArkansasPresent
-CaliforniaPresent
-ColoradoPresent
-ConnecticutPresent
-DelawarePresent
-FloridaPresent
-GeorgiaPresent
-HawaiiPresentIntroducedAs: Heliothis zea. First reported: ~1930
-IdahoPresent
-IllinoisPresent
-IndianaPresent
-IowaPresent
-KansasPresent
-KentuckyPresent
-LouisianaPresent
-MainePresent
-MarylandPresent
-MassachusettsPresent
-MichiganPresent
-MinnesotaPresent
-MississippiPresent
-MissouriPresent
-MontanaPresent
-NebraskaPresent
-NevadaPresent
-New HampshirePresent
-New JerseyPresent
-New MexicoPresent
-New YorkPresent
-North CarolinaPresent
-North DakotaPresent
-OhioPresent
-OklahomaPresent
-OregonPresent
-PennsylvaniaPresent
-Rhode IslandPresent
-South CarolinaPresent
-South DakotaPresent
-TennesseePresent
-TexasPresent
-UtahPresent
-VermontPresent
-VirginiaPresent
-WashingtonPresent
-West VirginiaPresent
-WisconsinPresent
-WyomingPresent

South America

ArgentinaPresent
BoliviaPresent
BrazilPresent, Widespread
-BahiaPresent
-CearaPresent
-Distrito FederalPresent
-GoiasPresent
-Mato GrossoPresent
-Mato Grosso do SulPresent
-Minas GeraisPresent
-ParaPresent
-ParanaPresent
-PernambucoPresent
-Rio de JaneiroPresent
-Rio Grande do SulPresent
-RoraimaPresent
-Santa CatarinaPresent
-Sao PauloPresent
ChilePresent, Widespread
-Easter IslandPresent
ColombiaPresent
EcuadorPresent, Widespread
Falkland IslandsPresent, Few occurrences
French GuianaPresent
GuyanaPresent
ParaguayPresent, Widespread
PeruPresent
SurinamePresent
UruguayPresent, Widespread
VenezuelaPresent

Risk of Introduction

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Helicoverpa zea was recently added to the EPPO A1 list of quarantine pests and is also considered as a quarantine pest by APPPC. Originally, H. zea was considered as practically synonymous with Helicoverpa armigera, an A2 quarantine pest (EPPO/CABI, 1996). The addition to the EPPO list harmonizes it with EU Directive Annex I/A1.

Phytosanitary Measures

For the related H. armigera, EPPO (OEPP/EPPO, 1990) makes recommendations on phytosanitary measures which would also be suitable for H. zea. According to these, imported propagation material should derive from an area where H. armigera does not occur or from a place of production where H. armigera has not been detected during the previous 3 months.

Bibliographies are included in the monograph by Hardwick (1965) (2000 titles on H. zea), and the reviews by Fitt (1989) (194 titles), and King and Coleman (1989) (159 references). Most of the basic research on H. zea was done in the early 1900s and published under early synonyms. Many references to H. zea are made in publications relating to the cultivation/protection of specific crops, for example, Chiang (1978), Centre for Overseas Pest Research (1983) and Pitre (1985).

Habitat List

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CategorySub-CategoryHabitatPresenceStatus
Terrestrial ManagedCultivated / agricultural land Present, no further details Harmful (pest or invasive)
Terrestrial ManagedProtected agriculture (e.g. glasshouse production) Present, no further details Harmful (pest or invasive)
Terrestrial ManagedManaged forests, plantations and orchards Present, no further details Harmful (pest or invasive)
Terrestrial ManagedManaged grasslands (grazing systems) Present, no further details Natural
Terrestrial ManagedDisturbed areas Present, no further details Natural
Terrestrial ManagedRail / roadsides Present, no further details Natural
Terrestrial Natural / Semi-naturalNatural forests Present, no further details Natural
Terrestrial Natural / Semi-naturalNatural grasslands Present, no further details Natural
Terrestrial Natural / Semi-naturalRiverbanks Present, no further details Natural

Hosts/Species Affected

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Helicoverpa zea is polyphagous in feeding habits but it shows a definite preference in North America for young maize cobs, and particularly for the cultivars grown as sweetcorn and popcorn, and also for sorghum. Most hosts are recorded from the Fabaceae (41% of total), Solanaceae (14%), Poaceae (9%), Asteraceae (8%) and Malvaceae (8%) (Cunningham and Zalucki, 2014); in total, more than 100 plant species are recorded as hosts. A feeding preference is shown for flowers and fruits of host plants.

For further information see Barber (1937)Neunzig (1963)Davidson and Peairs (1966) and Matthews (1991).

