Invasive Species Compendium

Detailed coverage of invasive species threatening livelihoods and the environment worldwide


Agrotis ipsilon
(black cutworm)



Agrotis ipsilon (black cutworm)


  • Last modified
  • 14 July 2018
  • Datasheet Type(s)
  • Invasive Species
  • Pest
  • Natural Enemy
  • Preferred Scientific Name
  • Agrotis ipsilon
  • Preferred Common Name
  • black cutworm
  • Taxonomic Tree
  • Domain: Eukaryota
  •   Kingdom: Metazoa
  •     Phylum: Arthropoda
  •       Subphylum: Uniramia
  •         Class: Insecta
  • Summary of Invasiveness
  • A. ipsilon is already endemic in all areas of the world where it can thrive.

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Top of page
Black cutworm larva with a cut maize plant.
CaptionBlack cutworm larva with a cut maize plant.
CopyrightArmon J. Keaster
Black cutworm larva with a cut maize plant.
LarvaBlack cutworm larva with a cut maize plant.Armon J. Keaster
Black cutworm moth.
CaptionBlack cutworm moth.
CopyrightArmon J. Keaster
Black cutworm moth.
AdultBlack cutworm moth.Armon J. Keaster
Black cutworm moth on lilac.
TitleAdult on lilac
CaptionBlack cutworm moth on lilac.
CopyrightArmon J. Keaster
Black cutworm moth on lilac.
Adult on lilacBlack cutworm moth on lilac.Armon J. Keaster


Top of page

Preferred Scientific Name

  • Agrotis ipsilon (Hufnagel, 1766)

Preferred Common Name

  • black cutworm

Other Scientific Names

  • Agrotis aureolum Schaus, 1898
  • Agrotis bipars Walker, 1857
  • Agrotis frivola Wallengren, 1860
  • Agrotis pepoli Bertolini, 1974
  • Agrotis spinula
  • Agrotis suffusa (Schiffermiller)
  • Agrotis telifera Donzel, 1837
  • Agrotis ypsilon Hufnagel
  • Bombyx idonea Cramer, 1780
  • Bombyx spinula Esper, 1786
  • Euxoa ipsilon Hufnagel
  • Euxoa ypsilon Hufnagel
  • Exarnis ypsilon (Hübner)
  • Feltia ipsilon Hufnagel
  • Feltia ypsilon Hufnagel
  • Lycophotia ypsilon
  • Noctua aneituma Walker, 1865
  • Noctua suffusa Denis & Schffermuller, 1775
  • Noctua ypsilon (S.A. von Rottenberg, 1776)
  • Peridroma suffusa (Butler)
  • Phalaena ipsilon Hufnagel
  • Phalaena ipsilon Hufnagel, 1766
  • Phalaena ypsilon (Cramer)
  • Phalaena ypsilon Rottenberg, 1776
  • Rhyacia ipsilon Hufnagel
  • Rhyacia pernigrata Warren, 1912
  • Rhyacia ypsilon (Rottenberg)
  • Scotia ipsilon Hufnagel
  • Scotia ypsilon (Hufnagel)

International Common Names

  • English: dark sword grass moth; gram cutworm; greasy cutworm; liance rustic; overflow worm; silver y-moth; tobacco cutworm; ypsilon dart
  • Spanish: gusano cortador negro; gusano grasiento (Arg); gusano trozador (Mexico); trozador
  • French: noctuelle ypsilon; ver gris-noir
  • Portuguese: lagarta rosea

Local Common Names

  • Germany: Eulen, Ypsilon-
  • Indonesia: ulat tanah
  • Italy: nottua dei legumi; nottua della canapa
  • Japan: tamana-yaga
  • Netherlands: zwartbruine aardrups
  • Turkey: bozkurt

EPPO code

  • AGROYP (Agrotis ipsilon)

Summary of Invasiveness

Top of page A. ipsilon is already endemic in all areas of the world where it can thrive.

Taxonomic Tree

Top of page
  • Domain: Eukaryota
  •     Kingdom: Metazoa
  •         Phylum: Arthropoda
  •             Subphylum: Uniramia
  •                 Class: Insecta
  •                     Order: Lepidoptera
  •                         Family: Noctuidae
  •                             Genus: Agrotis
  •                                 Species: Agrotis ipsilon

Notes on Taxonomy and Nomenclature

Top of page The genus Agrotis was named by Ochsenheimer in 1816. The current scientific name for this pest is Agrotis ipsilon (Hufnagel). Many references to Agrotis ypsilon can be found in the literature. Other non-preferred scientific names exist (see list) and can be found from time to time in the older literature, but none occurs so frequently as does A. ypsilon.


Top of page Eggs

Eggs are ribbed and about 0.45 mm high. Newly laid eggs are whitish yellow and become darker as hatching approaches.


The general body colour of the larvae is usually uniform above the spiracles and varies from light grey to black without distinct stripes or markings. The subventral and ventral areas are lighter in colour, with numerous pale flecks. The abdominal segments are nearly equal in width, and there is an indistinct, narrow, pale, mid-dorsal stripe. The head is pale-brownish with black coronal stripes and reticulation (early instars bear dark brown freckles instead of coronal stripes and dark reticulation). The skin bears convex, rounded, distinctly isolated, coarse granules with smaller granules interspersed between the larger granules. The spiracles are black. The setigerous tubercles on the abdomen are large, with the anterior dorsal tubercle only one-third as large as the posterior dorsal tubercle; they are heavily pigmented with black.

There are six or seven instars (usually six). Third instars are approximately 7 mm, fourth instars approximately 10-12 mm, fifth instars approximately 20-30 mm, and sixth or seventh instars approximately 35-50 mm.

The diagnostic characteristics of these larvae are the heterogeneous, convex granules and the relatively large D2 tubercles.


