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Datasheet

Aphis craccivora
(groundnut aphid)

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Datasheet

Aphis craccivora (groundnut aphid)

Summary

  • Last modified
  • 13 November 2018
  • Datasheet Type(s)
  • Invasive Species
  • Pest
  • Vector of Plant Pest
  • Natural Enemy
  • Preferred Scientific Name
  • Aphis craccivora
  • Preferred Common Name
  • groundnut aphid
  • Taxonomic Tree
  • Domain: Eukaryota
  •   Kingdom: Metazoa
  •     Phylum: Arthropoda
  •       Subphylum: Uniramia
  •         Class: Insecta

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Pictures

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PictureTitleCaptionCopyright
A. craccivora colony on cowpea.
TitleColony
CaptionA. craccivora colony on cowpea.
CopyrightJames Litsinger
A. craccivora colony on cowpea.
ColonyA. craccivora colony on cowpea.James Litsinger
Apterous viviparous A. craccivora females have a shiny black or dark brown body with prominent cauda and brown to yellow legs. Immatures slightly dusted with wax, adults without wax.
TitleAdults and nymphs
CaptionApterous viviparous A. craccivora females have a shiny black or dark brown body with prominent cauda and brown to yellow legs. Immatures slightly dusted with wax, adults without wax.
CopyrightE. Neering
Apterous viviparous A. craccivora females have a shiny black or dark brown body with prominent cauda and brown to yellow legs. Immatures slightly dusted with wax, adults without wax.
Adults and nymphsApterous viviparous A. craccivora females have a shiny black or dark brown body with prominent cauda and brown to yellow legs. Immatures slightly dusted with wax, adults without wax.E. Neering
Alate viviparous A. craccivora females have abdomens with dorsal cross bars. Alatae 1.4-1.9 mm
TitleAlate adult
CaptionAlate viviparous A. craccivora females have abdomens with dorsal cross bars. Alatae 1.4-1.9 mm
CopyrightNatural History Museum, London
Alate viviparous A. craccivora females have abdomens with dorsal cross bars. Alatae 1.4-1.9 mm
Alate adultAlate viviparous A. craccivora females have abdomens with dorsal cross bars. Alatae 1.4-1.9 mmNatural History Museum, London
Apterous adults have six-segmented antennae. Apterae 1.2-2.3 mm.
TitleApterous adult
CaptionApterous adults have six-segmented antennae. Apterae 1.2-2.3 mm.
CopyrightNatural History Museum, London
Apterous adults have six-segmented antennae. Apterae 1.2-2.3 mm.
Apterous adultApterous adults have six-segmented antennae. Apterae 1.2-2.3 mm. Natural History Museum, London
Infestation of A. craccivora and A. fabae (black bean aphid).
TitleInfestation
CaptionInfestation of A. craccivora and A. fabae (black bean aphid).
CopyrightJames Litsinger
Infestation of A. craccivora and A. fabae (black bean aphid).
InfestationInfestation of A. craccivora and A. fabae (black bean aphid).James Litsinger

Identity

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Preferred Scientific Name

  • Aphis craccivora Koch, 1854

Preferred Common Name

  • groundnut aphid

Other Scientific Names

  • Aphis atronitens Cockerell, 1903
  • Aphis beccarii del Guercio, 1917
  • Aphis cistiella Theobald, 1923
  • Aphis citricola del Guercio, 1917
  • Aphis dolichi Montrouzier, 1861
  • Aphis hordei del Guercio, 1913
  • Aphis isabellina del Guercio, 1917
  • Aphis kyberi Hottes, 1930
  • Aphis laburni Theobald
  • Aphis leguminosae Theobald, 1915
  • Aphis loti Kaltenbach, 1862
  • Aphis mimosae Ferrari, 1872
  • Aphis oxalina Theobald, 1925
  • Aphis papilionacearum van der Goot, 1918
  • Aphis robiniae Macchiati, 1885
  • Doralida loti (Kaltenbach)
  • Doralina craccivora (Koch)
  • Doralina salsolae Börner, 1940
  • Doralis laburni (Kaltenbach)
  • Doralis meliloti Börner, 1939
  • Doralis robiniae (Macchiati)
  • Pergandeida craccivora Koch
  • Pergandeida loti (Kaltenbach)
  • Pergandeida robiniae (Macchiati)

International Common Names

  • English: African bean aphid; bean aphid; black legume aphid; black lucerne aphid; cowpea aphid; lucerne aphid; oriental pea aphid
  • Spanish: afido del mata-ratón; pulgón negro (leguminosas, alfalfa, fréjol, caupi)
  • French: puceron de l'arachide; puceron noir (lucerne, gourgane); puceron oriental du pois

Local Common Names

  • Denmark: vikkebladlus
  • Germany: kundebohnen-blattlaus; robinien-blattlaus; schwarze klee-blattlaus
  • Iran: schatte schabdar
  • Israel: knimat hakitnyiot
  • Japan: mame-aburamusi
  • Netherlands: grijszwarte bladluis; zwarte wikkeluis

EPPO code

  • APHICR (Aphis craccivora)

Taxonomic Tree

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  • Domain: Eukaryota
  •     Kingdom: Metazoa
  •         Phylum: Arthropoda
  •             Subphylum: Uniramia
  •                 Class: Insecta
  •                     Order: Hemiptera
  •                         Suborder: Sternorrhyncha
  •                             Unknown: Aphidoidea
  •                                 Family: Aphididae
  •                                     Genus: Aphis
  •                                         Species: Aphis craccivora

Notes on Taxonomy and Nomenclature

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A. craccivora was first described by Koch in 1854. There are a number of synonyms in the literature (Eastop and Hille Ris Lambers, 1976).

Description

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A. craccivora is a relatively small aphid. Apterous viviparous females have a shiny black or dark brown body with a prominent cauda and brown to yellow legs. Immatures are slightly dusted with wax, adults without wax. Six-segmented antennae. Distal part of femur, siphunculi and cauda black. Apterae 1.4-2.2 mm.

Alate viviparous A. craccivora females have abdomens with dorsal cross bars. Alatae 1.4-2.1 mm (Blackman and Eastop, 2000).

Distribution

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Probably palearctic warm temperate in origin, A. craccivora now is virtually worldwide. It is particularly well distributed in the tropics, where it is one of the most common aphid species (CIE, 1983; Blackman and Eastop, 2000). A. craccivora has expanded its distribution in recent decades, now north to Siberia (Russia) and Alberta (Canada), south to Chile and Argentina.

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.