Host Plants and Other Plants Affected

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Plant nameFamilyContextReferences
Abelmoschus esculentus (okra)MalvaceaeMain
Abelmoschus esculentus (okra)MalvaceaeOther
Abutilon theophrasti (velvet leaf)MalvaceaeOther
Acalypha (Copperleaf)EuphorbiaceaeWild host
Alcea rosea (Hollyhock)MalvaceaeOther
Allium cepa (onion)LiliaceaeOther
Amaranthus (amaranth)AmaranthaceaeOther
Amaranthus palmeri (Palmer amaranth)AmaranthaceaeWild host
Arachis hypogaea (groundnut)FabaceaeOther
Asparagus officinalis (asparagus)LiliaceaeOther
Astragalus crassicarpusWild host
Brachiaria texanaPoaceaeWild host
Brassica oleracea (cabbages, cauliflowers)BrassicaceaeOther
Brassica oleracea var. botrytis (cauliflower)BrassicaceaeOther
Brassica oleracea var. capitata (cabbage)BrassicaceaeOther
Brassica oleracea var. viridis (collards)BrassicaceaeOther
Cajanus cajan (pigeon pea)FabaceaeMain
Canna indica (canna lilly)CannaceaeWild host
Cannabis sativa (hemp)CannabaceaeOther
Capsicum (peppers)SolanaceaeOther
Capsicum annuum (bell pepper)SolanaceaeMain
Capsicum frutescens (chilli)SolanaceaeWild host
Castilleja indivisaScrophulariaceaeWild host
Chenopodium quinoa (quinoa)ChenopodiaceaeOther
Chrysanthemum (daisy)AsteraceaeOther
Cicer arietinum (chickpea)FabaceaeOther
Citrullus lanatus (watermelon)CucurbitaceaeOther
CitrusRutaceaeOther
Cleome gynandraCapparaceaeWild host
Cleome spinosaCapparaceaeWild host
Coronilla variaOther
Croptilon divaricumWild host
Crotalaria (rattlepods)FabaceaeWild host
Croton hirtusEuphorbiaceaeWild host
Cucumis melo (melon)CucurbitaceaeOther
Cucumis sativus (cucumber)CucurbitaceaeOther
Cucurbita pepo (marrow)CucurbitaceaeOther
Cynara cardunculus var. scolymus (globe artichoke)AsteraceaeUnknown
Datura stramonium (jimsonweed)SolanaceaeWild host
Desmodium rigidumWild host
Desmodium tortuosum (Florida beggarweed)FabaceaeWild host
Digitaria sanguinalis (large crabgrass)PoaceaeWild host
Erythrina herbaceaFabaceaeWild host
Ficus carica (common fig)MoraceaeOther
Fragaria (strawberry)RosaceaeOther
Fragaria ananassa (strawberry)RosaceaeOther
Fragaria chiloensis (Chilean strawberry)RosaceaeWild host
Geranium carolinianum (Carolina geranium)GeraniaceaeOther
Geranium dissectum (cutleaf geranium)GeraniaceaeOther
Gerbera (Barbeton daisy)AsteraceaeOther
Gladiolus (sword lily)IridaceaeOther
Glycine max (soyabean)FabaceaeMain
Gossypium (cotton)MalvaceaeMain
Helianthus annuus (sunflower)AsteraceaeMain
Helianthus debilis (beach sunflower)AsteraceaeWild host
Hibiscus rosa-sinensis (China-rose)MalvaceaeOther
Ipomea cordatotrilobaWild host
Ipomoea purpurea (tall morning glory)ConvolvulaceaeOther
Jacquemontia tamnifolia (Smallflower morningglory)Wild host
Kummerowia stipulaceaWild host
Lactuca sativa (lettuce)AsteraceaeOther
Lamium amplexicaule (henbit deadnettle)LamiaceaeWild host
Lathyrus hirsutusFabaceaeWild host
Lathyrus latifolius (broad-leaved everlasting pea (UK))FabaceaeWild host
Lespedeza juncea var. sericea (Sericea lespedeza)FabaceaeWild host
Linaria canadensisScrophulariaceaeWild host
Linum usitatissimum (flax)Wild host
Lonicera japonica (Japanese honeysuckle)CaprifoliaceaeWild host
Ludwigia decurrensWild host
Lupinus (lupins)FabaceaeWild host
Lupinus texensisWild host
Medicago lupulina (black medick)FabaceaeWild host
Medicago polymorpha (bur clover)FabaceaeWild host
Medicago sativa (lucerne)FabaceaeOther
Melilotus albus (honey clover)FabaceaeWild host
Melilotus officinalis (yellow sweet clover)FabaceaeWild host
Nicotiana tabacum (tobacco)SolanaceaeOther
Oenothera (evening primrose)OnagraceaeWild host
Panicum miliaceum (millet)PoaceaeOther
Panicum scopariumWild host
Persicaria pensylvanicaPolygonaceaeWild host
PetitiaWild host
Phaseolus (beans)FabaceaeMain
Phaseolus lanatusMain
Phaseolus vulgaris (common bean)FabaceaeMain
Physalis (Groundcherry)SolanaceaeWild host
Pisum sativum (pea)FabaceaeMain
Prunus persica (peach)RosaceaeMain
Pyrus communis (European pear)RosaceaeMain
Rosa (roses)RosaceaeMain
Ruellia ciliatifloraWild host
Ruellia nudiflora var. runyoniWild host
Saccharum officinarum (sugarcane)PoaceaeOther
Salix (willows)SalicaceaeOther
Sesamum indicum (sesame)PedaliaceaeMain
Setaria italica (foxtail millet)PoaceaeWild host
SidaMalvaceaeWild host
Sida spinosa (teaweed (USA))MalvaceaeWild host
Solanum carolinense (horsenettle)SolanaceaeWild host
Solanum lycopersicum (tomato)SolanaceaeOther
Solanum lycopersicum (tomato)SolanaceaeMain
Solanum melongena (aubergine)SolanaceaeMain
Solanum rostratum (prickly nightshade)SolanaceaeWild host
Solanum tuberosum (potato)SolanaceaeMain
Sorghum bicolor (sorghum)PoaceaeMain
Sorghum halepense (Johnson grass)PoaceaeWild host
Spinacia oleracea (spinach)ChenopodiaceaeOther
Stachys agrariaWild host
Tagetes (marigold)AsteraceaeOther
Trifolium (clovers)FabaceaeWild host
Trifolium campestre (Hop trefoil)FabaceaeWild host
Trifolium hybridum (alsike clover)FabaceaeWild host
Trifolium incarnatum (Crimson clover)FabaceaeWild host
Trifolium pratense (red clover)FabaceaeWild host
Trifolium repens (white clover)FabaceaeWild host
Trifolium resupinatum (Shaftal clover)FabaceaeWild host
Triticum aestivum (wheat)PoaceaeMain
Verbena neomexicanaWild host
Vicia sativa (common vetch)FabaceaeOther
Vicia villosa (hairy vetch)FabaceaeWild host
Vigna unguiculata (cowpea)FabaceaeOther
Vitis (grape)VitaceaeMain
Xanthium (Cocklebur)AsteraceaeWild host
Xanthium strumarium (common cocklebur)AsteraceaeWild host
Zea mays (maize)PoaceaeMain
Zea mays subsp. mays (sweetcorn)PoaceaeMain