Pupae are brown to dark brown and approximately 17-25 mm in length and 5-6 mm in width. Pupae appear almost black in colour just before the moth emerges.


The forewings are long and narrow, darker than the hindwings and marked with black dashes or 'daggers': the basal two-thirds of the forewing is dark, with the outer third pale grey to brown; orbicular is tear-shaped; reniform has a distinct black wedge- or dagger-shaped black marking on its outer margin; claviform is small, dark, oblong, and filled with dark scales. There is a zigzag line of pale scales on a dark background in the subterminal area. The male antennae are plumose (feathered), and the female antennae are filiform. The wingspread is approximately 35-50 mm.


Top of page A. ipsilon is one of the most widely distributed species in the cutworm complex. It is generally considered to be worldwide in distribution (see Distribution Map).

Distribution Table

Top of page

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

Continent/Country/RegionDistributionLast ReportedOriginFirst ReportedInvasiveReferenceNotes


AfghanistanPresentCIE, 1969
ArmeniaPresentSafaryan, 1986; Azaryan and Sevumyan, 1971
AzerbaijanPresentDruzhelyubova, 1976; Abdinbekova and Akhmedov, 1981
BangladeshPresentAlam and Ahmad, 1975; APPPC, 1987; Islam et al., 1991
CambodiaPresentWaterhouse, 1993
ChinaPresentCIE, 1969; Chiang, 1977; APPPC, 1987
-AnhuiPresentChen, 1990
-GansuPresentXu and He, 2000
-GuangdongPresentWu, 1977
-GuangxiPresentZhou et al., 1991
-HebeiPresentAnon., 1977
-HenanPresentZhang, 1982
-Hong KongPresentCIE, 1969; APPPC, 1987
-HubeiPresentChen et al., 1989
-HunanPresentZhou and Chen , 2004; Li and Xiong, 2005
-HunanPresentZhou and Chen , 2004; Li and Xiong, 2005
-ShaanxiPresentXu and He, 2000
-ShandongPresentWang et al., 1985; Wang et al., 1993
-ShanxiPresentWang et al., 1988
-SichuanPresentLiu et al., 2009
-TibetPresentCIE, 1969
-XinjiangPresentZhao, 1985; Anon, 1976
-YunnanPresentYang et al., 1991
-ZhejiangPresentYe et al., 2000
IndiaPresentPresent based on regional distribution.
-Andhra PradeshPresentMurthy et al., 1982
-AssamPresentBorah et al., 1982
-BiharPresentSinha et al., 1979; Das and Ram, 1988
-DelhiPresentBhattacherjee & Gupta, 1971; Prasad et al., 1983
-GujaratPresentChari and Patel, 1972; Patel et al., 1991
-HaryanaPresentVerma et al., 1972; Yadav et al., 1984
-Himachal PradeshWidespreadThakur & Kashyap, 1992; Sukumaran, 1987
-Indian PunjabPresentSingh, 1977; Gill, 1987
-Jammu and KashmirPresentCIE, 1969; Bhat et al., 1994
-KarnatakaPresentSingh, 1986; Mutalikdesai et al., 1973
-Madhya PradeshPresentSaxena & Rawat, 1968; Meshram et al., 1990
-MaharashtraPresentPatil and Pokharkar, 1979; Patil et al., 1991
-OdishaPresentSontakke et al., 1989
-RajasthanPresentKhan and Sharma, 1971
-Tamil NaduPresentSantharam & Kumaraswami,1984; Abraham et al., 1972
-TripuraPresentDas, 1988
-Uttar PradeshPresentSrivastava et al., 1975; Nag and Nath, 1990
-UttarakhandPresentBisht et al., 2005
-West BengalPresentPramanik and Basu, 1971; Mandal and Biswas, 1992
IndonesiaWidespreadUhan, 1989; CIE, 1969; APPPC, 1987; Waterhouse, 1993
-JavaPresentVan Der Goot, 1924; Franssen, 1936
-SulawesiPresentVoute, 1937; Franssen, 1935
IranWidespreadCIE, 1969; Nikkhoo and Moiini, 1991
IraqPresentCIE, 1969
IsraelPresentCIE, 1969
JapanPresentCIE, 1969; Oku and Kobayashi, 1978; Hirai, 1991
-HokkaidoPresentTsutsui et al., 1985; CIE, 1969; Goto et al., 1986
-HonshuPresentOku et al., 1975; Saito & Fuse, 1987; CIE, 1969
JordanPresentCIE, 1969
Korea, DPRPresentCIE, 1969
Korea, Republic ofPresentKim & Kim,1981; CIE, 1969; Lee et al., 1970; APPPC, 1987
LebanonPresentCIE, 1969
MalaysiaPresentCIE, 1969; Waterhouse, 1993
MyanmarPresentWaterhouse, 1993
PakistanWidespreadHashmi et al., 1973; CIE, 1969; Baloch, 1976
PhilippinesPresentBarron & Litsinger,1987; CMI,1969; Waterhouse, 1993
Saudi ArabiaPresentCIE, 1969
SingaporePresentWaterhouse, 1993; AVA, 2001
Sri LankaPresentCIE, 1969
SyriaPresentCIE, 1969
TaiwanPresentWang, 1982; CIE, 1969; Liu and Yang, 1987
ThailandPresentCIE, 1969; APPPC, 1987; Waterhouse, 1993
TurkeyPresentSengonca, 1982; Turhan et al., 1983; CIE, 1969
United Arab EmiratesPresentKaakeh et al., 2007
VietnamPresentCIE, 1969; APPPC, 1987; Waterhouse, 1993


