Invasive Species Compendium

Detailed coverage of invasive species threatening livelihoods and the environment worldwide

Datasheet

Diuraphis noxia
(Russian wheat aphid)

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Datasheet

Diuraphis noxia (Russian wheat aphid)

Summary

  • Last modified
  • 15 November 2018
  • Datasheet Type(s)
  • Invasive Species
  • Pest
  • Vector of Plant Pest
  • Preferred Scientific Name
  • Diuraphis noxia
  • Preferred Common Name
  • Russian wheat aphid
  • Taxonomic Tree
  • Domain: Eukaryota
  •   Kingdom: Metazoa
  •     Phylum: Arthropoda
  •       Subphylum: Uniramia
  •         Class: Insecta

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Pictures

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PictureTitleCaptionCopyright
A yellow-green to grey-green spindle-shaped aphid, 1.4-2.6 mm long as an adult, integument covered with a waxy white exudate.
TitleAdult
CaptionA yellow-green to grey-green spindle-shaped aphid, 1.4-2.6 mm long as an adult, integument covered with a waxy white exudate.
CopyrightUSDA
A yellow-green to grey-green spindle-shaped aphid, 1.4-2.6 mm long as an adult, integument covered with a waxy white exudate.
AdultA yellow-green to grey-green spindle-shaped aphid, 1.4-2.6 mm long as an adult, integument covered with a waxy white exudate.USDA
D. noxia causes characteristic white to purple streaking and leaf-rolling on wheat and barley leaves; young plants are often stunted and even killed.
TitleSymptoms on wheat
CaptionD. noxia causes characteristic white to purple streaking and leaf-rolling on wheat and barley leaves; young plants are often stunted and even killed.
CopyrightUSDA
D. noxia causes characteristic white to purple streaking and leaf-rolling on wheat and barley leaves; young plants are often stunted and even killed.
Symptoms on wheatD. noxia causes characteristic white to purple streaking and leaf-rolling on wheat and barley leaves; young plants are often stunted and even killed.USDA
D. noxia parasitized by Aphidiidae.
TitleParasitized by Aphidiidae
CaptionD. noxia parasitized by Aphidiidae.
CopyrightUSDA
D. noxia parasitized by Aphidiidae.
Parasitized by AphidiidaeD. noxia parasitized by Aphidiidae.USDA
Aphelinus asychis stinging D. noxia.|Aphelinus asychis stinging Diuraphis noxia.
TitleAphelinus asychis|Attacked by A. asychis|Attacking D. noxia.
CaptionAphelinus asychis stinging D. noxia.|Aphelinus asychis stinging Diuraphis noxia.
CopyrightAngela M.I. de Farias
Aphelinus asychis stinging D. noxia.|Aphelinus asychis stinging Diuraphis noxia.
Aphelinus asychis|Attacked by A. asychis|Attacking D. noxia.Aphelinus asychis stinging D. noxia.|Aphelinus asychis stinging Diuraphis noxia.Angela M.I. de Farias
L. ninae larvae, preying on Diuraphis noxia.|Leucopis ninae larvae preying on D. noxia.
TitleL. ninae on D. noxia|Parasitized by L. ninae
CaptionL. ninae larvae, preying on Diuraphis noxia.|Leucopis ninae larvae preying on D. noxia.
CopyrightUSDA
L. ninae larvae, preying on Diuraphis noxia.|Leucopis ninae larvae preying on D. noxia.
L. ninae on D. noxia|Parasitized by L. ninaeL. ninae larvae, preying on Diuraphis noxia.|Leucopis ninae larvae preying on D. noxia.USDA

Identity

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Preferred Scientific Name

  • Diuraphis noxia (Kurdjumov, 1913)

Preferred Common Name

  • Russian wheat aphid

Other Scientific Names

  • Brachycolus noxia Mordvilko
  • Brachycolus noxius Mordvilko
  • Brachysiphoniella noxius (Mordvilko ex Kurdjumov)
  • Cavahyalopterus graminearium Mimeur
  • Cavahyalopterus noxius (Mordvilko)
  • Cuernavaca noxia (Mordvilko)
  • Cuernavaca noxius
  • Diuraphis muehlei (Borner)
  • Diuraphis noxius (Mordvilko ex Kurdjumov)
  • Holcaphis noxius (Mordvilko ex Kurdjumov)
  • Quernavaea noxia

International Common Names

  • English: barley aphid; Russian grain aphid

Local Common Names

  • Germany: Russische Getreide-Blattlaus; Russische Weizen-Blattlaus

EPPO code

  • BRAYNO (Diuraphis noxia)

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: Diuraphis
  •                                         Species: Diuraphis noxia

Notes on Taxonomy and Nomenclature

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D. noxia was first recognized as a separate species by Mordvilko (Kurdjumov, 1913), which he named Brachycolus noxius. Since then it has been described under various names by Aizenberg (1935), Mimeur (1942), Bodenheimer and Swirsky (1957) and Anon. (1963). Eastop and Hille Ris Lambers (1976), Blackman and Eastop (1984) and Stoetzel (1987) all place it in the genus Diuraphis Aizenburg.

Description

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D. noxia is a yellow-green to grey-green spindle-shaped aphid, 1.4-2.6 mm long as an adult. Although its integument is slick just after moult, it soon becomes covered with a waxy white exudate. The eyes and distal third of the antennae are dark; the head and thorax of winged adults are also dark. It has six-segmented antennae about half the length of the body. The rostrum reaches to about the middle coxae. The cornicles are pale and very truncated (50-60 µm long), being about as wide as long, so that they are inconspicuous. The cauda is elongate, and above the cauda is a supracaudal process on the 8th abdominal tergite, which gives it the appearance of having two caudae and distinguishes it from all other cereal aphid species. The supracaudal process is almost as long as the cauda on apterous adults, but it is smaller and less conspicuous on alates and nymphs.

Blackman and Eastop (2000), Stoetzel (1987), Pike et al. (1990) and Olsen et al. (1993) give descriptions and keys to D. noxia and other cereal aphids. Aalbersberg et al. (1987) and Olsen et al. (1993) provide keys to the immature stages.

Distribution

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Although found in Xinjiang-Uigur Autonomous Region of the Peoples Republic of China for about 70 years, D. noxia has not spread to the major wheat-growing area of central China, but there is concern that it may do so in the future (Zhang, 1991). D. noxia has recently been found in Chile and Argentina but has not yet become a pest (Reed and Kindler, 1994). D. noxia has been found in Kenya where it is considered a pest, and biological control is being investigated.