Growth Stages

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Flowering stage, Fruiting stage, Vegetative growing stage

Symptoms

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Fruiting structures are consumed or damaged and feeding damage can facilitate entry by diseases and other insect pests. In cotton the square (flower bud), flowers and young bolls are attacked and larvae excavate the interior. This can cause the reproductive tissue to abscise and, in severe cases, cause total yield loss for cotton growers (Pozo-Valdivia et al., 2021). Young shoots and leaves can also be damaged, especially in the absence of fruiting structures.

Young maize plants have serial holes in the leaves following whorl feeding on the apical leaf. On larger plants the silks are grazed and eggs can be found stuck to the silks. As the ears develop, the soft milky grains in the top few centimetres of the cobs are eaten; usually only one large larva per cob can be seen because the larvae are cannibalistic. Ear damage is often localized to the tip but can increase the incidence of disease.

Sorghum heads are grazed. Legume pods are holed and the seeds eaten. Bore holes can be seen in tomato fruits, cotton bolls, cabbage and lettuce hearts, soyabean pods and flower heads.

List of Symptoms/Signs

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SignLife StagesType
Fruit / external feeding
Fruit / internal feeding
Growing point / external feeding
Growing point / internal feeding; boring
Inflorescence / external feeding
Inflorescence / internal feeding
Leaves / external feeding
Seeds / external feeding
Seeds / internal feeding

Biology and Ecology

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Eggs are laid mostly on the silks of maize plants in small numbers (one to three), stuck to the plant tissues. However, in cotton, they are mostly laid on the leaves throughout the plant, but are more common near blooms (Braswell et al., 2019a, b). Choice of oviposition site by the female seems to be governed by a combination of physical and chemical cues. Female fecundity can be dependent upon the quality and quantity of larval food, and also on the quality of adult nutrition. Up to 3000 eggs have been laid by a single female in captivity, but a few hundred to a thousand per female is more usual in the wild (Quaintance and Brues, 1905; Bilbo et al., 2018). Hatching occurs after 2-4 days and the eggs change colour from green through red to grey. The tiny grey larvae first eat the eggshell and most of the newly hatched larvae disperse from the plant or die (Zalucki et al., 2002; Braswell et al., 2019a, b). Those that remain wander actively for a while before starting to feed on the plant. In maize, they usually feed on the silks initially and then on the young tender kernels after entering the tip of the husk. By the third instar the larvae become cannibalistic and usually only one larva survives per cob. Feeding damage is typically confined to the tip of the cob. In cotton, small larvae initially feed on squares (flower buds), but they can consume small bolls. Once larvae gain a sufficient size, they begin to feed on larger sized bolls (Braswell et al., 2019a). Larval development usually takes 14-25 (mean 16) days, but under cooler conditions up to 60 days may be required. In the final instar (usually sixth) feeding ceases and the fully fed caterpillar leaves the cob and descends to the ground. It then burrows into the soil and forms an earthen cell, where it rests in a prepupal state for a day or two, before finally pupating. Two basic types of pupal diapause are recognized, one in relation to cold and the other in response to arid conditions. In the tropics, pupation takes 10-14 (mean 13) days; the male takes 1 day longer than the female. Diapausing pupae are viable as far north as 40-45°N in the USA.

Adults are nocturnal in habit and emerge in the evenings. Maize fields in the USA regularly produce 40,000 to 50,000 adult moths per hectare. Flying adults respond to light radiation at night and are attracted to light traps (Hardwick, 1968), especially the ultraviolet type, in company with many other local noctuids. Sex aggregation pheromones have been identified and synthesized for most of the Heliothis/Helicoverpa pest species, and pheromone traps can be used for population monitoring. Adult longevity is recorded as being about 17 days in captivity; they drink water and feed on nectar from both floral and extrafloral nectaries. The moths fly strongly and are regular seasonal migrants, flying hundreds of kilometres from the USA into Canada. They migrate by flying high with prevailing wind currents.

The life cycle can be completed in 28-30 days at 25°C and in the tropics there may be up to 10-11 generations per year. All stages of the insect are to be found throughout the year if food is available, but development is slowed or stopped by either drought or cold. In the northern USA there are only two generations per year, in Canada only one generation.

For more information, see Hardwick (1965)Beirne (1971)Balachowsky (1972)Allemann (1979)King and Saunders (1984) and Fitt (1989).

