Helicoverpa zea (bollworm)
Index
- Pictures
- Identity
- Taxonomic Tree
- Notes on Taxonomy and Nomenclature
- Description
- Distribution
- Distribution Table
- Risk of Introduction
- Habitat List
- Hosts/Species Affected
- Host Plants and Other Plants Affected
- Growth Stages
- Symptoms
- List of Symptoms/Signs
- Biology and Ecology
- Natural enemies
- Notes on Natural Enemies
- Means of Movement and Dispersal
- Pathway Causes
- Pathway Vectors
- Plant Trade
- Impact Summary
- Impact
- Economic Impact
- Detection and Inspection
- Similarities to Other Species/Conditions
- Prevention and Control
- References
- Links to Websites
- Distribution Maps
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Top of pagePreferred 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
Top of page- Domain: Eukaryota
- Kingdom: Metazoa
- Phylum: Arthropoda
- Subphylum: Uniramia
- Class: Insecta
- Order: Lepidoptera
- Family: Noctuidae
- Genus: Helicoverpa
- Species: Helicoverpa zea
Notes on Taxonomy and Nomenclature
Top of pageThe 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
Top of pageEgg
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
Top of pageHelicoverpa zea is confined to the New World. It occurs throughout the Americas from Canada to Argentina (International Institute of Entomology, 1993).
Distribution Table
Top of pageThe 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 2022Continent/Country/Region | Distribution | Last Reported | Origin | First Reported | Invasive | Reference | Notes |
---|---|---|---|---|---|---|---|
Europe |
|||||||
Netherlands | Absent, Confirmed absent by survey | ||||||
Russia | Absent, Confirmed absent by survey | ||||||
Slovenia | Absent | ||||||
Switzerland | Absent, Intercepted only | ||||||
United Kingdom | Absent, Intercepted only | Original citation: Seymour and (1978) | |||||
North America |
|||||||
Antigua and Barbuda | Present | ||||||
Bahamas | Present | ||||||
Barbados | Present | ||||||
Bermuda | Present | ||||||
Canada | Present, Localized | ||||||
-British Columbia | Present | ||||||
-Manitoba | Present | ||||||
-New Brunswick | Present | ||||||
-Nova Scotia | Present | ||||||
-Ontario | Present | ||||||
-Quebec | Present | ||||||
-Saskatchewan | Present | ||||||
Costa Rica | Present | ||||||
Cuba | Present | ||||||
Dominica | Present | ||||||
Dominican Republic | Present | ||||||
El Salvador | Present | ||||||
Guadeloupe | Present | ||||||
Guatemala | Present | ||||||
Haiti | Present | ||||||
Honduras | Present | ||||||
Jamaica | Present | ||||||
Martinique | Present, Widespread | ||||||
Mexico | Present, Widespread | ||||||
Montserrat | Present | ||||||
Nicaragua | Present | ||||||
Panama | Present | ||||||
Puerto Rico | Present | ||||||
Saint Kitts and Nevis | Present, Localized | ||||||
Saint Lucia | Present | ||||||
Saint Vincent and the Grenadines | Present | ||||||
Trinidad and Tobago | Present, Widespread | ||||||
U.S. Virgin Islands | Present | ||||||
United States | Present, Widespread | ||||||
-Alabama | Present | ||||||
-Arizona | Present | ||||||
-Arkansas | Present | ||||||
-California | Present | ||||||
-Colorado | Present | ||||||
-Connecticut | Present | ||||||
-Delaware | Present | ||||||
-Florida | Present | ||||||
-Georgia | Present | ||||||
-Hawaii | Present | Introduced | As: Heliothis zea. First reported: ~1930 | ||||
-Idaho | Present | ||||||
-Illinois | Present | ||||||
-Indiana | Present | ||||||
-Iowa | Present | ||||||
-Kansas | Present | ||||||
-Kentucky | Present | ||||||
-Louisiana | Present | ||||||
-Maine | Present | ||||||
-Maryland | Present | ||||||
-Massachusetts | Present | ||||||
-Michigan | Present | ||||||
-Minnesota | Present | ||||||
-Mississippi | Present | ||||||
-Missouri | Present | ||||||
-Montana | Present | ||||||
-Nebraska | Present | ||||||
-Nevada | Present | ||||||
-New Hampshire | Present | ||||||
-New Jersey | Present | ||||||
-New Mexico | Present | ||||||
-New York | Present | ||||||
-North Carolina | Present | ||||||
-North Dakota | Present | ||||||
-Ohio | Present | ||||||
-Oklahoma | Present | ||||||
-Oregon | Present | ||||||
-Pennsylvania | Present | ||||||
-Rhode Island | Present | ||||||
-South Carolina | Present | ||||||
-South Dakota | Present | ||||||
-Tennessee | Present | ||||||
-Texas | Present | ||||||
-Utah | Present | ||||||
-Vermont | Present | ||||||
-Virginia | Present | ||||||
-Washington | Present | ||||||
-West Virginia | Present | ||||||
-Wisconsin | Present | ||||||
-Wyoming | Present | ||||||
South America |
|||||||
Argentina | Present | ||||||