BeninPresentCIE, 1969
Burkina FasoPresentCIE, 1969
CongoPresentFoscolo and Lefevre, 1939; CIE, 1969
Côte d'IvoirePresentCIE, 1969
EgyptWidespreadAwadallah et al., 1976; Nasr et al., 1980
Equatorial GuineaPresentCIE, 1969
KenyaPresentCIE, 1969; Khaemba, 1979; Muruli et al., 1980
LiberiaPresentBreniere, 1976
LibyaPresentKruger, 1934; CIE, 1969
MadagascarPresentCIE, 1969
MalawiPresentBallard, 1913; CIE, 1969
MaliPresentCIE, 1969
MauritiusPresentSugar Ind. Res. Inst., 1987; Dove and Williams, 1971
MoroccoPresentCIE, 1969
RéunionPresentCIE, 1969
Saint HelenaPresentCIE, 1969
SenegalPresentCIE, 1969; Collingwood et al., 1980
South AfricaPresentBrain, 1918; CIE, 1969
SudanPresentSiddig, 1987; CIE, 1969
TogoPresentCIE, 1969
TunisiaPresentHannothiaux, 1965; CIE, 1969
ZimbabwePresentTaylor, 1982; Mitchell et al., 1971; Blair, 1975

North America

CanadaPresentPresent based on regional distribution.
-AlbertaPresentCIE, 1969; Anon, 1979
-British ColumbiaPresentCIE, 1969; Anon, 1979
-ManitobaPresentCIE, 1969; Anon, 1979
-New BrunswickPresentAnon, 1979
-Newfoundland and LabradorPresentAnon, 1979
-Northwest TerritoriesPresentAnon, 1979
-Nova ScotiaPresentSpecht, 1972; CIE, 1969; Anon, 1979
-OntarioPresentCIE, 1969; Harris et al., 1973; Anon, 1979
-Prince Edward IslandPresentAnon, 1979
-QuebecPresentCIE, 1969; Anon, 1979
-SaskatchewanPresentCIE, 1969; Anon, 1979
-Yukon TerritoryPresentAnon, 1979
MexicoWidespreadRodriguez del Bosque & Loera-Gallardo, 1993
USAPresentPresent based on regional distribution.
-AlabamaPresentFoster & Gaylor , 1987
-ArizonaPresentCIE, 1969
-ArkansasPresentSelman & Barton, 1972
-CaliforniaPresentClement et al., 1982
-ColoradoPresentCIE, 1969
-ConnecticutPresentBritton, 1907; CIE, 1969
-DelawarePresentMilliron, 1958; CIE, 1969
-GeorgiaPresentReed, 1915; CIE, 1969
-HawaiiPresentCIE, 1969; Murdoch et al., 1990
-IdahoPresentCIE, 1969
-IllinoisPresentSherrod et al., 1979; Story et al., 1984
-IndianaPresentLund and Turpin, 1977; Johnson et al., 1984
-IowaPresentShowers et al., 1985; Levine et al., 1982
-KansasPresentSmith, 1943; CIE, 1969
-KentuckyPresentAnon., 1908; CIE, 1969
-LouisianaPresentJones, 1918; CIE, 1969
-MainePresentHarvey, 1891; CIE, 1969
-MarylandPresentHarrison et al., 1978; Harrison et al., 1980
-MassachusettsPresentFranklin, 1919; CIE, 1969
-MichiganPresentLarson, 1989
-MinnesotaPresentCook, 1920; CIE, 1969
-MississippiPresentKent, 1889; CIE, 1969
-MissouriPresentLevine et al., 1982; Legg and Keaster, 1984
-MontanaPresentCooley, 1906; CIE, 1969
-NebraskaPresentMuma, 1946; CIE, 1969
-NevadaPresentCIE, 1969
-New HampshirePresentCIE, 1969
-New JerseyPresentWillson et al., 1981
-New MexicoPresentCIE, 1969
-New YorkPresentWillson et al., 1981
-North CarolinaPresentKulash, 1958; CIE, 1969
-North DakotaPresentCIE, 1969
-OhioPresentWillson & Eisley, 1992; Levine et al., 1982
-OklahomaPresentCIE, 1969
-OregonPresentMiller and West, 1987
-PennsylvaniaPresentTietz, 1951; CIE, 1969
-Rhode IslandPresentCIE, 1969
-South CarolinaPresentCIE, 1969
-South DakotaPresentCIE, 1969
-TennesseePresentStanley, 1965; CIE, 1969
-TexasPresentSchuster & Boling, 1973
-UtahPresentKnowlton, 1958; CIE, 1969
-VermontPresentPerkins, 1894; CIE, 1969
-VirginiaPresentRoberts & Snider, 1978; Clark et al., 1993
-WashingtonPresentCIE, 1969
-West VirginiaPresentCIE, 1969
-WisconsinPresentCIE, 1969
-WyomingPresentScott, 1918; CIE, 1969

Central America and Caribbean

Dominican RepublicPresentSchotman, 1989
HondurasPresentSchotman, 1989

South America

ArgentinaPresentGalarza, 1972; Patruele, 1988
BoliviaPresentSquire, 1972
BrazilWidespreadBarbosa and Hooker, 1983
-Espirito SantoPresentFigueroa & Prado, 1973; Smith et al., 1973
-Minas GeraisPresentBarrigossi et al., 1988
-ParaPresentCosta et al., 1982
-ParanaPresentSantos et al., 1992
-PernambucoPresentLopes, 1990
-Rio de JaneiroPresentCIE, 1969
-Rio Grande do SulPresentLink et al., 1987; Bertels, 1972
-Santa CatarinaPresentCIE, 1969
-Sao PauloPresentSilveira-Neto et al., 1975; Machado et al., 1987
ChilePresentCarillo et al., 1988a; Carillo et al., 1988b; CIE, 1969
ColombiaPresentSchotman, 1989
EcuadorPresentCIE, 1969
PeruPresentCIE, 1969
UruguayPresentZerbino, 1986; CIE, 1969
VenezuelaPresentCIE, 1969