Natural enemies

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Natural enemyTypeLife stagesSpecificityReferencesBiological control inBiological control on
Actinoplagia koehleri Parasite
Agelaius phoeniceus Predator
Archytas incertus Parasite
Archytas marmoratus Parasite Larvae|Pupae
Archytas platonicus Parasite
Archytas scutellatus Parasite
Aspergillus niger Antagonist
Ateloglutus chilensis Parasite
Athrycia cinerea Parasite
Bacillus circulans Pathogen Arthropods|Larvae
Bacillus subtilis Pathogen Arthropods|Larvae
Bacillus thuringiensis Pathogen Arthropods|Larvae
Bacillus thuringiensis serovar. alesti Pathogen Arthropods|Larvae
Bacillus thuringiensis serovar. israelensis Pathogen Arthropods|Larvae
Bacillus thuringiensis serovar. kurstaki Pathogen Arthropods|Larvae
Bacillus thuringiensis serovar. thuringiensis Pathogen Arthropods|Larvae
Beauveria bassiana Pathogen
Boettcheria latisterna Parasite
Brachymeria incerta Parasite
Brachymeria ovata Parasite Arthropods|Pupae
Brachymeria robusta Parasite
Bracon mellitor Parasite
Bracon platynotae Parasite
Calleida decora Predator Arthropods|Larvae
Calosoma sayi Predator Arthropods|Larvae
Campoletis argentifrons Parasite
Campoletis flavicincta Parasite
Campoletis grioti Parasite
Campoletis sonorensis Parasite
Carcelia illota Parasite
Cardiochiles seminiger Parasite Arthropods|Larvae
Chaetogaedia analis Parasite
Chaetogaedia monticola Parasite
Chelonus blackburni Parasite Arthropods|Larvae
Chelonus curvimaculatus Parasite Arthropods|Larvae
Chelonus insularis Parasite Arthropods|Larvae
Chelonus narayani Parasite Arthropods|Larvae Hawaii
Chetogena claripennis Parasite
Chetogena edwardsii Parasite Arthropods|Larvae
Chetogena floridensis Parasite
Chetogena omissa Parasite
Chetogena tachinomoides Parasite
Chrysoperla carnea Predator
Chrysoperla rufilabris Predator
Coleomegilla maculata Predator
Collops quadrimaculatus Predator
Compsilura concinnata Parasite
Conura igneoides Parasite
Cotesia congregata Parasite Arthropods|Larvae
Cotesia kazak Parasite Arthropods|Larvae
Cotesia marginiventris Parasite Arthropods|Larvae
Cryptus albitarsis Parasite
cytoplasmic polyhedrosis viruses Pathogen Arthropods|Larvae
Diapetimorpha introita Parasite
Dusona lacticincta Parasite
Encarsia porteri Parasite Eggs
Enicospilus concolor Parasite
Entomophaga aulicae Pathogen
Erythemis simplicicollis Predator
Eucelatoria australis Parasite
Eucelatoria bryani Parasite Arthropods|Larvae Hawaii
Eucelatoria rubentis Parasite
Euplectrus comstockii Parasite
Euplectrus platyhypenae Parasite
Exorista mella Parasite
Geocoris punctipes Predator Eggs
Geocoris uliginosus Predator Eggs
Glyptapanteles militaris Parasite
Gonia capitata Parasite
Goniophthalmus halli Parasite
Gymnochaetopsis fulvicauda Parasite
Helicobia rapax Parasite
Helicoverpa armigera nuclear polyhedrosis virus Pathogen Adults|Larvae
Helicoverpa armigera nucleopolyhedrovirus Pathogen
Heliothis nucleopolyhedrosis virus Pathogen
Hippodamia convergens Predator
Hyposoter exiguae Parasite
Hyposoter rivalis Parasite
Ichneumon promissorius Parasite
Incamyia charlini Parasite
Incamyia spinicosta Parasite
Iridovirus Pathogen Arthropods|Larvae
Lebia analis Predator Arthropods|Larvae
Lespesia aletiae Parasite Arthropods|Larvae
Lespesia archippivora Parasite Arthropods|Larvae
Lespesia frenchii Parasite
Leuconostoc mesenteroides Pathogen
Linnaemya comta Parasite
Lydella minense Parasite Arthropods|Larvae
Megaselia nigriceps Parasite
Meloboris fuscifemora Parasite
Metaplagia occidentalis Parasite
Metarhizium anisopliae Pathogen
Metavoria orientalis Parasite
Meteorus arizonensis Parasite
Meteorus autographae Parasite Arthropods|Larvae
Meteorus laphygmae Parasite
Microplitis croceipes Parasite Arthropods|Larvae
Microplitis demolitor Parasite Arthropods|Larvae
Microplitis melianae Parasite
Microplitis rufiventris Parasite Arthropods|Larvae
Muscina levida Parasite
Muscina stabulans Parasite
Nabis alternatus Predator
Nabis roseipennis Predator
Nemorilla pyste Parasite Arthropods|Larvae
Netelia spinipes Parasite
Nomuraea rileyi Pathogen Arthropods|Larvae USA; South Carolina soyabeans
Notoxus monodon Predator
Nucleopolyhedrosis virus Pathogen Arthropods|Larvae
Ophion flavidus Parasite
Orius insidiosus Predator Eggs Hawaii
Orius tristicolor Predator
Oxyopes salticus Predator
Paecilomyces tenuipes Pathogen
Palexorista laxa Parasite Arthropods|Larvae
Paniscus Parasite
Paratriphleps laeviusculus Predator USA cotton; maize; tomatoes
Pelegrina galathea Predator
Peleteria pygmaea Parasite
Peucetia viridans Predator
Phidippus audax Predator
Philonthus alumnus Predator
Phyllobaenus pubescens Predator
Plagiomima spinosula Parasite
Podisus maculiventris Predator
Podisus nigrispinus Predator
Podisus placidus Predator
Polistes metricus Predator
Pristomerus pacificus appalachianus Parasite
Pristomerus spinator Parasite
Pseudatomoscelis seriatus Predator
Rogas perplexus Parasite Arthropods|Larvae
Sagaritis provancheri Parasite
Sinophorus eruficinctus Parasite
Siphona plusiae Parasite
Solenopsis invicta Predator
Spallanzania hebes Parasite
Spanagonicus albofasciatus Predator
Spilochalcis femorata Parasite
Staphylococcus epidermidis Pathogen
Steinernema carpocapsae Parasite
Steinernema feltiae Parasite
Steinernema riobrave Parasite
Stenotrophomonas maltophilia Pathogen
Telenomus heliothidis Parasite
Telenomus remus Parasite
Telenomus spodopterae Parasite Eggs
Toxoneuron bicolor Parasite Arthropods|Larvae
Trichogramma atopovirilia Parasite Eggs
Trichogramma brevicapillum Parasite Eggs
Trichogramma chilonis Parasite Eggs
Trichogramma deion Parasite Eggs
Trichogramma evanescens Parasite Eggs
Trichogramma exiguum Parasite Eggs
Trichogramma maltbyi Parasite Eggs
Trichogramma minutum Parasite Eggs
Trichogramma parkeri Parasite Eggs
Trichogramma perkinsi Parasite Eggs
Trichogramma pretiosum Parasite Eggs California; Nicaragua; Nova Scotia; Texas; USA; Texas cotton
Trichogramma thalense Parasite Eggs
Vairimorpha necatrix Pathogen
Vespula pensylvanica Predator
virus-like particles Parasite
Voria aurifrons Parasite
Winthemia quadripustulata Parasite
Winthemia rufiventris Parasite Arthropods|Larvae
Winthemia rufopicta Parasite
Winthemia sinuata Parasite
Zele melleus Parasite
Zelus tetracanthus Predator
Zenillia blanda Parasite