Continent/Country/RegionDistributionLast ReportedOriginFirst ReportedInvasiveReferenceNotes

Asia

AfghanistanPresentCIE, 1983
BangladeshPresentCIE, 1983; APPPC, 1987
Brunei DarussalamPresentWaterhouse, 1993
CambodiaPresentWaterhouse, 1993
ChinaPresentAPPPC, 1987
-GansuPresentLiu et al., 2005
-GuangdongPresentCIE, 1983
-HebeiPresentCIE, 1983
-HeilongjiangPresentCIE, 1983
-HenanPresentCIE, 1983
-Hong KongPresentCIE, 1983
-JiangxiPresentCIE, 1983
-JilinPresentCIE, 1983
-LiaoningPresentCIE, 1983
-Nei MengguPresentCIE, 1983
-ShandongPresentCIE, 1983
-ZhejiangPresentCIE, 1983
GazaPresentCIE, 1983
IndiaPresentGreathead and Greathead, 1992
-Andhra PradeshPresentVenkateswarlu et al., 2003
-AssamPresentPurnima and Dutta, 2002; Shimantini et al., 2016
-BiharPresentKumar et al., 1998
-ChhattisgarhPresentOudhia, 2001
-DelhiPresentCIE, 1983
-GujaratPresentCIE, 1983
-HaryanaPresentKalra et al., 2002
-Himachal PradeshPresentCIE, 1983
-Indian PunjabPresentCIE, 1983
-Jammu and KashmirPresentPandey, 2004
-JharkhandPresentRabindra and Devendera, 2007
-KarnatakaPresentCIE, 1983
-KeralaPresentCIE, 1983
-MaharashtraPresentCIE, 1983
-OdishaWidespreadCIE, 1983
-RajasthanPresentVerma, 2006
-SikkimPresentDeka et al., 2014
-Tamil NaduPresentCIE, 1983
-Uttar PradeshPresentCIE, 1983
-West BengalPresentBiswas et al., 2008
IndonesiaPresentWaterhouse, 1993
-Irian JayaPresentCIE, 1983
-JavaPresentCIE, 1983
-SumatraPresentCIE, 1983
IranPresentCIE, 1983
IraqPresentCIE, 1983
IsraelPresentCIE, 1983
JapanPresentPresent based on regional distribution.
-HokkaidoPresentCIE, 1983
-HonshuPresentCIE, 1983
-KyushuPresentCIE, 1983
JordanPresentCIE, 1983
Korea, DPRPresentCIE, 1983
Korea, Republic ofPresentCIE, 1983; APPPC, 1987
LaosPresentCIE, 1983; Waterhouse, 1993
LebanonPresentCIE, 1983; CIE, 1983
MalaysiaPresentWaterhouse, 1993
-Peninsular MalaysiaPresentCIE, 1983
-SarawakPresentCIE, 1983
MyanmarPresentFAO, 1984; APPPC, 1987; Waterhouse, 1993
NepalPresentCIE, 1983
PakistanWidespreadCIE, 1983
PhilippinesWidespreadCIE, 1983; Waterhouse, 1993
Saudi ArabiaPresentCIE, 1983
SingaporePresentCIE, 1983; APPPC, 1987; Waterhouse, 1993
Sri LankaPresentCIE, 1983
SyriaPresentCIE, 1983
TaiwanPresentCIE, 1983
ThailandPresentCIE, 1983; APPPC, 1987; Waterhouse, 1993
TurkeyPresentCIE, 1983; Akyürek et al., 2011
VietnamPresentCIE, 1983; APPPC, 1987; Waterhouse, 1993
YemenPresentCIE, 1983

Africa

AngolaPresentCIE, 1983
BeninPresentCIE, 1983
CameroonPresentCIE, 1983
Congo Democratic RepublicPresentCIE, 1983
EgyptPresentCIE, 1983
EritreaPresentCIE, 1983
EthiopiaPresentCIE, 1983
GambiaPresentCIE, 1983
GhanaPresentCIE, 1983
GuineaPresentCIE, 1983
KenyaPresentCIE, 1983
LiberiaPresentCIE, 1983
MadagascarPresentCIE, 1983
MalawiPresentCIE, 1983
MauritiusPresentCIE, 1983
MoroccoPresentCIE, 1983
MozambiquePresentCIE, 1983
NigeriaPresentCIE, 1983
RéunionPresentCIE, 1983
Saint HelenaPresentCIE, 1983
Sao Tome and PrincipePresentCIE, 1983
SenegalPresentCIE, 1983
Sierra LeonePresentCIE, 1983
SomaliaPresentCIE, 1983
South AfricaPresentCIE, 1983
Spain
-Canary IslandsPresentCIE, 1983
SudanPresentCIE, 1983
TanzaniaPresentBohlen, 1973; CIE, 1983
TogoPresentCIE, 1983
TunisiaPresentHalima Kamel & Hamouda, 1998
UgandaPresentCIE, 1983
ZambiaPresentCIE, 1983
ZimbabwePresentCIE, 1983

North America

CanadaPresentPresent based on regional distribution.
-AlbertaPresentCIE, 1983
-British ColumbiaPresentCIE, 1983
-QuebecPresentCIE, 1983
MexicoPresentCIE, 1983
USAPresentPresent based on regional distribution.
-ArizonaPresentCIE, 1983
-CaliforniaPresentCIE, 1983
-ColoradoPresentCIE, 1983
-DelawarePresentCIE, 1983
-FloridaPresentCIE, 1983
-GeorgiaPresentCIE, 1983
-HawaiiPresentCIE, 1983
-IdahoPresentCIE, 1983
-IllinoisPresentCIE, 1983
-IndianaPresentCIE, 1983
-IowaPresentCIE, 1983
-KansasPresentCIE, 1983
-LouisianaPresentCIE, 1983
-MarylandPresentCIE, 1983
-MassachusettsPresentCIE, 1983
-MissouriPresentCIE, 1983
-NebraskaPresentCIE, 1983
-New JerseyPresentCIE, 1983
-New MexicoPresentCIE, 1983
-New YorkPresentCIE, 1983
-North CarolinaPresentCIE, 1983
-North DakotaPresentCIE, 1983
-OklahomaPresentCIE, 1983
-OregonPresentCIE, 1983
-PennsylvaniaPresentCIE, 1983
-South CarolinaPresentCIE, 1983
-South DakotaPresentCIE, 1983
-TexasPresentCIE, 1983
-UtahPresentCIE, 1983
-VermontPresentCIE, 1983
-VirginiaPresentCIE, 1983
-WashingtonPresentCIE, 1983
-West VirginiaPresentCIE, 1983
-WisconsinPresentCIE, 1983
-WyomingPresentCIE, 1983

Central America and Caribbean

Antigua and BarbudaPresentCIE, 1983
CubaPresentCIE, 1983
JamaicaPresentMcDonald et al., 2003
NicaraguaPresentCIE, 1983
Puerto RicoPresentCIE, 1983
Saint Kitts and NevisPresentCIE, 1983
Trinidad and TobagoPresentCIE, 1983

South America

ArgentinaPresentCIE, 1983
BoliviaPresentCIE, 1983
BrazilPresentPresent based on regional distribution.
-Espirito SantoPresentMartins et al., 2016
-Rio Grande do SulPresentSturza et al., 2011
-Sao PauloPresentSousa-Silva et al., 1998
ChilePresentCIE, 1983
VenezuelaPresentCIE, 1983

Europe

AustriaPresentCIE, 1983
BelgiumPresentCIE, 1983
BulgariaPresentCIE, 1983
CyprusPresentCIE, 1983
Czechoslovakia (former)PresentCIE, 1983
DenmarkPresentCIE, 1983
FinlandPresentCIE, 1983
FrancePresentCIE, 1983
GermanyPresentCIE, 1983
GreecePresentCIE, 1983
HungaryPresentCIE, 1983
ItalyPresentCIE, 1983
NorwayPresentCIE, 1983
PolandPresentCIE, 1983
PortugalPresentCIE, 1983
RomaniaPresentCIE, 1983
Russian FederationPresentPresent based on regional distribution.
-Central RussiaPresentCIE, 1983
-Russia (Europe)PresentCIE, 1983
-Russian Far EastPresentCIE, 1983
SpainPresentCIE, 1983
SwedenPresentCIE, 1983
UKRestricted distributionCIE, 1983
Yugoslavia (former)PresentCIE, 1983

Oceania

AustraliaPresentPresent based on regional distribution.
-Australian Northern TerritoryPresentCIE, 1983
-New South WalesPresentCIE, 1983
-QueenslandPresentCIE, 1983
-South AustraliaPresentCIE, 1983
-TasmaniaPresentCIE, 1983
-VictoriaPresentCIE, 1983
-Western AustraliaPresentCIE, 1983
FijiPresentCIE, 1983
GuamPresentMoore and Miller, 2002
KiribatiPresentCIE, 1983
Marshall IslandsPresentCIE, 1983
New ZealandPresentCIE, 1983
Papua New GuineaPresentCIE, 1983; APPPC, 1987
SamoaPresentCIE, 1983
Solomon IslandsPresentCIE, 1983; APPPC, 1987
TongaPresentCIE, 1983
TuvaluPresentCIE, 1983

Risk of Introduction

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The main means of A. craccivora dispersal is wind-borne dispersal of the winged forms, with dispersal on plant material being of only minor importance. However, sanitary measures are important within crops to prevent the spread of viruses for which A. craccivora is a vector. Virus-infected planting material offers an easy means of transmission for the virus.