AlbaniaPresentKarsholt and Razowski, 1996
AustriaPresentCIE, 1969; Glaeser, 1971
BelgiumPresentCIE, 1969
BulgariaWidespreadDochkova, 1971; Nikolova, 1971; Dimitrov, 1975
CroatiaPresentBazok, 2007
CyprusPresentCIE, 1969
Czech RepublicPresentKarsholt and Razowski, 1996
Czechoslovakia (former)PresentSpitzer, 1970; CIE, 1969
DenmarkPresentCIE, 1969
EstoniaPresentKarsholt and Razowski, 1996
FinlandPresentCIE, 1969
FrancePresentNaibo,1984; Poitout,1982; Provost,1985; Rahn, 1973
-CorsicaPresentKarsholt and Razowski, 1996
GermanyPresentTerytze et al., 1987; CIE, 1969
GreecePresentCIE, 1969
HungaryPresentHerczig & Shultz, 1985; CIE, 1969; Camprag, 1977
IcelandPresentKarsholt and Razowski, 1996
IrelandPresentKarsholt and Razowski, 1996
ItalyWidespreadSannino & Balbiani, 1987; Zangheri et al., 1984
-SardiniaPresentKarsholt and Razowski, 1996
-SicilyPresentKarsholt and Razowski, 1996
LatviaPresentKarsholt and Razowski, 1996
LithuaniaPresentKarsholt and Razowski, 1996
LuxembourgPresentKarsholt and Razowski, 1996
MacedoniaPresentVasilev, 1987
MaltaPresentKarsholt and Razowski, 1996
NetherlandsPresentCIE, 1969
NorwayPresentCIE, 1969
PolandPresentCIE, 1969; Napiorkowska-Kowalik, 1973
PortugalWidespreadCIE, 1969
-AzoresPresentVieira, 2003
RomaniaWidespreadPeiu, 1977; Ionescu, 1985; Popescu, 1989
Russian FederationPresentKarsholt and Razowski, 1996
-Russia (Europe)PresentPospelov and Pukhaev, 1981
-Russian Far EastPresentDashevskii and Rybakova, 1979
-SiberiaPresentCIE, 1969
SloveniaPresentKarsholt and Razowski, 1996
SpainPresentGarrido et al., 1979; Caballero et al., 1989
SwedenPresentCIE, 1969
SwitzerlandPresentCIE, 1969; Freuler, 1984; Hachler, 1988; Hachler, 1989
UKWidespreadCIE, 1969; Bretherton, 1982
Yugoslavia (former)PresentCIE, 1969; Camprag, 1977
Yugoslavia (Serbia and Montenegro)PresentCIE, 1969


AustraliaPresentFarrow and McDonald, 1987
-New South WalesPresentCIE, 1969; Goodyer, 1985
-QueenslandWidespreadJarvis and Smith, 1946; CIE, 1969; Persson, 1977
-TasmaniaPresentCIE, 1969
-Western AustraliaPresentCIE, 1969
FijiPresentLever, 1938; Donald, 1939; Lever, 1940; Lever, 1943; Lever, 1945; APPPC, 1987
New CaledoniaPresentDelobel and Gutierrez, 1981
New ZealandPresentFox,1976; Pearson & Goldson,1980; Allan, 1975; Allan, 1987
Papua New GuineaPresentAPPPC, 1987

History of Introduction and Spread

Top of page A. ipsilon was initially described from specimens caught in Austria in 1766. Specimens in the Macleay Museum indicate that it was already present in Australia circa 1850. It is surmised that the species was already ubiquitous before records began.


Top of page Outbreaks of A. ipsilon frequently occur on wet soils or on ground that has been recently flooded. River bottoms or low areas in fields are often very susceptible to cutworm infestations. For this reason, this species has been known in some localities of the USA as the overflow worm.

Hosts/Species Affected

Top of page Agrotis ipsilon has a very wide host range, but seedling crop plants are most seriously damaged. It also feeds on grasses and weeds. In addition to the plants listed, which are mostly of economic importance, A. ipsilon has also been reared in the laboratory on the following species: amaranth, annual morning glory (Pharbitis purpurea), bluegrass (Poa pratensis), castor (Ricinus communis), Chenopodium album, creeping thistle, curled dock (Rumex crispus), dandelion (Taraxacum officinale), mint (Mentha pulegium), purple nettle (Lamium purpureum), red clover (Trifolium pratense), thornapple (Datura stramonium), violet (Viola odorata), and wormwood (Artemisia spp.).