Notes on Natural Enemies

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Kogan et al. (1989) provides a full list of natural enemy records and their relative importance in biological control is discussed in Reay-Jones (2019).

Means of Movement and Dispersal

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Natural Dispersal

Helicoverpa zea is a facultative seasonal nocturnal migrant, and adults migrate in response to poor local conditions for reproduction, when weather conditions are suitable. Three types of movement are practiced by Helicoverpa moths: short-range, long-range and migration. Short-range dispersal is usually within the crop and low over the foliage, and largely independent of wind currents. Long-range flights are higher (up to 10 m), further (1-10 km), and usually downwind, from crop to crop. Migratory flights occur at higher altitudes (up to 1-2 km) and may last for several hours. The moths can be carried downwind hundreds of kilometres; 400 km is not uncommon for such a flight. There is now evidence that many of them originate in Mexico as young adults and migrate northwards into the USA in the early spring. Probably three generations are required to effect the annual displacement from Mexico up to southern Ontario. Transatlantic dispersal is clearly a possibility for this moth, although it has not yet been demonstrated.

Vector Transmission (biotic)

Accidental Introduction

Air-freight transportation of agricultural produce from the New World to Europe is an ever-increasing commercial enterprise, especially with vegetables and ornamentals. Almost every year, caterpillars of H. zea are intercepted on this produce in the UK (Seymour, 1978).

Pathway Causes

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CauseNotesLong DistanceLocalReferences
Crop production Yes Yes
Cut flower trade Yes Yes

Pathway Vectors

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VectorNotesLong DistanceLocalReferences
Aircraft Yes Yes
Bulk freight or cargo Yes Yes

Plant Trade

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Plant parts liable to carry the pest in trade/transportPest stagesBorne internallyBorne externallyVisibility of pest or symptoms
Flowers/Inflorescences/Cones/Calyx Arthropods/Eggs Yes Pest or symptoms not visible to the naked eye but usually visible under light microscope
Flowers/Inflorescences/Cones/Calyx Arthropods/Larvae Yes Yes Pest or symptoms usually visible to the naked eye
Fruits (inc. pods) Arthropods/Eggs Yes Pest or symptoms not visible to the naked eye but usually visible under light microscope
Fruits (inc. pods) Arthropods/Larvae Yes Yes Pest or symptoms usually visible to the naked eye
Leaves Arthropods/Eggs Yes Pest or symptoms not visible to the naked eye but usually visible under light microscope
Leaves Arthropods/Larvae Yes Pest or symptoms usually visible to the naked eye
Stems (above ground)/Shoots/Trunks/Branches Arthropods/Eggs Yes Pest or symptoms not visible to the naked eye but usually visible under light microscope
Stems (above ground)/Shoots/Trunks/Branches Arthropods/Larvae Yes Pest or symptoms usually visible to the naked eye
Plant parts not known to carry the pest in trade/transport
Bark
Bulbs/Tubers/Corms/Rhizomes
Growing medium accompanying plants
Roots
Seedlings/Micropropagated plants
True seeds (inc. grain)
Wood

Impact Summary

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CategoryImpact
Cultural/amenity None
Economic/livelihood Negative
Environment (generally) Positive and negative
Human health Negative

Impact

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In North America it is reported that H. zea is the second most important economic pest species (preceded by codling moth, Cydia pomonella) (Hardwick, 1965). Fitt (1989) quotes the estimated annual cost of damage by H. zea and H. virescens together on all crops in the USA as more than US$ 1000 million, despite the expenditure of US$ 250 million on insecticide application.