Hosts/Species Affected

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A. craccivora is polyphagous, but with marked preference for Leguminosae, for example, Caragana, Lupinus, Medicago, Melilotus, Robinia, Trifolium and Vicia. It is found in small colonies on many other families, including Cruciferae.

Host Plants and Other Plants Affected

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Plant nameFamilyContext
Albizia julibrissin (silk tree)FabaceaeMain
Amaranthus hybridus (smooth pigweed)AmaranthaceaeWild host
AraceaeAraceaeOther
Arachis hypogaea (groundnut)FabaceaeMain
Brassica rapa subsp. oleifera (turnip rape)BrassicaceaeOther
Cajanus cajan (pigeon pea)FabaceaeMain
Capsicum (peppers)SolanaceaeOther
Chenopodium quinoa (quinoa)ChenopodiaceaeOther
Cicer arietinum (chickpea)FabaceaeOther
CitrusRutaceaeOther
Elettaria cardamomum (cardamom)ZingiberaceaeOther
Fabaceae (leguminous plants)FabaceaeOther
Gleditsia triacanthos (honey locust)FabaceaeMain
Glycyrrhiza (licorice)FabaceaeOther
Gossypium (cotton)MalvaceaeOther
Lablab purpureus (hyacinth bean)FabaceaeOther
Lens culinaris subsp. culinaris (lentil)FabaceaeMain
Lupinus (lupins)FabaceaeOther
Malpighia glabra (acerola)MalpighiaceaeOther
Medicago sativa (lucerne)FabaceaeMain
Momordica charantia (bitter gourd)CucurbitaceaeOther
Moringa oleifera (horse radish tree)MoringaceaeOther
Phaseolus (beans)FabaceaeOther
Phaseolus vulgaris (common bean)FabaceaeOther
Sesamum indicum (sesame)PedaliaceaeOther
Solanum americanumSolanaceaeWild host
Solanum lycopersicum (tomato)SolanaceaeOther
Solanum tuberosum (potato)SolanaceaeOther
Tagetes erecta (African marigold)AsteraceaeOther
Theobroma cacao (cocoa)SterculiaceaeOther
Trifolium (clovers)FabaceaeOther
Trigonella foenum-graecum (fenugreek)FabaceaeOther
Vicia (vetch)FabaceaeOther
Vicia faba (faba bean)FabaceaeOther
Vigna catjangFabaceaeOther
Vigna mungo (black gram)FabaceaeOther
Vigna radiata (mung bean)FabaceaeMain
Vigna unguiculata (cowpea)FabaceaeMain

Growth Stages

Top of page Flowering stage, Seedling stage, Vegetative growing stage

Symptoms

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Groundnut plants take on a bushy appearance due to attack by A. craccivora and infection with rosette virus. Rosette may take two forms, chlorotic rosette (white patches with green veins on young leaves and short internodes) and green rosette (darker appearance with stunting of leaflets and branches).

List of Symptoms/Signs

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SignLife StagesType
Growing point / external feeding
Leaves / abnormal colours
Leaves / abnormal patterns
Leaves / honeydew or sooty mould
Leaves / honeydew or sooty mould
Leaves / honeydew or sooty mould
Leaves / necrotic areas
Leaves / necrotic areas
Whole plant / dwarfing
Whole plant / external feeding

Biology and Ecology

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A. craccivora is anholocyclic almost everywhere. In the tropics it is exclusively anholocyclic, with only females (winged and wingless) normally encountered, and parthenogenetic reproduction occurring throughout the year. The aphid is ovoviviparous, with females retaining eggs inside their bodies and giving birth to small larvae. In areas with colder winters, overwintering may be as egg or hibernation. Males are alate and sexual forms have been recorded from Germany (Falk, 1960), Argentina (la Rossa et al., 1993) and India (Basu et al., 1969). Müller (1977) compares tropical and central European populations.

Young colonies of this small aphid concentrate on growing points of plants and are regularly tended by ants. Mutualism with ants described in India (Soans and Soans, 1971; Patro and Behera, 1991), Pakistan (Hamid et al., 1977) and Japan (Takeda et al., 1982). Hamid et al. (1977) found the ants Pheidole sp. and Monomorium indicum associated with the aphid at 90% of the sites examined; their symbiotic influence appeared to work in favour of the aphid.

Optimal development of A. craccivora is dependent on fairly specific climatic conditions, such as temperatures 24-28.5°C and around 65% RH (Réal, 1955; Mayeux, 1984). In the field, aphids do not generally survive periods of heavy rain. Abdel Malek et al. (1982) showed that optimum daylength for nymphal development was L:D 16:8, while photoperiod did not appear to affect alate production. However, plant chemistry, particularly a reduction in the intensity of hydrocarbon translocation, does influence formation of winged individuals (Mayeux, 1984). They have a preference, in groundnuts, for plants that are not drought stressed (Mayeux, 1984).

A. craccivora is capable of rapid population development. Talati and Butani (1980) investigated reproductive rate on groundnut in the laboratory in India, and observed that offspring from a single gravid adult aphid averaged 17-43 in 15 days. It had four nymphal instars on cowpeas in the laboratory. The total nymphal periods averaged 5.6, 5.1, 5.15 and 4.86 days in May-June, August-September, October-November and March-April, respectively. The durations of the total life cycle during the corresponding periods were 11.07, 11.15, 10.79 and 10.42 days (Patel and Srivastava, 1989). On cowpea, the intrinsic rate of increase in the laboratory was 0.27, with a generation time of 9.78 days (Rani and Remamony, 1998).

Detailed ecological studies of A. craccivora have been done by Réal (1955) on groundnuts in West Africa and Gutierrez et al. (1971, 1974a, b) on pasture in Australia.