Host Plants and Other Plants Affected

Top of page
Plant nameFamilyContext
Abelmoschus esculentus (okra)MalvaceaeMain
Agrostis (bentgrasses)PoaceaeWild host
Allium cepa (onion)LiliaceaeMain
Apium graveolens (celery)ApiaceaeMain
Arachis hypogaea (groundnut)FabaceaeMain
Asparagus officinalis (asparagus)LiliaceaeMain
Atropa belladonna (deadly nightshade)SolanaceaeWild host
Avena sativa (oats)PoaceaeMain
Beta vulgaris var. saccharifera (sugarbeet)ChenopodiaceaeMain
Brassica napus var. napus (rape)BrassicaceaeMain
Brassica nigra (black mustard)BrassicaceaeMain
Brassica oleracea (cabbages, cauliflowers)BrassicaceaeMain
Brassica oleracea var. gongylodes (kohlrabi)BrassicaceaeMain
Brassica oleracea var. italica (broccoli)BrassicaceaeMain
Brassica rapa subsp. chinensis (Chinese cabbage)BrassicaceaeMain
Brassica rapa subsp. rapa (turnip)BrassicaceaeMain
Brassicaceae (cruciferous crops)BrassicaceaeMain
Camellia sinensis (tea)TheaceaeMain
Capsicum annuum (bell pepper)SolanaceaeMain
Carthamus tinctorius (safflower)AsteraceaeMain
Chenopodium quinoa (quinoa)ChenopodiaceaeOther
Cicer arietinum (chickpea)FabaceaeMain
Citrullus lanatus (watermelon)CucurbitaceaeMain
Citrus sinensis (navel orange)RutaceaeMain
Coffea (coffee)RubiaceaeMain
Convolvulus arvensis (bindweed)ConvolvulaceaeWild host
Cucumis sativus (cucumber)CucurbitaceaeMain
Cucurbita pepo (marrow)CucurbitaceaeMain
Cynara cardunculus var. scolymus (globe artichoke)AsteraceaeMain
Daucus carota (carrot)ApiaceaeMain
Fragaria (strawberry)RosaceaeMain
Ginkgo biloba (kew tree)GinkgoaceaeWild host
Gladiolus hybrids (sword lily)IridaceaeOther
Glycine max (soyabean)FabaceaeMain
Gossypium (cotton)MalvaceaeMain
Helianthus annuus (sunflower)AsteraceaeMain
Hordeum vulgare (barley)PoaceaeMain
Ipomoea batatas (sweet potato)ConvolvulaceaeMain
Lactuca sativa (lettuce)AsteraceaeMain
Lens culinaris subsp. culinaris (lentil)FabaceaeMain
Linum usitatissimum (flax)Main
Malus domestica (apple)RosaceaeMain
Manihot esculenta (cassava)EuphorbiaceaeMain
Medicago sativa (lucerne)FabaceaeMain
Mentha (mints)LamiaceaeWild host
Mentha piperita (Peppermint)LamiaceaeMain
Mentha spicata (Spear mint)LamiaceaeMain
Musa (banana)MusaceaeMain
Nicotiana tabacum (tobacco)SolanaceaeMain
Papaver somniferum (Opium poppy)PapaveraceaeMain
Parthenium argentatum (Guayule)AsteraceaeMain
Phaseolus (beans)FabaceaeMain
Phaseolus vulgaris (common bean)FabaceaeMain
Pisum sativum (pea)FabaceaeMain
Polyphagous (polyphagous)Main
Prunus domestica (plum)RosaceaeOther
Prunus persica (peach)RosaceaeOther
Prunus salicina (Japanese plum)RosaceaeOther
Pyrus communis (European pear)RosaceaeOther
Raphanus sativus (radish)BrassicaceaeMain
Ricinus communis (castor bean)EuphorbiaceaeMain
Saccharum officinarum (sugarcane)PoaceaeMain
Sapium sebiferum (Chinese tallow tree)EuphorbiaceaeMain
Sesamum indicum (sesame)PedaliaceaeMain
Solanum lycopersicum (tomato)SolanaceaeMain
Solanum melongena (aubergine)SolanaceaeMain
Solanum tuberosum (potato)SolanaceaeMain
Sorghum bicolor (sorghum)PoaceaeMain
Stachys arvensis (staggerweed)LamiaceaeOther
Trifolium (clovers)FabaceaeWild host
Trifolium alexandrinum (Berseem clover)FabaceaeMain
Trifolium repens (white clover)FabaceaeMain
Triticum (wheat)PoaceaeMain
Vicia faba (faba bean)FabaceaeMain
Vigna unguiculata (cowpea)FabaceaeMain
Vitis (grape)VitaceaeMain
Zea mays (maize)PoaceaeMain
Zingiber (ginger)ZingiberaceaeMain

Growth Stages

Top of page Seedling stage, Vegetative growing stage


Top of page Early instar A. ipsilon larvae can create 'shotholes' while feeding on tender leaves of seedling plants. Third to seventh instars become negatively phototaxic and feed mostly at night. Damage from these instars is usually observed as a cutting of young seedlings, often causing death of the cut seedlings. Sometimes wilting is observed because of partial cutting. Larvae are destructive out of proportion to the actual plant material they consume because several plants may be cut by a single larva. A larva will often cut one plant, quickly move on to other plants and continue cutting. Relatively small populations of cutworms are capable of destroying entire stands of some crops, such as cotton or maize. When seedlings are too large to be cut, foliar feeding may reduce plant vigour and subsequent yield.

Difficulty in assessing A. ipsilon injury is due to their habit of tunnelling under the soil in the daytime and feeding at night. This characteristic makes determination of damaging larval populations difficult, and a great deal of damage can occur in a relatively short period before an infestation is suspected. As plants become larger, older instars will occasionally tunnel into the growing stalk, disrupting the xylem and phloem. Climbing behaviour in A. ipsilon is not well developed, but when plants become too large to cut or too difficult to tunnel into, larvae will cut off leaves. However, this leaf cutting does not usually cause economic damage to the plant. Fruit piercing may occur in plums, peaches and pears.

List of Symptoms/Signs

Top of page
SignLife StagesType
Fruit / external feeding
Leaves / external feeding
Stems / external feeding
Stems / internal feeding
Stems / lodging; broken stems
Whole plant / dwarfing
Whole plant / external feeding
Whole plant / wilt

Biology and Ecology

Top of page Female black cutworm moths usually deposit eggs singly or a few (as many as 30) together. The eggs are firmly attached to a substrate. Preferred substrates are densely growing plants relatively low to the ground and fine-textured plant debris in untilled fields. Damp, low-lying areas within untilled fields are particularly attractive for egg deposition and larval survival. Egg placement varies with the plant species and may be on the petioles, lower leaf surface, or stem. The eggs hatch in 3-6 days and the larvae move into the soil where they remain during the day.