D. noxia spread to 17 western states in the USA and three western provinces in Canada within 3 years of its introduction, it has not spread eastward in the USA or Canada (Webster and Amosson, 1994). D. noxia has not spread much into central or northern Europe.

D. noxia has been detected in South Australia for the first time where it has been found in cereal crops in the Mid-North (Government of South Australia, 2016). There is also a preliminary report of the pest in Victoria (IPPC, 2016).

A record of D. noxia in the UK (EPPO, 2006) published in previous versions of the Compendium is now considered erroneous and has been removed.

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

AfghanistanPresentCABI, 1991; EPPO, 2014
China
-XinjiangWidespreadZhang, 1991
IranPresentCABI, 1991; EPPO, 2014
IraqPresentAli et al., 1985; EPPO, 2014
IsraelPresentCABI, 1991; Hopper et al., 1997; EPPO, 2014
JordanPresentTanigoshi et al., 1995; EPPO, 2014
KazakhstanPresentGruber et al., 1991; Hopper et al., 1997; EPPO, 2014
KyrgyzstanWidespreadGruber et al., 1991; Hopper et al., 1997
PakistanPresentEPPO, 2014
Saudi ArabiaPresentAlhag et al., 1996
SyriaPresentEPPO, 2014
TurkeyPresentCABI, 1991; Gruber et al., 1991; EPPO, 2014
UzbekistanWidespreadHopper et al., 1997
YemenPresentStary and Erdelen, 1982; EPPO, 2014

Africa

AlgeriaPresentCABI, 1991; Miller et al., 1993; EPPO, 2014
EgyptPresentCABI, 1991; EPPO, 2014
EthiopiaPresentHaile and Megenasa, 1987; EPPO, 2014
KenyaPresentIntroduced1995 Invasive Macharia et al., 1999; IPPC-Secretariat, 2005
LibyaPresentCABI, 1991; EPPO, 2014
MoroccoPresentTanigoshi et al., 1995; Hopper et al., 1997; EPPO, 2014
South AfricaPresentIntroduced1978Walters et al., 1980; EPPO, 2014
ZimbabwePresentEPPO, 2014

North America

CanadaRestricted distributionEPPO, 2014
-AlbertaPresentJones et al., 1989; Butts, 1992; EPPO, 2014
-British ColumbiaPresentJones et al., 1989; EPPO, 2014
-SaskatchewanPresentJones et al., 1989; Butts, 1992; EPPO, 2014
MexicoPresentGilchrist et al., 1984; EPPO, 2014
USARestricted distributionEPPO, 2014
-ArizonaWidespreadWebster and Amosson, 1994
-CaliforniaWidespreadWebster and Amosson, 1994
-ColoradoPresentWebster and Amosson, 1994; EPPO, 2014
-IdahoWidespreadWebster and Amosson, 1994
-KansasPresentWebster and Amosson, 1994; EPPO, 2014
-MontanaPresentWebster and Amosson, 1994; EPPO, 2014
-NebraskaPresentWebster and Amosson, 1994; EPPO, 2014
-NevadaWidespreadWebster and Amosson, 1994
-New MexicoPresentWebster and Amosson, 1994; EPPO, 2014
-North DakotaRestricted distributionWebster and Amosson, 1994
-OklahomaPresentWebster and Amosson, 1994; EPPO, 2014
-OregonWidespreadWebster and Amosson, 1994
-South DakotaRestricted distributionWebster and Amosson, 1994
-TexasPresentIntroduced1986Webster and Amosson, 1994; EPPO, 2014
-UtahWidespreadWebster and Amosson, 1994
-WashingtonPresentWebster and Amosson, 1994; EPPO, 2014
-WisconsinPresentNAPIS, 2000
-WyomingPresentWebster and Amosson, 1994; EPPO, 2014

South America

ArgentinaPresentIntroduced1991Reed and Kindler, 1994; EPPO, 2014
ChilePresentIntroduced1987Reed and Kindler, 1994; EPPO, 2014

Europe

AlbaniaWidespreadEPPO, 2014
BulgariaPresentGruber et al., 1991
CroatiaPresentCuljak and Barcic, 2002
Czech RepublicPresentStary, 1996
FrancePresentGruber et al., 1991; Hopper et al., 1997; EPPO, 2014
GermanyPresentThieme et al., 2001
GreeceWidespreadHopper et al., 1997; EPPO, 2014
HungaryPresentBasky, 1993; EPPO, 2014
ItalyPresentHopper et al., 1997; EPPO, 2014
MacedoniaWidespreadHopper et al., 1997
MoldovaWidespreadGruber et al., 1991
PortugalPresentEPPO, 2014
-MadeiraPresentEPPO, 2014
Russian Federation
-Central RussiaWidespreadGrossheim, 1914
-Russia (Europe)WidespreadGrossheim, 1914; Kovalev et al., 1991
SlovakiaPresentBarta and Cagan, 2005; Barta and Cagan, 2007; Barta and Cagan, 2007
SpainPresentEPPO, 2014
UKAbsent, invalid recordEPPO, 2010; EPPO, 2014
UkrainePresentGruber et al., 1991; EPPO, 2014
Yugoslavia (Serbia and Montenegro)PresentHopper et al., 1997

Oceania

Australia
-South AustraliaPresentIntroduced Invasive Government of South Australia, 2016; IPPC, 2016In the Mid-North, south of Tarlee
-VictoriaRestricted distributionIPPC, 2016Preliminary report.

Risk of Introduction

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D. noxia was found in Chile in 1987 and in Argentina in 1991, and it is spreading rapidly (Reed and Kindler, 1994). Although it has not yet become a pest in Argentina and Chile, its rapid spread suggests that it may soon become one. D. noxia has not yet been found in several other major wheat-growing areas, such as Australia and central China. However, Hughes and Maywald (1990), using a climate-matching model, predicted potentially severe infestation of Australian cereals if D. noxia were introduced.

Although nymphs and adults are very unlikely to be transported alive on harvested grains or fodder, overwintering eggs could be transported in this way. However, D. noxia is anholocyclic in North America so far, despite the discovery of a few oviparae (Kiriac et al., 1990). Although it is possible that the sexual phase was lost after colonization, it appears more likely that the founders were also anholocyclic.