Natural enemies

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Natural enemyTypeLife stagesSpecificityReferencesBiological control inBiological control on
Adalia bipunctata Predator Adults/Nymphs
Aphelinus Parasite
Aphelinus asychis Parasite Adults/Nymphs
Aphelinus chaonia Parasite Adults/Nymphs
Aphidius absinthii Parasite
Aphidius colemani Parasite Adults/Nymphs
Aphidius matricariae Parasite Adults/Nymphs
Aphidius smithi Parasite Adults/Nymphs India; Jammu and Kashmir Medicago
Aphidius uzbekistanicus Parasite Adults/Nymphs
Aphidoletes aphidimyza Predator Adults/Nymphs
Ardilea convexa Parasite Adults/Nymphs
Asaphes vulgaris Parasite
Binodoxys nearctaphidis Parasite Adults/Nymphs
Bracon gelechiae Parasite
Brinckochrysa nachoi Predator Adults/Nymphs
Brumoides suturalis Predator Adults/Nymphs
Carabidae Predator Nymphs
Cheilomenes lunata Predator Adults/Nymphs
Cheilomenes propinqua vicina Predator Adults/Nymphs
Cheilomenes sexmaculata Predator Adults/Nymphs Manipur Lablab purpureus; Smithia sensitiva
Chlaenius panagaeoides Predator Adults/Nymphs
Chrysoperla carnea Predator Adults/Nymphs
Coccinella septempunctata Predator Adults/Nymphs India; Manipur Lablab purpureus; Smithia sensitiva
Coccinella septempunctata var. divaricata Predator Adults/Nymphs
Coccinella transversalis Predator Adults/Nymphs India; Manipur Lablab purpureus
Coccinella undecimpunctata Predator Adults/Nymphs
Coelophora inaequalis Predator Adults/Nymphs
Coelophora saucia Predator Adults/Nymphs Manipur
Cycloneda sanguinea Predator Adults/Nymphs
Deraeocoris pallens Predator
Diaeretiella rapae Parasite Adults/Nymphs
Ephedrus plagiator Parasite Adults/Nymphs China; Yunnan Vicia faba
Episyrphus balteatus Predator Adults/Nymphs
Eriopis connexa Predator Adults/Nymphs
Eupeodes confrater Predator Adults/Nymphs
Eupeodes corollae Predator Adults/Nymphs
Fusarium pallidoroseum Pathogen Adults/Nymphs
Harmonia axyridis Predator Adults/Nymphs
Harmonia octomaculata Predator Adults/Nymphs
Hippodamia convergens Predator Adults/Nymphs
Hippodamia variegata Predator Adults/Nymphs
Ischiodon scutellaris Predator Adults/Nymphs India Vicia faba
Leucopis Predator Adults/Nymphs
Lipolexis scutellaris Parasite Adults/Nymphs
Lysiphlebia japonica Parasite Nymphs
Lysiphlebia mirzai Parasite Adults/Nymphs
Lysiphlebus confusus Parasite Adults/Nymphs Lebanon Medicago sativa
Lysiphlebus delhiensis Parasite Adults/Nymphs
Lysiphlebus fabarum Parasite Nymphs Lebanon; Morocco Medicago sativa; Vicia faba
Lysiphlebus marismortui Parasite Adults/Nymphs
Lysiphlebus salicaphis Parasite Adults/Nymphs
Lysiphlebus testaceipes Parasite Nymphs Guadeloupe; India Citrus; Glyricidia maculata; groundnuts
Melanostoma fasciatum Predator
Metarhizium anisopliae Pathogen Pegu et al., 2012
Micraspis discolor Predator Adults/Nymphs
Micromus timidus Predator Adults/Nymphs
Motacilla caspica Predator Adults/Nymphs
Motacilla flava Predator Adults/Nymphs
Neozygites fresenii Pathogen Adults/Nymphs
Olla v-nigrum Predator Adults/Nymphs
Orius minutus Predator Adults/Nymphs China apples; Astragalus sinicus
Paragus serratus Predator Adults/Nymphs
Platynaspis luteorubra Predator Adults/Nymphs
Praon flavinode Parasite Adults/Nymphs
Praon volucre Parasite Adults/Nymphs
Scymnus bicolor Predator Adults/Nymphs
Spilocaria bissellata Predator Adults/Nymphs
Sticholotis substriata Predator Adults/Nymphs
Synonycha grandis Predator Adults/Nymphs India; Manipur Lablab purpureus
Tetragnatha laboriosa Predator
Toxares deltiger Parasite Adults/Nymphs India; Jammu and Kashmir Robinia pseudoacacia
Trioxys angelicae Parasite Nymphs Lebanon Medicago sativa
Trioxys hokkaidensis Parasite Adults/Nymphs
Trioxys indicus Parasite Nymphs Australia; India; Jammu and Kashmir Lupinus; Robinia pseudoacacia
Trioxys jaii Parasite Adults/Nymphs
Trioxys rishii Parasite Adults/Nymphs India; Jammu and Kashmir Robinia pseudoacacia

Notes on Natural Enemies

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Few aphid natural enemies are host-specific, rather they are attracted to aphids in particular habitats. Thus the important natural enemies attacking particular aphid pests on crops tend to vary according to the crop, the circumstances under which it is grown and the climate. This is particularly true of aphid pests attacking a range of different crops over large geographical areas. Besides, many parasitoids are members of species complexes, morphologically very similar but with different host preferences and geographical distributions. The natural enemies in the list represent a selection of species that have been considered as important by investigators and should not be taken as definitive.

Parasitoids reared from A. craccivora have been listed from the Mediterranean basin (Stary, 1976) and Yemen (Stary and Erdelen, 1982). Particularly important parasites of A. craccivora are Trioxys indicus, Lysiphlebus fabarum and L. testaceipes. Attia et al. (1983) reported peak parasitism of 10% on cowpeas in Egypt. Singh and Sinha (1983) reported 9.4% parasitism by T. indicus on pigeonpea in India, shortly after the appearance of A. craccivora leading to peak rates of 64.6% at later stages of infestation. It was sufficient to suppress aphid populations on pigeonpea. Tian et al. (1991) found 13% parasitism by T. indicus and L. japonicum in A. craccivora on cotton.

Alloxysta pleuralis is an important hyperparasite of Trioxys indicus; similarly, Pachyneuron aphidis on Lysiphlebus testaceipes.

Important predators include coccinellid beetles, e.g. Cheilomenes sexmaculata and Coccinella septempuncta, syrphid larvae, e.g. Ischiodon scutellaris, neuropteran larvae, e.g. Micromus timidus, and a predatory dipteran, e.g. Aphidoletes aphidimyza. Spiders may also be important in some areas. A natural enemy complex was described in West Bengal and Karnataka, India, by Ghosh et al. (1981) and Joshi et al. (1997), respectively.

Recorded fungal pathogens include Fusarium pallidoroseum on cowpeas in India (Hareendranath et al., 1987) and Entomophthora fresenii in China (Zhang, 1987). Between 11 and 66.7% of A. craccivora were infected with Neozygites fresenii on faba bean in Egypt during a survey in late 1998 (Sewify, 2000). In a study in India, with F. pallidoroseum applied against A. craccivora on cowpea, yield at 32 days after treatment showed that an increase in spore concentration resulted in a corresponding decrease in aphid population (Sunitha and Mathai, 1999).
 

Impact

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Introduction

A. craccivora is a cosmopolitan species with a worldwide distribution; abundant in subtropical and tropical regions, and in the Mediterranean. It is one of the commonest aphid species in the tropics.

A. craccivora is mainly found on plants in the Leguminosae. It is a major economic pest of groundnut and cowpea, particularly in the tropics, and a significant pest of mungbean (Vigna radiata), pigeonpea (Cajanus cajan), chickpea (Cicer arietinum), field and green beans (Vicia spp. and Phaseolus spp.), lupins (Lupinus angustifolius), lentil (Lens esculenta) and lucerne (Medicago sativa). It is also a minor pest on a range of other leguminous crops, and also some non-leguminous crops, such as cotton and citrus. There is little in the way of published information on financial losses caused by A. craccivora. However, it is an important pest in economic terms in groundnuts, cowpeas and a range of other leguminous crops. Crop losses caused by A. craccivora are due to direct feeding damage and to indirect damage, chiefly via the transmission of plant viruses.

Direct Feeding Damage

Direct damage caused by A. craccivora occurs on the seedling, vegetative and flowering stages of plants. Aphids prefer to feed on young leaves, shoots, flowers and immature seed pods. It is when aphid colonies concentrate on the growing tips of plants in the spring that the highest yield losses due to direct damage occur.