Reasons for the success and importance of this agricultural pest include its high fecundity, polyphagous larval feeding habits, high mobility of both larvae locally and adults with their facultative seasonal migration, and a facultative pupal diapause.

Damage is usually serious and costly because of the larval feeding preference for the reproductive structures and growing points rich in nitrogen (for example, maize cobs and tassels, sorghum heads, cotton bolls and buds, etc), and they have a direct influence on yield. Many of the crops attacked are of high value (cotton, maize, tomatoes). If this pest should become established in protected cultivation economic damage could be widespread.

Infestations of maize grown for silage or for grain are not of direct economic importance; losses are typically about 5% and no control measures are taken, but they serve as a focus, or reservoir of infestation. In many areas the first generation is not regarded as a pest (often on Trifolium) and it does not become an economic pest on cultivated crops until the second, third or even fourth generation.

Economic Impact

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In North America, H. zea has long been reported as a major economic pest. During the 1960s, it was noted as being the second most important economic pest species (preceded by codling moth, Cydia pomonella) (Hardwick, 1965). Fitt (1989) quoted the estimated annual cost of damage by H. zea and Heliothis virescens together on all crops in the USA as more than US$ 1000 million, despite the expenditure of US$ 250 million on insecticide application. During 2019, H. zea and H. virescens were estimated at nearly US$ 117 million in costs of control and direct losses in cotton (Cook and Threet, 2020), and over US$ 117 million in costs of control and direct losses in soyabean (Musser et al., 2020).

Reasons for the success and importance of this agricultural pest include its high fecundity, polyphagous larval feeding habits, high mobility of both larvae locally and adults with their facultative seasonal migration and a facultative pupal diapause (Fitt, 1989).

Damage is usually serious and costly because of the larval feeding preference for the reproductive structures and growing points rich in nitrogen (for example, sorghum heads, cotton bolls and buds, soyabean seeds, etc.), and they have a direct influence on yield. Many of the crops attacked are of high value (hemp, tobacco, tomatoes). If this pest should become established in protected cultivation economic damage could be widespread.

Infestations of maize grown for silage or for grain are not of direct economic importance, and no control measures are taken, but they serve as a focus, or reservoir of infestation. In many areas the first generation is not regarded as a pest (often on Trifolium) and it does not become an economic pest on cultivated crops until the second, third or even fourth generation.

Detection and Inspection

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Feeding damage is usually visible and the larvae can be seen on the surface of plants but often they are hidden within plant reproductive tissue (flowers, fruits, etc). Bore holes may be visible, but otherwise it is necessary to cut open the tissue to detect the pest. Because of morphological similarity, it is impossible to distinguish the larvae of H. zea from those of Helicoverpa armigera, already present in the EPPO region. Positive identification is achieved by rearing the larvae and examining the genitalia of the adult.

Similarities to Other Species/Conditions

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The adults are similar in appearance to Helicoverpa armigera but differ in several details in their genitalia (Hardwick, 1965); dissection and slide-mounting are required for specific morphological determination, and some aspects are comparative so that a series of closely related species have to be available for comparison. H. armigera and H. zea can be distinguished using molecular techniques. Where H. armigera is invasive in South America, it can hybridize with H. zea (Anderson et al., 2018). In Brazil, these hybrids are more common in areas with more maize and soyabean production (Cordeiro et al., 2020).

Prevention and Control

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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.

Control

Introduction

Control of H. zea has been advocated in the USA since the middle of the 19th century, and measures fall into two broad categories: those aimed at an overall pest population reduction and others aimed at the protection of a particular crop. In most situations it is now recommended that integrated pest management be used (Bottrell, 1979).

Cultural Control

Various cultural practices can be used to kill the different instars, including deep ploughing, discing and other methods of mechanical destruction, manipulation of sowing dates and use of trap crops.

Biological Control

In many areas, natural control of this pest may be quite effective for most of the time. Insect parasitoids attack the eggs (especially Trichogramma spp.) and larvae, and some predators can be important in reducing pest populations. King and Coleman (1989) discuss the prospects for long-term biological control of Heliothis/Helicoverpa spp., and clearly this should be an important component of any regional IPM programme.

The most frequently tried method of achieving biological control has been by augmentative releases of artificially reared parasites or predators, especially using Trichogramma spp. However, releases in cotton have not been consistently effective against heliothine populations. Microplitis croceipes could be more effective because it is less affected by organophosphate pesticides and synthetic pyrethroids.

There has also been interest in exploiting entomophagous pathogens such as Bacillus thuringiensis and Heliothis NPV. NPV is commercially applied worldwide for control of Helicoverpa species, especially in Australia and Brazil, but increasingly in the USA. A number of pest and beneficial arthropod species can aid in dispersing the virus beyond where it is applied (Black et al., 2019). In cotton, maize and soyabean, transgenic crop varieties expressing the active B. thuringiensis toxin have been used commercially.

Host-Plant Resistance

The development of crop cultivars resistant or tolerant to damage by Helicoverpa spp. has major potential in their management, particularly for communities with few resources. Many crops possess some genetic potential that can be exploited by breeders to produce varieties less subject to pest damage. Resistance can take three basic forms: antixenosis, antibiosis and tolerance. Varieties of crop hosts showing resistance to Helicoverpa spp. have been identified or developed in cotton, chickpeas, soyabean, tomato, maize, sorghum, millet and tobacco.