Hosts/Species Affected

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Kindler and Springer (1989) found that D. noxia could develop on 47 out of 48 cool-season grass species and 18 out of 32 warm-season grass species to which it was exposed in the greenhouse. They found that it did not develop on any of 27 legume species or 17 forb species to which it was exposed. Pike and Allison (1991) compiled a list of host plants from the literature, comprising 140 grass species with varying degrees of suitability for D. noxia reproduction. Zea mays does not appear suitable for D. noxia reproduction, and Sorghum bicolor is only marginally suitable (Webster et al., 1987). Thus it appears that D. noxia is restricted to grasses. Furthermore, it reproduces best and does most damage on cool-season grasses.

Host Plants and Other Plants Affected

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Growth Stages

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

Symptoms

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Although D. noxia feeds on leaves and flowers/seedheads of grasses, it appears to inject a polypeptide toxin that affects the entire plant (Hewitt et al., 1984). Colonies are found most frequently on the youngest leaves or on newly emerged flowers/seedheads. Even very small colonies of D. noxia cause characteristic white to purple streaking and leaf-rolling on wheat and barley leaves. The streaks usually extend most of the long axis of the leaf and are irregularly distributed across the short axis of the leaf. Rolling extends in severity from simple folding of the leaf along the mid-vein, to one side of the leaf rolled in upon itself, to the whole leaf being tightly rolled around the aphid colony. Large colonies can roll the flag leaf to the point where the tip of the inflorescence becomes trapped, giving it a fish-hook shape. Young plants are often stunted and even killed. Plants attacked after flowering show few to no obvious symptoms. Duration of infestation may have more impact than aphid density on yield loss (Burd and Burton, 1992; Kieckhefer and Gellner, 1992).

List of Symptoms/Signs

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SignLife StagesType
Inflorescence / discoloration panicle
Leaves / abnormal colours
Leaves / leaves rolled or folded
Leaves / necrotic areas

Biology and Ecology

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Hughes (1988, 1996) and Araya et al. (1990) review the literature on D. noxia, and Poprawski et al. (1992) give a complete bibliography from 1886 to 1992.

D. noxia is multivoltine with a development time from birth to reproduction of 11 days at 20°C. With a daily fecundity of 1-3 nymphs per day and adult longevity of about 80 days in the laboratory, D. noxia has a high potential capacity for increase with rm = 0.06/day for a temperature regime of 5-15°C (Michels and Behle, 1989). Aalbersberg et al. (1987) and Kieckhefer and Elliott (1989) found somewhat similar values for development rate, survival, and reproduction, although differences in techniques make comparisons problematic.

Plant growth stage, temperature, and their interaction affect D. noxia life span and reproduction (Girma et al., 1990). Reproduction is greatest at 18-21°C on wheat in growth stages from jointing to heading. D. noxia develops faster, lives longer, produces more progeny, and thus has a higher rate of increase on wheat than on rye (Behle and Michels, 1990). D. noxia from wheat have higher levels of reproduction when shifted to less suitable host plants, than those grown for several generations on the less suitable plants (Schotzko and Smith, 1991; Worrall and Scott, 1991; Robinson, 1993).

In the northern part of the USA and in Canada, overwintering mortality can be 100% if temperatures are low enough for long enough (Butts, 1992; Armstrong et al., 1992). For D. noxia, supercooling point is weakly if at all related to ability to survive cold winters; most winter mortality occurs at temperatures well above the supercooling point (Butts, 1992). Winter survival can depend on the details of aspect and snow cover so that within a field, survival can be high on the south-facing sides of furrows receiving greater solar insolation (Hammon and Peairs, 1992).

The geographical range of D. noxia (particularly the areas where it is a pest) is restricted to regions of low rainfall. Furthermore, even in areas with low rainfall, D. noxia is rarely a problem in irrigated cereals, and populations decline after heavy rainfall. These observations suggest that precipitation and/or humidity may directly or indirectly reduce survival or reproduction of D. noxia.

Alates are formed when plants are water stressed, not when aphids are crowded (Baugh and Phillips, 1991). Messina (1993) found only 15-35% of variation in alate production was explained by crowding, and prenatal experience has a strong effect on alate formation. D. noxia density has little effect on per capita growth rate (Messina, 1993). The rapid spread of D. noxia across the western USA and its ability rapidly to infest newly germinated wheat suggest a high dispersal rate. However, quantitative measurements are lacking, and it seems likely that it was present in many areas at low numbers before being detected, so that its rate of spread may have been less than it would appear.

Wheat and barley can provide suitable habitat for 9-11 months of the year, but wild grasses are very important for persistence and growth of D. noxia populations (Kriel et al., 1986; Aalbersberg et al., 1988a; Armstrong et al., 1991; Montandon et al., 1993). D. noxia can oversummer on a variety of wild grass species (Kindler and Springer, 1989; Clement et al., 1990; Messina et al., 1993b), which thus provide a bridge for infestation of autumn-planted cereals.

In Eurasia, some populations have both sexual and asexual generations (i.e. are holocyclic) with sexual forms and egg laying in the fall so that the aphids overwinter as eggs (Grossheim, 1914; Basky, 1993). However, in other European populations (e.g. in southern France) and in all populations in the USA only asexual reproduction has been found (i.e. populations are anholocyclic), although a few ovipariae have been observed in USA populations (Kiriac et al., 1990). Besides differences in life-cycle among populations, D. noxia populations vary in the damage done to host plants (Bush et al., 1989; Puterka et al., 1992, 1993), in fecundity (Webster et al., 1993a), and in randomly amplified polymorphic DNA markers (Black et al., 1992; Puterka et al., 1993).

Despite initial indications that D. noxia could vector diseases such as Barley yellow dwarf virus (Rybicki and von Wechmar, 1982; von Wechmar and Rybicki, 1984), subsequent experiments showed little if any persistent transmission of viruses (Kriel et al., 1986; Fouche et al., 1984; Hewitt et al., 1984; Halbert et al., 1992; Damsteegt et al., 1992).