Groundnuts
A. craccivora is the most important pest of groundnuts in the tropics. Sap removal and physiological reactions of plants to aphid feeding cause direct damage. The removal of sap weakens the plant, causing poor and stunted growth, leaf curling and distorted leaf growth, wilting and reduced resistance to drought conditions, all resulting in yield losses. Damage due to irritants and toxins, produced by aphid feeding on the leaves and growing points, manifests itself in necrosis and other adverse reactions. A brown necrosis may be induced in groundnuts, for example, while disturbances in fruiting and a reduction in the root system can occur. Direct feeding damage on groundnuts by large numbers of aphids can also result in partial sterility of plants (Mayeux, 1984).

A. craccivora is probably the most injurious insect species of groundnut throughout Africa (Mayeux, 1984; Attia et al., 1986; Tarimo and Karel, 1987; Wightman and Wightman, 1994). The presence of aphids, known not to be carrying plant viral diseases, caused groundnut yields to drop by about 48%, compared with controls, in a study in Niger. The effects of aphid feeding became insignificant after the 42nd day of seedling growth, compared with controls, by which time 73% of the pods had been formed. The economic damage due to aphids varies with the stage of plant development, with most damage caused if aphids infest the growing points of groundnut early in the plant's development (Mayeux, 1984). In southern Africa, groundnut yields of 0.6-0.7 tons dry pods/ha are typical, well below what can be achieved on research farms (for example, 2.0 t/ha), mainly because of losses due to crop pests and diseases (Wightman and Wightman, 1994).

In Asia, A. craccivora appears in groundnut early in the rainy season, when it can cause considerable direct feeding damage. Yield losses of ca 16% were recorded in groundnut in India due to a complex of insect pests, the predominant one being A. craccivora (Jagtap et al., 1984).

Cowpeas and other crops
A. craccivora is the most important aphid pest of cowpeas worldwide. Direct feeding damage is due to the removal of sap from leaves, pods, seeds and other aerial plant parts. Symptoms include plant stunting and seed shrivelling. In tropical regions, direct feeding damage can result in large yield reductions.

In greenhouse studies on cowpea seedlings in Nigeria, infestation with A. craccivora at the young and mid-fill pod stage caused significantly more pod shrivelling compared with uninfested pods. Infestation at the mature pod stage alone caused relatively little pod shrivelling. However, all experimental levels of infestation caused significant reductions in seed yield, irrespective of the age of the pod (Ofuya, 1989). The use of resistant cowpea varieties, combined with the presence of natural enemies, can be an effective means of pest control in Nigeria (Ofuya, 1995; Bottenberg et al., 1998).

In field and greenhouse studies in Kenya on cowpea, the influence of the duration of infestation by A. craccivora on aphid-resistant (ICV-12) and aphid-susceptible (ICV-1) cultivars was compared with uninfested controls. Significant interactions were found between cultivar and duration of infestation for plant yields, quantified as pods per plants and seeds per pod. Reductions in seeds per pod were significantly reduced by aphid infestations of 7 days or more. The cultivar selection and aphid density at initial infestation were the major factors on cowpea growth and yield (Annan et al., 1995, 1996). In further studies, it was found that fertilizer application and cultivar selection were both important for cowpea growth and yield, as well as aphid population dynamics and bionomics. Aphid counts were higher in all fertilizer treatments compared with untreated controls. Cultivar selection was the most important factor determining yield, with aphid-resistant cultivars giving best yields. With susceptible cultivars, fertilizer application in some cases counteracted the adverse effects of A. craccivora feeding, but in other cases actually aggravated the plant growth deformities and yield reductions caused by aphids (Annan et al., 1997).

Infestations of A. craccivora on cowpeas produced deleterious effects on plant physiology, manifested in reduced growth (quantified by leaf height, leaf area growth rate and plant height) and losses in yield (Annan et al., 1995). In Canadian laboratory studies, short-term feeding was shown to significantly alter carbon dioxide exchange and photoassimilate partitioning. If infestation lasted 10 days or more, significant reductions in plant growth and respiration resulted. If infestations were terminated, plants could compensate for aphid-induced physiological changes (Hawkins et al., 1987, 1988). Direct feeding damage caused by inoculation of cowpea with A. craccivora, in a Chinese study, resulted in a reduction in plant height (to 41.9% of the controls), reduction in the green leaf area index from the 7th day after inoculation, and delayed production of harvestable pods by 30 days (Chang and Thrower, 1981).

In Pakistan, A. craccivora infestations ranged from 14 to 76% on chickpea, and from 11 to 100% on lentil, with indigenous varieties exhibiting the greatest levels of resistance (Mushtaq, 1977). In a study of the efficacy of synthetic pyrethroid pesticides on greengram (Vigna radiata) in Assam, India, against A. craccivora, the yield in infested and untreated field plots was 2.52 q/ha. This compared with yields of 6.39-11.57 q/ha in plots treated with eight pyrethroid insecticides. Even in untreated plots, however, aphid populations declined toward the end of the study, probably due to rising temperature and maturity of the crop (Borah, 1996).

A. craccivora was reported to be the most damaging pest of cowpeas in Egypt, particularly early in the growing season with aphids migrating from other food plants (Attia et al., 1986). Untreated faba bean fields in Egypt suffered slight to heavy damage due to direct feeding by A. craccivora (Bishara et al., 1984). The aphid is also a significant pest of mungbean in Egypt (Farghali et al., 1996).

In pasture grasses, A. craccivora can reach large population levels, causing direct feeding damage. Large colonies have been reported in the inflorescences of lucerne in Russia at the beginning of the flowering period, with infestation of plants sometimes reaching 60-70% (Keston, 1975). A. craccivora is also an important pest of cotton in the Zaravshan valley of Uzbekistan (Khushbaktov, 1995).

The economic injury level (EIL) of A. craccivora has been calculated as 10-16 aphids/plant on green gram (cv. AAU 34) in India (Sarma et al., 2000), 10.82 aphids/plant on black gram (cv. T-9) in India (Deka and Dutta, 2001) and 8.6 aphids/plant on faba bean in Egypt, depending on the market price of bean and control cost during the season (Abdou et al., 2012).

Factors Affecting Yield Loss

Environmental factors affect yield losses due to direct feeding damage, which is related to aphid population development. The optimal development time of A. craccivora in experimental studies occurs at around 65% relative humidity and in temperatures between 24-28.5°C (Mayeux, 1984).

In a study on field beans (Vicia faba) in Hungary during 1986 and 1987, A. craccivora was common and of economic importance, with 22.3 aphids/plant on average recorded in 1986. In this year, rapid population increase was attributed to relatively high temperatures (16.8-17.6°C) in late April and early May. In 1987, the average temperature was lower (11.1-13.3°C) during this period, while rainfall was relatively high (125 mm), and aphid reproduction did not begin until the beginning of June (Kuroli et al., 1988).

High rainfall is not favourable to A. craccivora populations since aphids do not survive periods of heavy rain. Severe tornadoes can stop swarms of aphids, while heavy rains can destroy colonies on young plants, where foliage is not plentiful enough to protect them (Mayeux, 1984). Natural enemies can also limit population build up of aphids.

The sowing date of crops influences A. craccivora direct feeding damage. Groundnuts sown early have a better chance of producing a good yield, as they go through a key period of growth before aphid populations build up to high levels. Sowing date was found to be an important factor determining the yield of field beans in the Sudan, with seed yield/plant being positively correlated with number of pods and negatively with percentage infection with Sudanese broad bean mosaic virus (SBBMV), which is transmitted by A. craccivora and Acyrthosiphon sesbaniae (Salih et al., 1973). The close spacing of groundnuts can deter A. craccivora and is also an important cultural factor influencing yield loss (Mayeux, 1984).