In maize, resistant genotypes have been identified which have a high concentration of maysin (rhamnosyl-6-C-(4-ketofucosyl)-5,7,3',4'-tetrahydroxyflavone), a C-glycosyl flavone, in silk tissue. Quantitative trait loci for maysin production were identified on chromosomes one (p1) and nine (umc105a) (Byrne et al., 1996).

In cotton, gossypol glands on the calyx crowns of flower buds confers considerable resistance to H. zea (Calhoun et al., 1997).

Transgenic maize, cotton and soyabean containing genes encoding delta-endotoxins from Bacillus thuringiensis (Bt) kurstaki [Bacillus thuringiensis serovar. kurstaki] have been commercialized in various parts of the world. H. zea is targeted with Bt maize and cotton in the USA, and Bt maize, cotton, and soyabean in Brazil. Two types of Cry toxins (in the families Cry1A and Cry2A) have been used extensively, and H. zea has evolved resistance to these toxins in the USA (Dively et al., 2016; Reisig et al., 2018). Increasingly, these Cry traits are being pyramided with Vip3Aa20 in maize and Vip3Aa19 in cotton. H. zea is effectively controlled with the Vip3Aa toxin (Leite et al., 2018; Yang et al., 2020).

There is little knowledge of the interactions between natural enemies of Helicoverpa and host-plant resistance, but it cannot be assumed that resistance will always be compatible with natural control. For example, laboratory tests using resistant tomato plants containing an alkaloid (alpha-tomatine) were found to be toxic to Hyposoter exiguae, a parasite of H. zea. The parasite acquired the alkaloid from its host after the host had ingested the alkaloid (Campbell and Duffey, 1979).

Chemical Control

Chemical control of the larvae has been the most widely used and generally successful method of managing H. zea on most crops, but it is not easy because larvae feed within plant structures. The early history of chemical control of H. zea is given by Hardwick (1965). Pesticide resistance has been known for some years and is quite widespread (Fitt, 1989) especially on cotton crops.

For cotton, chlorantraniliprole is the most efficacious foliar insecticide, because it is systemic in the plant with long-lasting residual (Reisig et al., 2019; Babu et al., 2021). In soyabean, a suite of insecticides, including chlorantraniliprole, emamectin benzoate, indoxacarb, spinosad, spinetoram are effective. Foliar insecticides are not recommended in maize, as H. zea rarely limits yield (Reay-Jones and Reisig, 2014; Bibb et al., 2018; Olivi et al., 2019), however in sweetcorn, multiple sprays, with rotations and mixtures of chlorantraniliprole, pyrethroids, spinosad and spinetoram applications are generally made 3 days after silk emergence and applied on a weekly basis until silks dry down.

Sterile Backcrosses

Sterile male offspring are produced when certain species are crossed, for example, Heliothis subflexa and Heliothis virescens. This fact has been exploited and evaluated on the island of St. Croix, Virgin Islands, where after a three-year release programme, suppression was achieved.

Pheromonal Control

Mating of H. zea was reduced by 50% in a 12 ha maize field treated with hollow fibres containing (Z)-9-tetradecenyl formate (Mitchell and McLaughlin, 1982). Likewise, (Z)-11-hexadecenal, a component of the H. virescens pheromone, reduced the mating of females of H. zea by 85% (Mitchell et al., 1976).

References

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Allemann, D.V., 1979. Maize pests in the USA. In: Maize, [ed. by Hafliger, E.]. Basle, Switzerland: CIBA-GEIGY Ltd. 58-63.

Anderson, C.J., Oakeshott, J.G., Tay, W.T., Gordon, K.H.J., Zwick, A., Walsh, T.K., 2018. Hybridization and gene flow in the mega-pest lineage of moth, Helicoverpa. Proceedings of the National Academy of Sciences of the United States of America, 115(19):5034-5039. DOI: 10.1073/pnas.1718831115

Anon., 1997. Insect control guide. Ohio, USA: Meister Publishing Company. 442 pp.

Babu, A., Reisig, D.D., Pes, M.P., Ranger, C.M., Chamkasem, N., Reding, M.E., 2021. Effects of chlorantraniliprole residual on Helicoverpa zea in Bt and non-Bt cotton. Pest Management Science, 77(5):2367-2374. DOI: 10.1002/ps.6263

Balachowsky, A.S., 1972. Entomologie appliquee a l'agriculture. Tome II. Lepidopteres. Deuxieme volume. Zygaenoidea-Pyraloidea-Noctuoidea [ed. by Balachowsky, A.S.]. Paris, Masson et Cie, France. viii + 1059-1634.

Barber, G.W., 1937. Seasonal availability of food plants of two species of Heliothis in eastern Georgia. Journal of Economic Entomology, 30(1):150-158. DOI: 10.1093/jee/30.1.150

Barber, G.W., 1941. Technical Bulletin. United States Department of Agriculture. Washington, D.C. 791, 17 pp.

Behere, G.T., Tay, W.T., Russell, D.A., Heckel, D.G., Appleton, B.R., Kranthi, K.R., Batterham, P., 2007. Mitochondrial DNA analysis of field populations of Helicoverpa armigera (Lepidoptera: Noctuidae) and of its relationship to H. zea. BMC Evolutionary Biology, 7. https://doi.org/10.1186/1471-2148-1187-1117.