Natural enemies

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Natural enemyTypeLife stagesSpecificityReferencesBiological control inBiological control on
Adalia bipunctata Predator Adults/Larvae/Nymphs South Africa wheat
Aeolothrips intermedius Predator Adults/Nymphs USSR wheat
Aphelinus Parasite Adults/Nymphs
Aphelinus albipodus Parasite Adults/Eggs/Larvae/Nymphs/Pupae Washington
Aphelinus asychis Parasite Adults/Nymphs Texas
Aphelinus toxopteraphidis Parasite Adults/Nymphs USSR wheat
Aphelinus varipes Parasite Adults/Nymphs Australia
aphid lethal paralysis virus Pathogen Adults/Nymphs
Aphidius Parasite Adults/Nymphs
Aphidius colemani Parasite Adults/Nymphs Washington
Aphidius ervi Parasite Adults/Nymphs USSR wheat
Aphidius hortensis Parasite Adults/Nymphs
Aphidius matricariae Parasite Adults/Nymphs Washington
Aphidius nigripes Parasite Adults/Nymphs
Aphidius picipes Parasite Adults/Nymphs USSR wheat
Aphidius rhopalosiphi Parasite Adults/Nymphs Washington
Aphidius setiger Parasite Adults/Nymphs
Aphidius urticae Parasite Adults/Nymphs USSR wheat
Aphidius uzbekistanicus Parasite Adults/Nymphs USSR wheat
Aphidoletes aphidimyza Predator Adults/Nymphs USSR wheat
Beauveria bassiana Pathogen Adults/Nymphs
Cantharis lateralis Predator USSR wheat
Cheilomenes intermedia Predator Adults/Eggs/Larvae/Nymphs/Pupae
Cheilomenes literate Predator Adults/Eggs/Larvae/Nymphs/Pupae
Cheilomenes lunata Predator Adults/Nymphs
Cheilomenes vincia Predator Adults/Eggs/Larvae/Nymphs/Pupae
Chrysopa abbreviata Predator Adults/Nymphs USSR wheat
Chrysopa formosa Predator Adults/Nymphs USSR wheat
Chrysopa hummeli Predator Adults/Nymphs USSR wheat
Chrysopa phyllochroma Predator Adults/Nymphs USSR wheat
Chrysoperla Predator Adults/Nymphs
Chrysoperla carnea Predator Adults/Nymphs USSR wheat
Chrysoperla rufilabris Predator Adults/Nymphs
Coccinella Predator Adults/Eggs/Larvae/Nymphs/Pupae
Coccinella novemnotata Predator
Coccinella septempunctata Predator Adults/Nymphs North America; USSR wheat
Coccinella transversoguttata biinterupta Predator Adults/Eggs/Larvae/Nymphs/Pupae
Coccinella undecimpunctata Predator Adults/Nymphs
Coccinellina ancoralis Predator Adults/Nymphs
Coccinula quatuordecimpustulata Predator Adults/Nymphs USSR wheat
Conidiobolus coronatus Pathogen Adults/Nymphs
Conidiobolus obscurus Pathogen Adults/Nymphs
Conidiobolus thromboides Pathogen Adults/Nymphs
Cycloneda ancoralis Predator Adults/Nymphs
Diaeretiella rapae Parasite Adults/Nymphs USSR; Washington wheat
Diaretus obsoletus Parasite Adults/Eggs/Larvae/Nymphs/Pupae
Ephedrus Parasite Adults/Eggs/Larvae/Nymphs/Pupae
Ephedrus persicae Parasite Adults/Nymphs
Ephedrus plagiator Parasite Adults/Nymphs USSR; Washington wheat
Episyrphus balteatus Predator Adults/Nymphs USSR wheat
Eriopis connexa Predator Adults/Eggs/Larvae/Nymphs/Pupae
Erynia neoaphidis Pathogen Adults/Nymphs
Erynia radicans Pathogen Adults/Nymphs
Eupeodes corollae Predator Adults/Nymphs USSR
Eupeodes luniger Predator Adults/Eggs/Larvae/Nymphs/Pupae
Eupeodes volucris Predator Adults/Eggs/Larvae/Nymphs/Pupae
Exochomus concavus Predator Adults/Nymphs
Exochomus nigromaculatus Predator Adults/Nymphs
Exochomus quadripustulatus Predator Adults/Nymphs
Harmonia axyridis Predator Adults/Nymphs
Hippodamia Predator Adults/Nymphs
Hippodamia convergens Predator Adults/Nymphs South Africa wheat
Hippodamia sinuata Predator Adults/Nymphs
Hippodamia tredecimpunctata Predator Adults/Nymphs USSR wheat
Hippodamia undecimnotata Predator Adults/Nymphs North America
Hippodamia variegata Predator Adults/Nymphs North America
Ischiodon scutellaris Predator Adults/Nymphs
Lecanicillium lecanii Pathogen Adults/Nymphs
Leucopis Predator Adults/Nymphs
Leucopis caucasica Predator Adults/Nymphs USSR wheat
Leucopis glyphinivora Predator Adults/Nymphs USSR wheat
Leucopis ninae Predator Adults/Nymphs USSR wheat
Leucopis pallidolineata Predator Adults/Nymphs USSR wheat
Leucopis puncticornis Predator Adults/Eggs/Larvae/Nymphs/Pupae
Lioadalia Predator Adults/Eggs/Larvae/Nymphs/Pupae
Lioadalia flavomaculata Predator Adults/Nymphs
Lioadalia intermedia Predator Adults/Eggs/Larvae/Nymphs/Pupae
Lioadalia signifera Predator Adults/Eggs/Larvae/Nymphs/Pupae
Lysiphlebus testaceipes Parasite Adults/Nymphs
Malachius geniculatus Predator Adults/Nymphs USSR wheat
Meliscaeva auricollis Predator Adults/Eggs/Larvae/Nymphs/Pupae
Monoctonus washingtonensis Parasite Adults/Eggs/Larvae/Nymphs/Pupae
Nabis ferus Predator Adults/Nymphs USSR wheat
Nabis punctatus Predator Adults/Nymphs
Oenopia conglobata Predator Adults/Nymphs North America
Orius majusculus Predator Adults/Nymphs USSR wheat
Orius niger compressicornis Predator Adults/Eggs/Larvae/Nymphs/Pupae USSR wheat
Paecilomyces fumosoroseus Pathogen Adults/Eggs/Larvae/Nymphs/Pupae
Pandora neoaphidis Pathogen Adults/Nymphs
Platypalpus pictitarsis Predator Adults/Nymphs USSR wheat
Praon Parasite Adults/Eggs/Larvae/Nymphs/Pupae
Praon necans Parasite Adults/Eggs/Larvae/Nymphs/Pupae
Praon sp. nr callaphis Parasite Adults/Eggs/Larvae/Nymphs/Pupae
Praon volucre Parasite Adults/Nymphs USSR wheat
Praon yakimanum Parasite Adults/Eggs/Larvae/Nymphs/Pupae
Propylea quatuordecimpunctata Predator Adults/Nymphs
Propylea quatuordecimpunctata Predator Adults/Nymphs North America; USSR wheat
Psyllobora vigintiduopunctata Predator Adults/Eggs/Larvae/Nymphs/Pupae
Scaeva pyrastri Predator Adults/Nymphs USSR wheat
Scymnus Predator Adults/Nymphs
Scymnus caurinus Predator Adults/Eggs/Larvae/Nymphs/Pupae
Scymnus fenderi Predator Adults/Eggs/Larvae/Nymphs/Pupae
Scymnus frontalis Predator Adults/Nymphs North America
Scymnus morelleti Predator Adults/Nymphs
Scymnus nigrinus Predator Adults/Nymphs USSR wheat
Sphaerophoria menthastri Predator Adults/Nymphs USSR wheat
Sphaerophoria rueppellii Predator Adults/Eggs/Larvae/Nymphs/Pupae USSR wheat
Sphaerophoria scripta Predator Adults/Nymphs USSR wheat
Sphaerophoria sp. nr annulipes Predator Adults/Eggs/Larvae/Nymphs/Pupae
Syrphidae Predator Adults/Eggs/Larvae/Nymphs/Pupae
Syrphus Predator Adults/Nymphs