Indirect Damage

Honeydew
Indirect damage caused by A. craccivora is due to the production of honeydew and the transmission of plant viruses. Aphid honeydew is a digestive product with a very high sugar content. It builds up on plants when high aphid numbers occur, and acts as a substrate for fungal growth, particularly of sooty moulds. Plant respiration and photosynthetic efficiency are adversely affected (Mayeux, 1984).

Virus transmission
A. craccivora is an important vector of plant viral disease, transmitting over 30 plant viruses, including groundnut rosette, groundnut (peanut) mottle and subterranean clover stunt. It also a vector of a range of viruses of beans, cardamoms, groundnuts, peas, beet, cucurbits and Cruciferae (Blackman and Eastop, 2000). A. craccivora is polyphagous, and reservoirs of aphids and associated disease infection can exist on non-crop plants throughout the year.

Groundnut rosette virus
Groundnut rosette virus
(GRV) is a complex of at least five viruses, varying in distribution. GRV is transmitted in a persistent manner by A. craccivora, and may persist in aphids for more than 10 days (Mayeux, 1984). Groundnut rosette can cause serious morphological disturbances to groundnut plants, which take on a bushy appearance. Other symptoms include yellowing, mottling, leaf mosaic, and stunting and distortion of the shoots. It can account for extensive yield losses. If plants are infected when young, they may produce no nuts. The complex of GRV strains, along with an assistor luteovirus (GRAV) and satellite RNAs cause groundnut rosette disease. Distinct disease types have been recognised, dependent on the GRV strains involved: groundnut chlorotic rosette disease, groundnut green rosette disease and groundnut mosaic rosette disease. A. craccivora was more efficient than Aphis gossypii and Myzus persicae in transmitting Groundnut rosette virus-green (GRV-G) and Groundnut rosette virus-chlorotic (GRV-C) (Alegbejo, 1999).

The GRV complex was first described in Africa in 1907, when it strongly affected groundnut production. It has continued to do so, for example, infecting 80-90% of groundnut plants in the Belgian Congo in 1939 (Réal, 1955); and causes high pod yield losses (Bock, 1973; Olorunju et al., 1991). A. craccivora is now considered the most important insect pest of groundnuts in Africa because it transmits GRV persistently (Wightman and Wightman, 1994). Alegbejo and Abo (2002) reviewed the aetiology, ecology, transmission and control of groundnut rosette disease, which appears to be restricted to Africa.

Major yield losses result when secondary spread within the crop occurs due to aphid vectors. GRV can spread rapidly within a groundnut field. In a study in Tanganyika, low infestation of aphids was noted on 16 January, while the first plants with rosette infestation were found on 22 January. A tenfold increase in infestation occurred in the next 2 weeks accompanied by rapidly increasing aphid numbers, with 65% of plants in that field infested with GRV by late February (Evans, 1954).

In a study in Niger in 1981, yields in groundnut crops were 1350 kg/ha following low aphid infestations, compared with 183 kg/ha following high infestations of aphids carrying GRV (Mayeux, 1984). A typical groundnut yield in Africa would be around 450-670 kg/ha, but using insecticides and other management practices yields over 1300 kg/ha can exceptionally be achieved.

Groundnut varieties resistant to GRV were found in Africa in the early 1950s. Systematic plant breeding programmes have been in operation since then (Evans, 1954; Mayeux, 1984). In studies conducted in Nigeria, eight genotypes that were either resistant or susceptible to GRV were planted and infested with viruliferous A. craccivora. Infestation with rosette resulted in a seed yield 33 times higher in the resistant genotypes than in the susceptible ones, while yields were comparable in the two groups under rosette disease-free conditions (Olorunju et al., 1991). It has been reported that GRV resistant varieties from one region in Africa may succumb if grown in another region (Anon., 2000).

Sowing as early as possible in the rainy season, close crop spacing, and the use of insecticides are the recommended methods for controlling GRV in groundnuts in Africa (Farrell, 1976). In Nigeria, for example, dense crops were less heavily colonized than sparse crops (A'Brook, 1964, 1968). In a survey of groundnuts in Malawi, Zambia and Zimbabwe, economic losses due to GRV were low, which was attributed to A. craccivora, the principal vector, being controlled by natural processes, and to the practice of early and synchronous sowing (Wightman and Wightman, 1994).

Insecticides have given yield increases of over 650 kg/ha of shelled groundnuts in rainfed forests and over 1000 kg/ha under irrigation in rosette-prone crops. Insecticide spraying must be done before disease symptoms appear and before aphids can easily be found, however, to give maximum yield advantages; while some farmers may be reluctant to spray seemingly uninfested crops (Anon., 2000).

In India, the incidence of GRV was 4.9% in bunched and 3.3% in spreading varieties of groundnut, while incidence of GRV was nearly twice as high in irrigated compared with rain-fed crops. The loss in yield varied from 27 to 100%. There was a positive correlation between infestation with A. craccivora and the incidence of rosette disease. Close spacing and late weeding were highly effective in reducing aphid infestation and virus infection, and in increasing yield (Kousalya et al., 1971, 1973).

Infections of Groundnut rosette assistor virus (GRAV), in the absence of GRV and its satellite RNAs, reduced leaf area, decreased plant height, reduced the dry weight of haulms and reduced seed weight in four genotypes of groundnut, indicating that infections of GRAV alone can affect plant growth and contribute to yield losses (Naidu and Kimmins, 2007).

Peanut mottle and groundnut stunt viruses

Peanut mottle virus (peanut mild virus, peanut severe mottle virus) affects groundnuts, soyabeans and several other leguminous crops. It causes mottling with necrosis and mosaic symptoms. Leaf crinkling or other malformations can occur, while photosynthetic efficiency is reduced. A. craccivora transmits this virus in a non-persistent manner, and aphids can remain infective for 2 hours (Bock, 1973; Brunt et al., 1996).

In a survey of groundnuts in Hubei, China, it was reported that infestation by groundnut (peanut) mild mottle virus, transmitted by A. craccivora, was between 1 and 98%. In experimental trials, the mild mottle virus reduced groundnut yields by up to 23% (Xu et al., 1983). Infection with Peanut mottle virus led to a mean yield loss of 20%/plant in Georgia, USA, in 1973 (Kuhn and Demski, 1975). Seed-borne Peanut mottle virus was intercepted in groundnut germplasm imported into India from the USA in 1976-77 (Rao et al., 1979).

A. craccivora was found to be an important vector of Peanut stunt virus in lucerne in Sudan. Reservoirs of aphids and virus infection could spread to other leguminous crops, and it has been recommended that lucerne cultivation should not be introduced into the main groundnut-growing areas of the Sudan (Ahmed and El Sadig, 1985).

A. craccivora was reported transmitting five isolates of Peanut stripe virus (PStV) on groundnut in China (Chen et al., 1999).

Other viruses

Subterranean clover stunt virus (SCSV) infects all varieties of Trifolium subterranean, and many species of Medicago and Trifolium. It is spread in a persistent manner by A. craccivora and several other aphids (Brunt et al., 1996). In Australia and New Zealand, A. craccivora is an important pest of pasture grasses, largely because it is a vector of SCSV, which causes mild or severe stunting, marginal chlorosis and puckering of leaflets, and stunting and yellowing of new growth (Gutierrez et al., 1974a, b; Ashby et al., 1982). A. craccivora is also a vector of SCSV in broad bean in Tasmania, where extensive clover plantings potentially act as a reservoir of aphids and virus infection (Johnstone, 1978).