Beirne, B.P., 1971. Pest insects of annual crop plants in Canada. I. Lepidoptera. II. Diptera. III. Coleoptera. In: Memoirs of the Entomological Society of Canada, 78. 124 pp.

Bibb, J.L., Cook, D., Catchot, A., Musser, F., Stewart, S.D., Leonard, B.R., Buntin, G.D., Kerns, D., Allen, T.W., Gore, J., 2018. Impact of corn earworm (Lepidoptera: Noctuidae) on field corn (Poales: Poaceae) yield and grain quality. Journal of Economic Entomology, 111(3):1249-1255. DOI: 10.1093/jee/toy082

Bilbo, T.R., Reay-Jones, F.P.F., Reisig, D.D., Musser, F.R., Greene, J.K., 2018. Effects of Bt corn on the development and fecundity of corn earworm (Lepidoptera: Noctuidae). Journal of Economic Entomology, 111(5):2233-2241. DOI: 10.1093/jee/toy203

Black, J.L., Lorenz, G.M., Cato, A.J., Faske, T.R., Popham, H.J.R., Paddock, K.J., Bateman, N.R., Seiter, N.J., 2019. Field studies on the horizontal transmission potential by voluntary and involuntary carriers of Helicoverpa armigera nucleopolyhedrovirus (Baculoviridae). Journal of Economic Entomology, 112(3):1098-1104. DOI: 10.1093/jee/toz012

Bottrell, D.G., 1979. Guidelines for integrated control of maize pests. In: FAO Plant Production and Protection Paper, 91. Rome, Italy: FAO.

Braswell, L.R., Reisig, D.D., Sorenson, C.E., Collins, G.D., 2019a. Development and dispersal of Helicoverpa zea (Lepidoptera: Noctuidae) on non-Bt and Bt pyramided cotton. Environmental Entomology, 48(2):465-477. DOI: 10.1093/ee/nvz006

Braswell, L.R., Reisig, D.D., Sorenson, C.E., Collins, G.D., 2019b. Helicoverpa zea (Lepidoptera: Noctuidae) oviposition and larval vertical distribution in Bt cotton under different levels of nitrogen and irrigation. Journal of Economic Entomology, 112(3):1237-1250. DOI: 10.1093/jee/toz023

Braswell, L.R., Reisig, D.D., Sorenson, C.E., Collins, G.D., 2019c. Helicoverpa zea (Lepidoptera: Noctuidae) preference for plant structures, and their location, within Bt cotton under different nitrogen and irrigation regimes. Journal of Economic Entomology, 112(4):1741-1751. DOI: 10.1093/jee/toz105

Byrne, P.F., McMullen, M.D., Snook, M.E., Musket, T.A., Theuri, J.M., Widstrom, N.W., Wiseman, B.R., Coe, E.H., 1996. Quantitative trait loci and metabolic pathways: genetic control of the concentration of maysin, a corn earworm resistance factor, in maize silks. Proceedings of the National Academy of Sciences of the United States of America, 93(17):8820-8825. DOI: 10.1073/pnas.93.17.8820

CABI/EPPO, 1998. Distribution maps of quarantine pests for Europe (edited by Smith, I.M., Charles, L.M.F.). Wallingford, UK: CAB International. xviii + 768 pp.

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Distribution References

CABI, Undated. Compendium record. Wallingford, UK: CABI

EPPO, 2022. EPPO Global database. In: EPPO Global database, Paris, France: EPPO. 1 pp. https://gd.eppo.int/

Marchiori C H, Oliveira A M S, Costa M C R, 2002. Insects collected in maize crop in Itumbiara, south of Goiás state, Brazil. (Insetos coletados em cultivar de milho em Itumbiara, sul de Goiás, Brasil.). Arquivos do Instituto Biológico (São Paulo). 69 (Suplemento, Resumos), 233-234. http://www.biologico.sp.gov.br/ARQUIVOS/V69_supl_RE/marchiori.PDF

Murúa M G, Scalora F S, Navarro F R, Cazado L E, Casmuz A, Villagrán M E, Lobos E, Gastaminza G, 2014. First record of Helicoverpa armigera (Lepidoptera: Noctuidae) in Argentina. Florida Entomologist. 97 (2), 854-856. http://www.fcla.edu/FlaEnt/ DOI:10.1653/024.097.0279

Olivares T S, Angulo A O, Badilla Q R, 2012. Taxonomic notes and new register of moths for Easter Island (Lepidoptera: Noctuoidea). Entomological News. 122 (2), 157-164. http://www.bioone.org/loi/entn DOI:10.3157/021.122.0208

Seebens H, Blackburn T M, Dyer E E, Genovesi P, Hulme P E, Jeschke J M, Pagad S, Pyšek P, Winter M, Arianoutsou M, Bacher S, Blasius B, Brundu G, Capinha C, Celesti-Grapow L, Dawson W, Dullinger S, Fuentes N, Jäger H, Kartesz J, Kenis M, Kreft H, Kühn I, Lenzner B, Liebhold A, Mosena A (et al), 2017. No saturation in the accumulation of alien species worldwide. Nature Communications. 8 (2), 14435. http://www.nature.com/articles/ncomms14435

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GISD/IASPMR: Invasive Alien Species Pathway Management Resource and DAISIE European Invasive Alien Species Gatewayhttps://doi.org/10.5061/dryad.m93f6Data source for updated system data added to species habitat list.

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