Notes on Natural Enemies

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Coccinellids were found associated with D. noxia throughout its Eurasian range (Hopper et al., 1997). Coccinella septempunctata was the species most frequently found in association with D. noxia, Hippodamia variegata the next most frequent, and Propylea quattuordecimpunctata the next most frequent (Hopper et al., 1997). Although some of the other species, for example Scymnus spp., are smaller and thus may be more able to attack D. noxia in rolled leaves (Kauffman and LaRoche, 1994), they were much rarer during exploration for introductions into USA. Yu and Liang (1998) described 17 species of coccinellid attacking D. noxia in Xinjiang, China. Both larvae and adults of coccinellids can prey on aphids; however, they attack other prey as well: for example, eggs of Lepidoptera. Furthermore, these predators do not in general show strong prey preferences or great differences in development when exposed to various aphid species (Michels and Flanders, 1992; Formusoh and Wilde, 1993).

Syrphids were associated with D. noxia throughout its Eurasian range (Hopper et al., 1997). The most common species were Episyrphis balteatus, Eupeodes corollae and Sphaerophoria scripta (Hopper et al., 1997). Syrphid adults eat nectar, honeydew and pollen, but their larvae eat aphids. The number of eggs laid by adult female E. balteatus and E. corollae increases with aphid abundance (Chambers, 1991). E. balteatus and E. corollae larvae can eat 413±5 and 311±6 D. noxia, respectively, over their development times of 9-10 and 7-8 days (Rojo et al., 1996). Furthermore, third instars can crawl 2 m/h and live 3-6 days without aphids (Rojo et al., 1996).

Leucopis spp. were found in association with D. noxia throughout its Eurasian range, however they were not as abundant as syrphids. Most have been identified as L. ninae. Like syrphids, adult Leucopis spp. eat nectar, honeydew, and pollen, but their larvae eat aphids, coccids, and psyllids (Tanasijtshuk, 1986). Although they occur in a variety of habitats, their small size has led some to argue that they should excel at attacking D. noxia in the rolled leaves which this aphid often causes. Adult female L. ninae can lay 20±4 eggs per day and 78±6 eggs over a 9-day period (Dabire, 1995). Females oviposit preferentially in aphid colonies and lay more eggs where aphid density is higher (Dabire, 1995). Although females tend to lay more eggs in leaf sheaths, neither they nor their larvae prefer D. noxia over other cereal aphids (Dabire, 1995). Larvae can consume 66±10 D. noxia over their development time of 9±1 days in the laboratory at 20°C. The number of aphids consumed has a great impact on larval development and survival and on adult size and fecundity (Dabire, 1995). Only 4% of second instars find aphid colonies 10 cm away (Dabire, 1995).

Aphelinus spp. were collected from D. noxia throughout most of its Eurasian range, although they were not found in several trips to the Middle East (Hopper et al., 1997). They are solitary endoparasitoids of nymphs and adults of many aphid species: A. asychis attacks at least 39 aphid species (Wilbert, 1964; Kalina and Stary, 1976). A. asychis females search for their hosts by walking and occasionally flitting from leaf to leaf. Females are only weakly attracted by volatiles from the plant-host complex (Farias, 1995). A. asychis females can parasitize 17±5 D. noxia per day and 230±30 D. noxia over their lifetime of 40±5 days in the laboratory at 20°C (K Hopper, Beneficial Insects Introduction Research Laboratory, USDA, USA, unpublished data). Adult females of Aphelinus spp. kill a few aphids per day by feeding on them to obtain nutrients for egg production (Cate et al., 1973, Bai and Mackauer 1990). At 20°C, Aphelinus spp. take about 3 weeks to develop from egg to adult emergence in D. noxia (Lajeunesse and Johnson, 1992). Wasp larvae kill their hosts and cause hardening of the aphid exoskeleton to form black, ellipsoidal mummies 7-10 days after oviposition at 20°C.

Aphelinus spp. are relatively rare on cereal aphids other than D. noxia (Rabasse and Dedryver, 1983). However, in the Montpellier region (of France), levels of parasitism of D. noxia by Aphelinus spp. can reach 55% in spring in wheat (Farias, 1995). The taxonomy of the genus Aphelinus needs revision. A. albipodus was only distinguished from A. varipes late in the project to introduce natural enemies for control of D. noxia in the USA; and it appears from reciprocal crosses among populations from different regions that what has been identified as A. asychis is also a species complex (Kazmer et al., 1996). A. varipes and A. albipodus abundances appear to vary inversely in space and time with abundance of the A. asychis complex (Farias, 1995). Although parasitism of D. noxia by Aphelinus spp. can reach high levels (Farias, 1995), there are often few mummies per colony (Chen and Hopper, 1997).

Parasitoid species in the braconid subfamily Aphidiinae were collected from D. noxia throughout its Eurasian range (Hopper et al., 1997). Diaeretiella rapae was the most frequently collected from D. noxia, and Aphidius spp. the next most frequently collected; other species were much more rare (Hopper et al., 1997). Aphidiines are solitary endoparasitoids of nymphs and adults of a wide variety of aphid species: D. rapae and Aphidius colemani collected from D. noxia successfully parasitized 7 and 9 species, respectively, of 14 aphid species to which they were exposed (Elliot et al., 1994a, b). Unlike aphelinids, they do not host feed. A. colemani develops in 12 days at 20°C and produces 316 offspring per female on D. noxia (Prinsloo et al., 1993). Aphidius matricariae develops on D. noxia in 273.1 ± 5.9 degree-days above a threshold of 4.5 ± 0.4°C (Miller and Gerth, 1994). D. rapae and Aphidius spp. can parasitize 40-100 aphids per day and 212-532 aphids over their lifetimes of 7-21 days in the laboratory at 20°C; D. rapae produces a maximum of 60 mummies per day at 21°C (Hågvar and Hofsvang, 1991). They are attracted to volatiles from the plant-host complex in general and from D. noxia on wheat in particular (Farias, 1995; Reed et al., 1995). Parasitism of D. noxia by aphidiines was often lower than that by aphelinids when both were present (Farias, 1995; Chen and Hopper, 1997).