Bean common mosaic virus (BCMV) (bean mosaic virus, bean western mosaic virus, mungbean mosaic virus) infects Phaseolus spp. and a range of other bean and pea crops worldwide. The related Bean yellow mosaic virus (BYMV) also infects Phaseolus and peas, causing necrosis and mottling, although it also causes mottling in groundnuts. A. craccivora transmits BCMV and BYMV in a non-persistent manner (Brunt et al., 1996). For example, A. craccivora transmits both BCMV and BYMV in green beans (Phaseolus vulgaris) in New Zealand (Malone and Hartley, 1978). BCMV was identified occurring on cowpea (Yadav, 2010) and French bean (Phaseolus vulgaris) (Verma and Gupta, 2010) in India, and was transmitted through sap, by seed and by aphids, including A. craccivora.

In the Philippines, mungbeans were most susceptible to mungbean mosaic virus at an early growth stage. A. craccivora was the only insect vector, transmitting the virus in a non-persistent manner even at low aphid densities. The earlier the plant became infected, the higher the percentage of infected seeds (Khan and Lapis, 1989). Mungbean infected in Iran by a virus described as a mungbean strain of Bean common mosaic virus (M-BCMV), which caused deformation, puckering, rolling, blistering and mosaic symptoms, reduced yields by 31-75%, particularly when infection occurred before pod set (Kaiser and Mossahebi, 1974).

A. craccivora was recorded transmitting Soybean mosaic virus (SbMV) in Illinois, USA (Halbert et al., 1981, 1986).

A. craccivora is a significant vector of several viruses of lupins, including BYMV and Cucumber mosaic virus (CMV). In Australia, aphids decrease the grain yield of lupins directly, when large numbers of aphids colonize plants late in the growing season, but mainly cause economic damage via the spread of CMV infection. Incoming viruliferous alatae initiate the infection and yield loss is greatest when aphids arrive early in the crop's development. Intensive insecticide treatments against A. craccivora (and also Myzus persicae), in Australian field experiments, decreased CMV infection, and increased grain yields by up to 35% (Bwye et al., 1997).

In Iran, A. craccivora is an important vector of a number of viruses of leguminous crops, including BYMV, CMV, Alfalfa mosaic virus (AMV), and Pea-leaf roll virus (PLRV) in lentil. In field inoculation studies in lentil with these viruses, yield was reduced by 46-94%. Transmission of CMV and PLRV by A. craccivora in cowpeas and broad beans can also cause yield losses of this order (Kaiser, 1973a, b).

Six aphid species, including A. craccivora, transmitted Bean leafroll virus (BLRV) in a circulative persistent manner on Vicia fabae in Egypt (El-Beshehy and Azza, 2013), and A. craccivora transmitted Cowpea aphid-borne mosaic virus (CABMV) in a non-persistent manner in cowpea in Saudi Arabia (Damiri et al., 2013).

A new virus involved with leaf crinkle disease of urdbean (Vigna mungo) was observed in Himachal Pradesh, India. The virus was transmitted by A. craccivora and via seed and sap (Sharma et al., 2014).

An unknown virus isolated from Senna hirsuta in Nigeria and tentatively designated as Senna mosaic virus (SeMV) was transmitted by A. craccivora in a non-persistent manner (Owolabi and Proll, 2001).

An unknown or uncommon member of the family Luteoviridae causing yellowing and stunting symptoms on chickpea and faba bean was observed in Ethiopia. The name Chickpea chlorotic stunt virus was proposed for this disease. A. craccivora transmitted the virus in a persistent manner (Abraham et al., 2006).

In field studies in Turkey, 10% of aphids of A. craccivora were viruliferous with Plum pox virus-T (Caglayan et al., 2013).

A. craccivora has also been recorded as a vector of Onion yellow dwarf virus (OYDV) (Kumar et al., 2009), Papaya ringspot virus (PRSV) (Kalleshwaraswamy and Kumar, 2008; Kumar et al., 2010; Isha Bhoyer et al., 2014), Pepper veinal mottle virus (PVMV) (Fajinmi et al., 2011), Watermelon mosaic virus (WMV) and Zucchini yellow mosaic virus (ZYMV) (Haque et al., 2004), Sunflower mosaic virus (SMV) (Singh et al., 2005), Faba bean necrotic yellows virus (FBNYV) (Vega-Arreguín et al., 2007) and Faba bean necrotic stunt virus (FBNSV) (Grigoras et al., 2009).

Threatened Species

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Threatened SpeciesConservation StatusWhere ThreatenedMechanismReferencesNotes
Sesbania tomentosaNational list(s) National list(s); USA ESA listing as endangered species USA ESA listing as endangered speciesHawaiiPest and disease transmissionUS Fish and Wildlife Service, 2010

Risk and Impact Factors

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  • Pest and disease transmission

Detection and Inspection

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On groundnut, very young rolled up leaves of seedlings should be examined for nymphs early in the season.

Similarities to Other Species/Conditions

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Some other members of the craccivora group are morphologically very similar, particularly A. loti (regarded as a synonym by some authors). Heie (1986) summarizes the morphological differences between A. craccivora and A. loti. Differences are most evident between the oviparous forms, most noticeably fewer scent plaques (between 40 and 100) in A. loti compared to 110-230 in A. craccivora. Among the males, A. craccivora are winged and A. loti are wingless.

Prevention and Control

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Chemical Control

Most major groups of insecticides have been used against this insect pest, including chlorinated hydrocarbons, organophosphates, carbamates and pyrethroids. The persistence and effectiveness of insecticides on the plants is an important factor. Control in groundnuts must be very effective between germination and the 40th day, and therefore systemics with satisfactory persistence through this growth stage are preferred. The high cost of systemics to farmers in the developing world emphasizes the need for early warning and forecast systems (Mayeux, 1984). Systemics will kill aphids effectively, but they may still have time to feed and transmit virus before dying. In such circumstances, it may be more effective to control aphids on wild hosts on which they feed before dispersing to crops.

The use of monitored applications of insecticides (which are carried out only when insect infestation/damage reaches or exceeds an action threshold) can provide effective control of pests while saving costs and reducing environmental pollution by a decrease in the number of applications made. A study in cowpea in Nigeria found that there was no significant difference in yields between calendar and monitored spray treatments. Fewer chemical applications were made in monitored spray treatments and in calendar spray treatments made four times every 10 days than in calendar spray treatments made five times every 7 days (Egho, 2010).

Dhingra (1993) described pest resistance to pyrethroid insecticides. Other sprays tried on crops include neem (Dimetry and El Hawary, 1995) and petroleum oil (El Sisi and El Hariry, 1991). Formulations of neem (Azadirachta indica) have been shown to be effective against A. craccivora and can be used as an alternative to chemical insecticides (Egho et al., 2009; Baidoo et al., 2012; Chaudhari et al., 2015).

Experimental work is focused on botanical insecticides, e.g. oil of Parkia roxburghii (Salam et al., 1995), fractions of Atriplex semibaccata (Barakat et al., 2005) and crude extracts of Halocnemon strobilacium (Abdallah et al., 2009). A mixture of alkaloids of Sophora alopecuroides and nicotine showed significant synergistic activity against A. craccivora in field trials (Huo et al., 2014).

Cultural Control

Mayeux (1984) reviewed crop protection methods in groundnut, including the use of early and dense sowings. Early sowings allow plants to start flowering before aphids appear, while dense sowings provide a barrier to aphids penetrating in from field edges. Mulching and other cultural practices used in IPM in groundnuts in the USA were described by Wightman and Ranga Rao (1994). Sanitary measures are important within crops and between seasons to prevent the spread of viruses for which A. craccivora is a vector. Virus-infected plant material should be removed after harvest and any volunteer plants or weeds that harbour viruses should also be destroyed.