The sudden appearances of A. ipsilon, often in large numbers, and its seasonal disappearances (especially in India, Sulawesi, Egypt and the Middle East) have long caused this pest to be recognized as a migrant (Odiyo, 1975). In many parts of the world, migration has been suggested as the source for black cutworm moth populations ovipositing on early spring crops (Rivnay, 1964; Meszaros and Nagy, 1968; Johnson, 1969; Odiyo, 1975; Sugimoto and Kobayashi, 1978; Pedgley and Yathom, 1993). Research in the USA has also supported a spring migration as the probable source of black cutworm moths infesting northern latitudes (Carey and Beegle, 1975; Kaster and Showers, 1982; Domino et al., 1983; Clement et al., 1985; Hutchins et al., 1988; Showers et al., 1989a,b). In addition to spring northward migration in the USA, autumn southward migration, combined with delayed reproduction, has also been suggested (Williams, 1926; Williams et al., 1942; Kaster and Showers, 1982; Story and Keaster, 1982a; Clement et al., 1985; Showers et al., 1993).

Female black cutworm moths usually deposit eggs singly or a few (as many as 30) together. The eggs are attached firmly to a substrate. Preferred substrates are densely growing plants relatively low to the ground and fine-textured plant debris in untilled fields. Damp, low-lying areas within untilled fields are particularly attractive for egg deposition and larval survival. Egg placement varies with the plant species and may be on the petioles, lower leaf surface, or stem. The eggs hatch in 3-6 days and the larvae move into the soil where they remain during the day. Larvae move to the surface at night and feed on young plants. There are six or seven instars, depending on temperature and adequacy of diet. Depending on temperature, larvae begin to pupate in 25-35 days. Pupation occurs in the soil at a depth of 2.0 cm to 10.0 cm and the pupal stage lasts about 12-15 days.

There may be several generations a year, but the spring generation is usually the most damaging because of the young vulnerable plants available at this time. When mature, spring-generation adults may stay in the same area, depositing eggs on weeds and grasses in crop lands, pastures, fence rows, vegetable fields, gardens, etc. Depending on geographical location, the mature spring generation may also move farther north on persistent low-level wind jets. Summer-generation adults will again deposit eggs on weeds, grasses, and turf grasses. The succeeding generation (autumn) can be found in habitats similar to those of the previous generations.

A. ipsilon is chiefly a pest of seedling plants. Third to seventh instars become negatively phototaxic, and most feeding in these instars is nocturnal. Larvae construct burrows or tunnels in the soil about 2.5-5 cm deep. During the night, the larva cuts a (seedling) plant(s), drags it into this tunnel and feeds upon it during the day. During the photophase in all these habitats, the larvae will usually seek a soil depth of 2.5-10.0 cm, where a moisture line exists. However, during extremely wet conditions, larvae may exhibit climbing behaviour on vegetation.

In the USA, deteriorating environmental conditions in late summer and early autumn trigger a southward movement on weather systems with near-surface northerly (southward displacement) airflow to the coastal regions of the Gulf of Mexico. Winter survival in the USA can occur north of the soil 0°C isotherm; this line will fluctuate annually in midcontinental USA.

Natural enemies

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Natural enemyTypeLife stagesSpecificityReferencesBiological control inBiological control on
Abacidus permundus Predator Larvae
Apanteles bourquini Parasite Larvae
Archytas marmoratus Parasite Pupae
Bacillus thuringiensis aizawai Pathogen Larvae
Bacillus thuringiensis darmstadiensis Pathogen Larvae
Bacillus thuringiensis entomocidus Pathogen Larvae
Bacillus thuringiensis finitimus Pathogen Larvae
Bacillus thuringiensis galleriae Pathogen Larvae
Bacillus thuringiensis sotto Pathogen Larvae
Bacillus thuringiensis subsp. dendrolimus Pathogen Larvae
Bacillus thuringiensis subtoxicus Pathogen Larvae
Bacillus thuringiensis thompsoni Pathogen Larvae
Bacillus thuringiensis thuringiensis Pathogen Larvae
Bacillus thuringiensis tolworthi Pathogen Larvae
Beauveria bassiana Pathogen
Campoletis argentifrons
Campoletis chlorideae
Campoletis flavicincta Parasite Larvae
Carabidae Predator Larvae
Chelonus inanitus Parasite Larvae
Chrysoperla carnea Predator
Cotesia marginiventris Parasite Larvae
Cotesia ruficrus Parasite Larvae Egypt; Middle East; Sulawesi aubergines; polyphagous; Vicia faba
Cotesia telengai Parasite Larvae
Ctenichneumon panzeri Parasite Larvae
Cyclotrachelus sodalis Predator Larvae
Enicospilus merdarius Parasite
Exorista larvarum Parasite Larvae
Gonia bimaculata Parasite Larvae
Gonia capitata Parasite Larvae
Granulosis virus Pathogen Larvae
Herpetogramma licarsisalis Predator
Heterorhabditis bacteriophora Parasite
Heterorhabditis heliothidis Parasite
Hexamermis arvalis Parasite
Hexamermis arvalis Parasite Larvae
Hexamermis heterocephalis Parasite
Hexamermis heterocephalis Parasite Larvae
Hexamermis tzihuensis Parasite
Hexamermis tzihuensis Parasite Larvae
Labidura riparia Predator
Lespesia archippivora Parasite Larvae
Linnaemya comta Parasite Larvae New Zealand pasture plants
Linnaemya comta Parasite
Linnaemya comta Parasite Larvae
Macrocentrus collaris Parasite Larvae New Zealand pasture plants
Metarhizium anisopliae Pathogen
Meteorus communis Parasite Larvae
Meteorus leviventris Parasite Larvae
Meteorus rubens Parasite Larvae Egypt Vicia faba
Microgaster kewleyi Parasite Larvae
Microplitis feltiae Parasite Larvae
Microplitis kewleyi Parasite Larvae
Microplitis rufiventris Parasite Larvae
Microplitis similis Parasite Larvae
Netelia fuscicornis Parasite Pupae
Noctuidonema guyanense Parasite Adults
Nomuraea rileyi Pathogen
Nucleopolyhedrosis virus Pathogen
Peleteria rubescens Parasite Larvae
Peleteria rubescens Parasite
Peleteria varia Parasite Larvae Sulawesi
Peleteria varia Parasite
Periscepsia carbonaria Parasite Larvae
Periscepsia carbonaria Parasite
Phryxe magnicornis Parasite Larvae
Phryxe vulgaris Parasite Larvae
Pseudogonia rufifrons Parasite Larvae
Pterostichus chalcites Predator Larvae
Serratia marcescens Pathogen
Siphona collini Parasite Larvae
Siphona cristata Parasite Larvae
Steinernema carpocapsae Parasite
Steinernema carpocapsae Parasite
Steinernema feltiae Parasite
Steinernema feltiae Parasite
Steinernema glaseri Parasite
Steinernema glaseri Parasite
Stelopolybia pallipes Predator Adults/Larvae
Telenomus nawaii Parasite Eggs Egypt
Telenomus remus Parasite Eggs
Trichogramma dendrolimi Parasite Eggs China; China; Shanxi cotton
Trichogramma evanescens Parasite Eggs
Trichogrammatoidea bactrae Parasite
Tritaxys heterocera Parasite Larvae Sulawesi
Turanogonia chinensis Parasite Larvae
Vairimorpha necatrix Pathogen
Zele nigricornis Parasite Larvae
Zelus tetracanthus Predator