Fungal pathogens were collected from D. noxia in fewer areas in Eurasia than predators and parasitoids (Hopper et al., 1997). This may be in part because some collectors lacked the training to collect and culture entomopathogenic fungi. Nonetheless, epizootics of fungal pathogens in D. noxia populations appear to be rare, perhaps because of the relatively arid environments in which this aphid is usually found (Wraight et al., 1993). However, D. noxia populations were found in the Jezreel Valley and upper Galilee Valley in Israel in 1994 that appeared to have been destroyed by fungal epizootics (Hopper et al., 1997). Prevalence of fungal pathogen infections has also been low in the US (Feng et al., 1991; Wraight et al., 1993). At least two viruses, aphid lethal paralysis virus and Rhopalosiphum padi virus, are known to attack D. noxia (Laubscher and Von Wechmar, 1991, 1992).

Impact Summary

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

Impact

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D. noxia has a great economic impact on cereal crops (Brooks et al., 1994). It is a phloem feeder like other aphids and the symptoms evident on plants are a result of this feeding mechanism. By feeding on the phloem, the aphid damages the plants through nutrient drainage (Dixon, 1985) which results in chlorosis, necrosis, wilting, stunting, and curling of the leaves, misshapen or nonappearance of new growth, and localised cell death at the site of aphid feeding. D. noxia further elicits an increase in essential amino acids in the phloem sap by triggering the breakdown of proteins in infested wheat leaves (Burd and Burton, 1992; du Toit, 1986; Ma et al., 1998; Miller et al., 2001). The damage to the foliar tissue is thought to play a role in the pest’s ability to increase nutritional quality of the host plant (Botha et al., 2006).

Barley and wheat are the most important cultivated hosts of D. noxia and considerable yield losses can occur in these crops. It is particularly injurious to late-sown barley in continental climates. Spring wheat suffers greatest yield loss when attacked during tillering to boot stage; winter wheat suffers greatest loss after vernalization (Gray et al., 1990). Early, heavy infestations on barley can cause total crop loss (Adisu et al., 2003) and significantly affect grain quality (Bregitzer et al., 2003). Aphid feeding on susceptible genotypes causes chlorosis and longitudinal streaking of leaves, and emerging leaves remain tightly rolled, which traps spikes and prevents their normal development (Mornhinweg, 2011). Infestation of wheat seedlings by D. noxia in the autumn can reduce the ability of seedlings to survive the winter (Thomas and Butts, 1990; Storlie et al., 1993). In general, damage is greatest when crop ripening coincides with peak aphid numbers.

D. noxia has been a pest of small grains in Russia since 1912, when the first serious outbreaks on barley were recorded. In the Crimea, D. noxia has reduced harvests by up to 75% in some years. In Eurasia generally, where it is thought to have originated, it is only occasionally a serious pest; with short-lived outbreaks being reported (Grossheim, 1914; Tuatay and Remaudière, 1964; Fernández et al., 1992).

Since its appearance in Texas in 1986 (Stoetzel, 1987), D. noxia has become a major pest of wheat and barley in the USA, causing over US$850 million in direct and indirect losses from 1987 to 1992 (Brooks et al., 1994). During the 1992/93 cropping season, over 7 million acres (20%) of dryland winter wheat and 1 million acres (33%) of barley were infested throughout the western USA (Webster and Amosson, 1994). In Canada, yield losses ranging from 25 to 37% without insecticide treatments in field trials (Butts et al., 1997).

Since its introduction in 1978, it has also become the major pest of wheat in South Africa (Walters et al., 1980), where it can cause over 80% yield loss if not controlled in the summer rainfall region (Tolmay and Prinsloo, 1994). In Ethiopia, it has been a major pest of barley for over two decades, with 20-30% yield loss in some areas (Haile and Megenasa, 1987). D. noxia is also a pest in Yemen (Erdelen, 1981).

Macedo et al. (2009) showed that D. noxia affected net photosynthesis of wheat. Wheat plants tolerant to D. noxia often exhibit increased photosynthetic rates, increased growth rates, increased stored root carbon and/or an enhanced ability to shunt stored carbon from roots to shoots (Burd and Elliott, 1996; Haile et al., 1999; Reidel and Blackmer, 1999; Smith and Chuang, 2013). In addition to direct feeding damage, major indirect losses in wheat and barley can be caused as a result of D. noxia transmitting Barley yellow dwarf virus (BYDV-PAV). However, D. noxia is a less effective vector of BYDV than Rhopalosiphum padi, R. maidis, Schizaphis graminum or Sitobion avenae (El-Yamani and Bencharki, 1997).

There have been conflicting reports on the effect of D. noxia on yield and yield components of host plants. For instance, in Canada, Ba-Angood and Stewart (1980) found that aphid feeding significantly reduced the number of grains per ear, whereas Milne and Delves (1999) in Australia found that aphid feeding did not affect grain numbers per ear. However, aphid feeding usually significantly reduces 1000-seed weight and grain yield per ha (e.g., Ba-Angood and Stewart, 1980; Milne and Delves, 1999; Damate, 2015). The percentage yield loss from D. noxia on irrigated wheat in Ethiopia ranged from 15 to 93%, depending on variety and season (Damate, 2015).

Damage ratings based on leaf chlorosis have frequently been used to evaluate resistance to D. noxia. However, some workers (e.g., Frank et al., 1989; Smith et al., 1991; Assad et al., 2004) suggest that leaf chlorosis may not be effective in separating resistant entries. To increase the accuracy of evaluation, leaf damage other than chlorosis, i.e. leaf rolling and leaf trapping, should also be considered (Nematollahy et al., 1998).