Insecticide applications interacted with cropping systems in minimizing the incidence of A. craccivora in India, when chickpeas were intercropped with barley or linseed (Prasad et al., 1988). However, cowpea and groundnut are usually not good companions, due to risk of A. craccivora spreading from cowpea to groundnut.

Field studies in Uganda evaluated the use of intercropping with green gram (Vigna radiata) or sorghum as a pest control strategy in cowpea. The response of pests varied. Populations of A. craccivora were significantly reduced in the cowpea+sorghum intercrop but were higher in the cowpea+green gram intercrop. It is concluded that to be effective, intercropping needs to be part of a pest management system that involves other control strategies and considers the pest profile (Nampala et al., 2002). In studies in Egypt, intercropping faba bean with coriander (Coriandrum sativum) significantly decreased the population of A. craccivora, significantly increased the numbers of associated predators and increased seed yields (Rizk, 2011).

Field studies on mung bean in India, found that two strains of Bacillus subtilis induced resistance to A. craccivora by enhancing the phenol and peroxidase concentrations in the host plant (Swarnali Bhattacharya et al., 2008).

Biological Control

A population explosion of A. craccivora on cowpeas was attributed to cypermethrin adversely affecting coccinellid and syrphid predators (Ofuya, 1987). Control with insecticides having less impact on natural enemies, which have lower toxicity against coccinellids, may be necessary in integrated pest management programmes. Coccinellids have often been cited as an important natural control factor in groundnuts in India and Africa (e.g. Agarwala and Bardhanroy, 1999). In Bangladesh, five larvae of the coccinellid Cheilomenes sexmaculata caused 73-95% suppression of infesting A. craccivora at high densities (490-640) and 86-100% reduction on caged bean plants in 7 days; while the efficacy of 15 larvae of C. sexmaculata per bean plant was significantly greater than two insecticide treatments (Bari and Sardar, 1998).

A potential parasitoid for biological control programmes is the braconid Trioxys indicus (Singh and Agarwala, 1992). It was introduced to Australia from India to control A. craccivora on lupins and other crops (Sandow, 1986). This parasitoid has a high fecundity and can withstand long periods of hot weather. Third-instar nymphs are most suitable for parasitism. However, the presence of hyperparasitoids such as Aphidencryptus spp. may limit its effectiveness in certain areas.

Experimental releases of an introduced aphid parasitoid, Lysiphlebus testaceipes, from the USA, were carried out in Shaanxi, China, in 1983 (Zheng and Tang, 1989). In the former USSR, Lysiphlebus fabarum, which appears in mid-April, reproduced at a rate paralleling that of the aphid and reached its peak of activity in June, when it parasitized up to 85% of an aphid population. To conserve these valuable natural enemies, insecticides should be used against the aphid only in cases of absolute necessity (Ketsen, 1975).

Releases of Coccinella septempunctata successfully reduced populations of A. craccivora and Aphis gossypii on protected crops of sweet pepper in Portugal (Valério et al., 2007).

In field studies in groundnut in India, populations of A. craccivora were reduced by releases of the reduviid predators Rhynocoris marginatus (Sahayaraj and Martin, 2003), Rhynocoris kumraii (Sahayaraj and Ravi, 2007) and the chrysopid predator Chrysoperla zastrowi sillemi (Baskaran and Rajavel, 2013), and by sprays with the fungus Verticillium lecanii (Sahayaraj and Namachivayam, 2011).

Field and laboratory studies in eastern Uganda confirmed the effectiveness of Syrphus ribesti as a potential biological control agent of Aphis craccivora on cowpea (Munyuli et al., 2006).

In experiments on soyabean in Egypt, releases of Chrysoperla carnea larvae on plants infected with healthy or Soybean mosaic virus (SbMV)-viruliferous aphids of A. craccivora resulted in an increase in the number and weight of seeds. It is suggested that C. carnea should be released in the early stage of the plants, at the first appearance of aphids, to prevent the spread of SbMV (El-Arnaouty et al., 2008).

Host-Plant Resistance

Large differences in susceptibility of chickpea genotypes to stunt disease and groundnut varieties to rosette virus complex have meant that the development of resistant plant material to these diseases has been a main aim of plant breeding programmes. Evans (1954) identified resistance in groundnuts to Groundnut rosette virus, while several resistant varieties are now available; resistance is manifest by longer aphid generation times and a much reduced fecundity. A strong negative relationship was found between the concentration of the tannin, procyanidin, in the leaf bud petioles of seven genotypes of groundnut and fecundity of A. craccivora (Grayer et al., 1992).

Resistance to A. craccivora was identified in the groundnut breeding line ICG 12991, with a lower rate of nymphal development, lower fecundity and smaller aphids than on control varieties (Minja et al., 1999). Resistance to A. craccivora in this genotype is controlled by a single recessive gene (Herselman et al., 2004). Germplasms ICG 12988 and ICG 12991 have shown resistance to groundnut rosette disease (Subrahmanyam et al., 2000). ICGV-SM 90704 is resistant to GRV but susceptible to A. craccivora (Merwe et al., 2001).

Integrated Pest Management

A. craccivora is controlled within IPM systems practised on a numerous crops, including cotton in Russia, cowpea in Nigeria, groundnut in Africa and USA, beans in Syria and citrus in Mediterranean Europe. Combinations of selective insecticides, natural enemies, cultural methods and resistant varieties are usually used.

In groundnuts, monitoring pest populations to ensure effective insecticide spray application is combined with the use of cultural methods and resistant cultivars (Mayeux, 1984). Lecoq (1983) described IPM in muskmelon, where cultural practices and partial host-plant resistance provided adequate protection. Mulching was found to be effective in delaying virus spread during the critical early growth and fruit-setting periods. Control on beans in Bangladesh was effective using malathion sprays along with natural predation by Menochiles sexmaculatus [C. sexmaculata] (Ahamad and Sardar, 1994). Shomirsaidov (1983) described IPM on cotton in Tajikistan using selective insecticides and effective natural enemies. Narzikulov (1982) calculated that IPM on cotton worked out 2-2.5 times cheaper than chemical control.

Insecticide applications interacted with cropping systems in minimizing the incidence of A. craccivora in India, when chickpeas were intercropped with barley or linseed (Prasad et al., 1988).

In a review on options for the management of cowpea pests, Adipala et al. (2000) concluded that selected combinations of agronomic, chemical and cultural control measures (IPM), particularly when combined with early planting, provide better control of cowpea pests than a single control strategy. Nabirye et al. (2003) described the results from ten IPM field schools with 10-20 farmers that were run in Uganda to determine the best practice for the management of cowpea pests, including A. craccivora. A strategy that combined early planting, close planting (30 × 20 cm) and three insecticide applications, once each at the budding, flowering and podding stages, resulted in the highest yields of 791 kg/ha with a 51% yield gain over the farmers' traditional practices.

The effects of certain agricultural practices on the infestation of six broad bean cultivars by A. craccivora were studied in Egypt. A. craccivora populations were lower at higher plant spacings (30 cm compared with 10 and 20 cm), low nitrogen fertilization levels (50 and 70 kg N/feddan) and early first irrigation (14, 21 and 28 days compared with 56 and 63 days) (Salman et al., 2007).

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26/10/15 Impact and Prevention and Control sections updated by:

Angela Whittaker, Consultant, UK.

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