Bolivia | Present | ||||||
Brazil | Present, Widespread | ||||||
-Bahia | Present | ||||||
-Ceara | Present | ||||||
-Distrito Federal | Present | ||||||
-Goias | Present | ||||||
-Mato Grosso | Present | ||||||
-Mato Grosso do Sul | Present | ||||||
-Minas Gerais | Present | ||||||
-Para | Present | ||||||
-Parana | Present | ||||||
-Pernambuco | Present | ||||||
-Rio de Janeiro | Present | ||||||
-Rio Grande do Sul | Present | ||||||
-Roraima | Present | ||||||
-Santa Catarina | Present | ||||||
-Sao Paulo | Present | ||||||
Chile | Present, Widespread | ||||||
-Easter Island | Present | ||||||
Colombia | Present | ||||||
Ecuador | Present, Widespread | ||||||
Falkland Islands | Present, Few occurrences | ||||||
French Guiana | Present | ||||||
Guyana | Present | ||||||
Paraguay | Present, Widespread | ||||||
Peru | Present | ||||||
Suriname | Present | ||||||
Uruguay | Present, Widespread | ||||||
Venezuela | Present |
Risk of Introduction
Top of pageHelicoverpa 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
Top of pageCategory | Sub-Category | Habitat | Presence | Status |
---|---|---|---|---|
Terrestrial | Managed | Cultivated / agricultural land | Present, no further details | Harmful (pest or invasive) |
Terrestrial | Managed | Protected agriculture (e.g. glasshouse production) | Present, no further details | Harmful (pest or invasive) |
Terrestrial | Managed | Managed forests, plantations and orchards | Present, no further details | Harmful (pest or invasive) |
Terrestrial | Managed | Managed grasslands (grazing systems) | Present, no further details | Natural |
Terrestrial | Managed | Disturbed areas | Present, no further details | Natural |
Terrestrial | Managed | Rail / roadsides | Present, no further details | Natural |
Terrestrial | Natural / Semi-natural | Natural forests | Present, no further details | Natural |
Terrestrial | Natural / Semi-natural | Natural grasslands | Present, no further details | Natural |
Terrestrial | Natural / Semi-natural | Riverbanks | Present, no further details | Natural |
Hosts/Species Affected
Top of pageHelicoverpa 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
Top of pageSymptoms
Top of pageFruiting 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
Top of pageSign | Life Stages | Type |
---|---|---|
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
Top of pageEggs 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
Top of pageNatural enemy | Type | Life stages | Specificity | References | Biological control in | Biological 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
Top of pageKogan 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
Top of pageNatural 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
Top of pageCause | Notes | Long Distance | Local | References |
---|---|---|---|---|
Crop production | Yes | Yes | ||
Cut flower trade | Yes | Yes |
Pathway Vectors
Top of pageVector | Notes | Long Distance | Local | References |
---|---|---|---|---|
Aircraft | Yes | Yes | ||
Bulk freight or cargo | Yes | Yes |
Plant Trade
Top of pagePlant parts liable to carry the pest in trade/transport | Pest stages | Borne internally | Borne externally | Visibility 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
Top of pageCategory | Impact |
---|---|
Cultural/amenity | None |
Economic/livelihood | Negative |
Environment (generally) | Positive and negative |
Human health | Negative |
Impact
Top of pageReasons 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
Top of pageIn 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
Top of pageFeeding 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
Top of pageThe 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
Top of pageDue 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
Top of pageAnon., 1997. Insect control guide. Ohio, USA: Meister Publishing Company. 442 pp.
Chiang, H.C., 1978. Pest management in corn. Annual Review of Entomology, 23:101-123.
Neunzig, H.H., 1963. Wild host plants and parasites. Journal of Economic Entomology, 52:135-139.
Neunzig, H.H., 1969. Technical Bulletin, North Carolina Agricultural Experiment Station. 196, 76 pp.
Distribution References
CABI, Undated. Compendium record. Wallingford, UK: CABI
Links to Websites
Top of pageWebsite | URL | Comment |
---|---|---|
GISD/IASPMR: Invasive Alien Species Pathway Management Resource and DAISIE European Invasive Alien Species Gateway | https://doi.org/10.5061/dryad.m93f6 | Data source for updated system data added to species habitat list. |
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