Detection and Inspection

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D. noxia is most easily detected in cereal crops by visual examination of plants. This aphid causes characteristic symptoms in cereals: leaves of infested plants are streaked and rolled. The streaks are white to purple, extend most of the long axis of the leaf, and are irregularly distributed across the short axis of the leaf. Purple, as opposed to white, streaks appear more frequently when the plant has been exposed to low temperatures. Rolling extends in severity from folding along the mid-vein, to one edge of the leaf rolled, to the leaf being rolled completely upon itself forming a straw-like tube. Heavily infested plants are stunted. Seed heads can be trapped by rolled flag leaves, giving the seed head a fish-hook shape. Symptoms are less severe on some wild grasses.

Like other aphids, alate D. noxia can be captured in suction traps, yellow pan traps, and sticky traps (Labonne et al., 1983; Aalbersberg et al., 1984; Melia et al., 1990; Armstrong et al., 1991; Halbert et al., 1992b; Basky, 1993). Nymphs and apterous adults are rarely captured in suction traps and are caught much less than alate adults in yellow pan traps and sticky traps (Labonne et al., 1983). D. noxia can also be sampled effectively by collecting plant material and placing it directly in Berlese funnels.

Similarities to Other Species/Conditions

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No other cereal aphids have both short, inconspicuous cornicles and a supracaudal process. Furthermore, no other pest causes the symptoms typical of D. noxia attack on cereals (see Symptoms, and Detection and Inspection Methods).

Prevention and Control

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Cultural Control

Walker and Peairs (1994) reviewed cultural control methods for D. noxia. These include planting date, planting density, furrow orientation, irrigation, fertilization, grazing, manipulation of crop residue, and alternative host plants.

Delaying planting beyond the period at the beginning of a season when alate D. noxia are abundant can reduce initial infestations. However, other agronomic constraints prevent large shifts in planting date. Although suction trap catches indicated sufficient autumn dispersal in Idaho (USA) to warrant delayed planting to avoid D. noxia infestation (Halbert et al., 1990), in more southerly regions, where early spring densities of alate D. noxia are higher, late-planted wheat may suffer more from spring infestations. D. noxia density is sometimes higher where plant density is lower (Walker and Peairs, 1994), so that increasing plant density could reduce D. noxia infestation.

North-South-oriented irrigation furrows in some cases give lower winter survival and slower development of D. noxia than East-West-oriented furrows because of reduced solar insolation (Hammon and Peairs, 1992). Thus, running irrigation furrows from north to south where possible in regions with marginal winter survival for D. noxia could reduce spring infestations and damage.

Nitrogen fertilization alone did not affect D. noxia numbers in several studies (Walker et al., 1991; Archer et al., 1995), although there was a shift to earlier reproduction with high-nitrogen fertilized wheat (Moon et al., 1995). On the other hand, Larson et al. (1990) found that nitrogen and sulphur together increased D. noxia infestation and Walker and Peairs (1994) found the opposite. It appears that individual nutrients can have varied effects depending on other nutrients, irrigation, and perhaps other variables. Irrigation in some cases reduced D. noxia population growth (Archer et al., 1995; Farias et al., 1995), but in others did not (Farias et al., 1995). A phenotypic study showed delayed development and less severity of symptoms with decreased numbers of aphids feeding on wheat plants treated with potassium phosphate in comparison with untreated plants for both resistant (Tugela DN) and susceptible (Scheepers) cultivars (Venter et al., 2014).

Although spring grazing can increase subsequent D. noxia population growth on the regrown plants (Messina et al., 1993a), it can also delay D. noxia growth and thus insecticide applications (Walker et al., 1990). Because D. noxia populations must survive the period between the harvest of one year's crop and the planting of the next, management of crop residues and alternative host plants appear promising. The current practice in much of the western USA is to plough under all crop residues and strictly control non-crop plants with herbicides. Although this does reduce reservoirs for D. noxia and other pests, it also reduces sources of natural enemies of these pests. Replacement of wild grasses currently used for forage and for conservation reserve programmes with grasses less suitable for D. noxia growth appears promising. Grass species vary in suitability for D. noxia development (Kindler et al., 1991a, 1992; Mowry et al., 1995). Furthermore, Kindler et al. (1991c, 1992) have found resistance to D. noxia within species of forage grasses. Fungal endophytes may provide some of this resistance (Kindler et al., 1991b; Clement et al., 1992).

Chemical Control

Imidacloprid is effective as a seed treatment for dryland wheat (Pike et al., 1993; van der Westhuizen et al., 1994), and Pike et al. (1993) consider it an environmentally safer approach than aerial or in-furrow applications of broader spectrum insecticides. When compared with several other foliar applications chlorpyrifos caused the greatest reduction in D. noxia density (Hill et al., 1993). Chlorpyrifos was effective at half the dose currently used in Canada, but residual impact on D. noxia survival was low (Hill et al., 1993).

Variation in D. noxia susceptibility to insecticides suggests the possibility that D. noxia may develop resistance to them (Brewer and Kaltenbach, 1995). However, Bayoun et al. (1995) found some insecticides not in current use against D. noxia that were effective for its control and relatively less toxic to natural enemies.

Biological Control

The limited evidence available suggests that indigenous natural enemies in South Africa and in the USA have had little impact to date on D. noxia population dynamics (Aalsbersberg et al., 1988b; Feng et al., 1992; Wraight et al., 1993). However, there are several reasons to expect that introducing Eurasian natural enemies would reduce D. noxia abundances in these countries. Introduced natural enemies have substantially controlled several introduced aphid pests, e.g. Therioaphis trifolii (spotted alfalfa aphid) and Chromaphis juglandicola (walnut aphid) (Laing and Hamai, 1976). Although these successes were in orchards or perennial crops, introduced natural enemies have also substantially reduced the abundance of introduced cereal pests, e.g. the cereal leaf beetle, Oulema melanopus in the USA (Haynes et al., 1974), and greenbug, Schizaphis graminum in Chile (Zuniga, 1990). Furthermore, a wide variety of predators and parasitoids attack cereal aphids, and many of these have been reported in association with D. noxia (Pike et al., 1991). Lastly, exclosure experiments suggest natural enemies (including predatory coccinellid, predatory syrphids and parasitoid wasps) play a major role in limiting D. noxia populations in southern France (Hopper et al., 1995).

Studies conducted from 1993 to 1994 in southeastern Wyoming (Brewer et al., 1998), after establishment of Aphelinus asychis and A. albipodus, documented high levels of parasitism of D. noxia by these parasitoids, especially A. albipodus. Lee et al. (2005) showed there may be regional differences in the importance of parasitoids in biological control of D. noxia.