Notes on Natural Enemies

Top of page A. ipsilon is a migrant moth of cosmopolitan distribution for which numerous parasites and some predators have been recorded. Most natural enemies are generalists and so few have any beneficial impact. Those listed have been studied in some detail and/or recorded more than once. The list also attempts to provide a broad geographical coverage, indicative of the host's distribution. For further information on these and other recorded natural enemies in Egypt, India and Indiana, USA, see El Heneidy et al. (1987), Singh (1982) and Schoenbolm and Turpin (1977), respectively.

In addition to the naturally occurring enemies, laboratory studies have been conducted on potential biological control agents (see Biological Control section).

Means of Movement and Dispersal

Top of page Natural dispersal

A. ipsilon is naturally migratory and takes advantage of prevailing winds to maintain its global spread. There is no evidence that humans or any other species contribute to its ubiquity.

Pathway Vectors

Top of page
VectorNotesLong DistanceLocalReferences
Soil, sand and gravellarvae, pupae in soil Yes

Plant Trade

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Plant parts not known to carry the pest in trade/transport
Fruits (inc. pods)
Growing medium accompanying plants
Seedlings/Micropropagated plants
Stems (above ground)/Shoots/Trunks/Branches
True seeds (inc. grain)

Wood Packaging

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Wood Packaging not known to carry the pest in trade/transport
Loose wood packing material
Processed or treated wood
Solid wood packing material with bark
Solid wood packing material without bark

Impact Summary

Top of page
Animal/plant collections Negative
Animal/plant products Negative
Biodiversity (generally) None
Crop production Negative
Environment (generally) None
Fisheries / aquaculture None
Forestry production Negative
Human health None
Livestock production Negative
Native fauna None
Native flora None
Rare/protected species None
Tourism Negative
Trade/international relations None
Transport/travel None


Top of page A. ipsilon has a very wide host range. Many of the plants attacked are not of economic importance, but damage to seedling maize, many vegetables, cotton, tobacco, turf grasses, and other crops can be economically significant. It is known to damage crops in North, Central and South America, Europe, Asia, Australasia, Oceanania, Africa and the Middle East.

A. ipsilon is a pest on many crops in tropical and subtropical regions all around the world, causing significant losses, in Chile (Carrillo et al., 2001), Brazil (Secchi, 2001), Egypt (Amin and Abdin, 1997), India (Verma and Verma, 2002), Myanmar (Morris and Waterhouse, 2001), Poland (Walczak, 2002), Spain (Amate et al., 1998) and USA (Amin and Abdin, 1997).

Environmental Impact

Top of page There is little evidence of environmental impact.

Impact: Biodiversity

Top of page There is little evidence of any effects on biodiversity.

Social Impact

Top of page A. ipsilon is a pest on playing fields and golf courses.

Detection and Inspection

Top of page Early instars can create 'shotholes' on the tender leaves of seedling plants. Third to seventh instars become negatively phototaxic and feed mostly at night. These instars usually cut young seedlings at the base, often causing death of the cut seedlings. Several plants may be cut by one larva. Fields should be checked for leaf-feeding, cut, wilting, and missing plants. The larvae tunnel under the soil in the daytime and feed at night, and this characteristic makes determination of damaging larval populations difficult. When damaged plants are found, the soil around the base of the plants should be examined for cutworms. Sometimes the larvae can be found beneath clods, in their tunnels, or in soil cracks. As plants become larger, older instars will occasionally tunnel into the growing stalk, disrupting the xylem and phloem. Climbing behaviour is not well developed, but when plants become too large to cut or too difficult to tunnel into, larvae may cut off leaves. Fruit piercing may occur in plums, peaches and pears.

Similarities to Other Species/Conditions

Top of page A. ipsilon can be distinguished from similar species by the convex granules on the abdominal body segments. In addition, tubercle II is transverse and two to three times as large as tubercle I.

Prevention and Control

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Cutworms are among the insects that continue to challenge the best efforts at pest management. The following practices may prove advantageous to some growers:

General Management Tactics

- If possible, avoid planting crops in fields with a known history of cutworm problems.
- Avoid planting crops (especially maize) following longstanding pastures, meadows, lucerne or red clover.
- Plough in the autumn and use shallow tillage to keep down late autumn and early spring vegetation (where conservation practices allow).
- Monitor larvae with larval cutworm bait traps (Story and Keaster, 1982, 1983; Munson et al., 1986).
- Monitor adults to predict attacks (Hachler and Brunetti, 2002).
- Monitor weather to predict attacks (Bhagat and Praveen Sharma, 2000; Zhou and Chen, 2004).
- Low mow grass to remove eggs, disposing of cuttings at a distance (Williamson and Potter, 1997).
- Topdressing with sand does not kill larvae but deters them from travelling (Williamson and Potter, 1997).
- Encourage predators by encouraging their other prey species nearby, e.g. by having conservation strips between fields or golf fairways (Frank and Shrewsbury, 2004).

Host-Plant Resistance

Three genes have been transferred from their original species to crop plants in order to make them resistant to A. ipsilon. One is the cowpea trypsin inhibitor (CpTi). Another is the barley trysin inhibitor (Bti-cme). Another is the Bacillus thuringiensis cry1Ac gene (Bt). These genes each code for the production of a different insecticidal toxin. The resultant crops are termed 'transgenic'. More than one of these genes can be transferred to give increased resistance.

Grass resistant to A. ipsilon was created by Agrobacterium-mediated transformation of Bermudagrass (Cynodon dactylon) with the Bacillus thuringiensis cry1Ac gene (Salehi et al., 2005).

Transgenic Bt+CpTI cotton was resistant to higher instars of A. ipsilon but could not control the pest effectively (Cui et al., 2002).
Transgenic (Bt) maize was found to be no more resistant to A. ipsilon than regular maize (Pilcher at al., 1997).

Transgenic Bt+CpTi tobacco gave 48% mortality to first-instar A. ipsilon larvae (Luo et al., 1999).

Transgenic wheat and tobacco with the barley trypsin inhibitor gene (Bti-cme) were resistant to A. ipsilon (Carbonero et al., 1998).

Studies of predators feeding on A. ipsilon larvae which were feeding on transgenic crops have been ambiguous. The predator Orius albidipennis was found to be both negatively (Hafez et al., 1997) and positively affected. However, daily feeding of four other predatory species with A. ipsilon larvae fed on transgenic cotton displayed a normal feeding function in agreement with a Holling type II functional response (Cui and Xia, 1997).

Chemical Control

Synthetic Insecticides

Clothianidin, a new synthetic chloronicotinyl insecticide, has been found to be effective as a seed treatment against A. ipsilon (Andersch and Schwarz, 2003).

The effectiveness of diazinon 20 EC, quinalphos 25 EC, chlorpyrifos 20 EC, fenitrothion 50 EC, deltamethrin 2.8 EC and malathion 5% dust against A. ipsilon on potatoes was tested in India (Tripathi et al., 2003). Chlorpyriphos 20 EC was the most effective. Chlorpyriphos, quinalphos, cypermethrin, phosalone and carbaryl have also been tested in similar conditions (Mishra, 2002).

Common alum, aluminium potassium sulfate (solid) and aluminium oxide (liquid) were found to be toxic to A. ipsilon larvae, and synergised the effectiveness of other insecticides (Youssef, 1997).

Natural Insecticides

Pills made of powdered Nerium oleander leaves, wheat bran, and cotton seed meal were effective for trapping and killing A. ipsilon on cotton in China (Ma et al., 2003).

Neem products were found effective for young seedlings of maize in India (Viji and Bhagat, 2001a).

A methanol extract of Melia azedarach fruits was found to be toxic to A. ipsilon (Schmidt et al., 1997).

Extract of Bassia muricata was found to be toxic to first-instar larvae (El-Sayed et al., 1998).

Leaf extracts of Lantana, Parthenium, Hyptis and Ipomoea carnea were found to be toxic to A. ipsilon and other pests (Ramesh-Chandra, 2004).

Toxin extracts from the marine bacterium Microcystis aeruginosa and the sea anenome Parasicyonis actinostoloides were found to be toxic to fourth-instar larvae (Nassar, 2000).

Extracts of Tephrosia nubica were found to be effective against against A. ipsilon (Sharaby and Ammar, 1997).

A root extract of Rumex nepalensis was found to be effective against A. ipsilon larvae (Thakur, 1997).

Biological Control

Cell lines of various organs of A. ipsilon larvae have been established for Baculovirus production (Goodman et al., 2001).

Kentucky Bluegrass (Poa pratensis) inhibits the growth of A. ipsilon larvae. Endophyte enhanced perennial ryegrass (Lolium perenne) inhibits foraging behavior. Turfgrass composed of a mixture of these is resistant to A. ipsilon (Richmond and Shetlar, 2001).

Endemic nematodes were investigated in India for effectiveness against A. ipsilon (Hussaini, 2003). Alginate formulations of entomopathogenic nematodes against A. ipsilon caused the maximum mortality (Hussaini et al., 2003). Nematodes should be rehydrated before use (Baur et al., 1997). Fresh manure reduced the effectiveness of nematodes against A. ipsilon on maize (Shapiro et al., 1999). Effectiveness of nematodes is also diminished on grass infected with the endophytic fungus Neotyphodium lolii (Kunkel et al., 2004).

Bacillus thuringiensis was most effective against first- and second-instar larvae of A. ipsilon (Young et al., 2000). Calcium oxide potentiated B. thuringiensis against A. ipsilon on potatoes in Egypt (Salama et al., 1999).

Volatile substances extracted by steam distillation from withered black poplar (Populus nigra) leaves showed strong attractive activity to A. ipsilon and other insects, and could be used in traps (Guo XianRu et al., 2001).


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