Currently, D. rapae is being released for biological control of D. noxia. Studies of the impact of aphidophagous parasitoids and predators on D. noxia populations soon after invasion by the pest indicated that native natural enemies were ineffective at controlling D. noxia (Rice and Wilde, 1991; Wraight et al., 1993). In contrast, Hopper et al. (1995) observed that natural enemies played a significant role in suppressing D. noxia populations in Europe, where the Russian wheat aphid is presumably native (Lee et al., 2005). So far, four species of introduced parasitoids are known to have established in the USA: Aphelinus asychis (in Texas, Michels and Whitaker-Deerberg, 1993), Aphelinus albipodus, Aphidius colemani and A. uzbekistanicus (not from D. noxia; Halbert et al., 1996). No predators are known to have established.

Researchers in biological control at universities, state departments of agriculture, the US Department of Agriculture, and the International Institute for Biological Control carried out an extensive programme of foreign exploration and importation of predators, parasitoids and entomopathogens into the USA (e.g., Gonzàlez, 1989; Hopper et al., 1990; Gruber et al., 1991; Hopper, 1992; Gonzàlez et al., 1993; Hopper et al., 1993; Hopper et al., 1993; McKinnon et al., 1993; Gonzàlez et al., 1994; Hopper et al., 1994a, b[N1] ; Pike et al., 1994; Reed and Kindler, 1994; Tanigoshi et al., 1995).

Syrphids are promising candidates for D. noxia biological control because of their frequent co-occurrence with D. noxia and because of the great consumption rate and vigour of their larvae. Leucopis spp. are less promising candidates than syrphids because of the lower number of aphids their larvae consume and the lower capacity of their larvae to find aphid colonies. However, because of their relatively high fecundity, the dependence of oviposition on aphid density, and their potential to attack aphids in rolled leaves, they should not be neglected. Aphelinus spp. are promising candidates for D. noxia biological control because they are active during the seasons when D. noxia is active, they occur throughout most of the geographic range of D. noxia, and they live long and can kill a large number of aphids. Aphidiines appear to be less promising than aphelinids for biological control of D. noxia because they parasitize relatively few of these aphids. However, they have been successful in the control of other aphid pests and thus were included among the introductions into the USA.

A more limited programme of introductions has been conducted in South Africa (Tolmay and Prinsloo, 1994). This included introductions of Aphelinus hordei from the Ukraine (via CSIRO, Australia) and Leucopis ninae from Pakistan, Iran, and China (via USDA, USA). Aalbersberg et al. (1989) found that planting a refugium around wheat, to provide prey/hosts for D. noxia natural enemies, increased natural enemy abundance in wheat in South Africa. However, the natural enemies were still not able to keep D. noxia in check, partly because of poor timing of prey/host dynamics in the refugium. Hughes et al. (1994) released A. varipes into Australia as a pre-emptive measure against D. noxia which may eventually be introduced. However, there is no evidence that A. varipes established in Australia.

Feng and Johnson (1991) and Wang and Knudsen (1993) explored the use of the fungal pathogen Beauveria bassiana for D. noxia control. Wraight et al. (1993) suggested that use of pathogens to control D. noxia in the semi-arid western USA will require increased humidity, e.g. from irrigation.

Host-Plant Resistance

Resistance has been identified in barley and different wheat genotypes, including bread and non-bread wheat (e.g., Du Toit, 1989; Calhoun et al. 1991b; Quick et al., 1991; Smith et al., 1991; Souza et al., 1991; Robinson et al., 1991; Robinson, 1992a, 1993; Sarafrazi and Ahmadi, 1993; Formusoh et al., 1992, 1994; Nematollahy et al., 1998; Kazemi et al., 2001; Najafi-Mirak et al., 2003; Bregitzer et al., 2005, 2008; Hamedanian et al., 2010; Pourhaji and Ahmadi, 1999; Moharramipour et al., 2002; Mornhinweg et al., 2006, 2007a, 2007b, 2008, 2009, 2011; Pourhaji et al., 2011). Webster et al. (1991) published a uniform rating system for classifying foliar damage, but other rating systems have also been used (Du Toit, 1988; Nkongolo et al., 1990; Souza et al., 1991; Robinson et al., 1992a; Burd et al., 1993; Porter et al., 1993). Resistance to foliar damage symptoms correlated well with resistance to yield loss (Calhoun et al., 1991b). In vitro tests (Zemetra et al., 1993; Dong et al. 1994) and behavioural tests (Girma et al., 1992) for screening accessions for resistance to D. noxia have been developed.

Resistance in some wheat lines appears to be mostly from antibiosis with some tolerance (Du Toit, 1989; Quisenberry and Schotzko, 1994). One resistant wheat line showed moderate tolerance, high antibiosis, and little antixenosis (Baker et al., 1992). Other lines are tolerant with low levels of antibiosis (Smith et al., 1992). Nematollahy et al. (1999) showed that out of 15 bread and non-bread wheat genotypes, ‘5172’, a T. monococcum genotype, and ‘4898’ a T. aestivum line, had the highest level of tolerance and antibiosis resistance. Barley resistance is in part from antibiosis (D. noxia feed and grow less on resistant lines (Robinson et al., 1991, 1992b; Webster et al., 1993b) but also from antixenosis (Robinson, 1992b).

Resistance varies with environment, e.g. susceptible seedling survival in harsh winters is compromised by D. noxia infestation (Thomas and Butts, 1990; Storlie et al., 1993) so it is best to screen for resistance in the target environment (Calhoun et al., 1991b). Resistance may be related to concentration of hydroxamic acid (DIBOA) in barley (Gianoli and Niemeyer, 1998).

The genetic mechanisms of resistance in various accessions has been investigated (Du Toit, 1989; Nkongolo et al., 1991a, b, 1992; Robinson et al., 1992b; Elsidaig and Zwer, 1993; Marais et al., 1994; Mornhinweg et al., 1993; Schroeder-Teeter et al., 1993-4; Nieto-Lopez and Blake, 1994; Assad et al., 1999Najafi-Mirak et al., 2003; Fazel-Najafabadi et al., 2009, 2015). Six major genes have been identified (Quick, 1994). More resistance genes may be available in perennial Triticeae (Kindler et al., 1993), wild Hordeum spp. (Kindler and Springer, 1991), and triticale (Webster, 1990; Scott et al., 1991). Effects of resistance on other characters have been explored (Zwer et al., 1994). The physiological/biochemical basis of resistance have also been studied (Miller et al., 1994).

There is a lot of research on screening resistance to D. noxia in the laboratory and greenhouse at the seedling stage, but reports of field screening are scarce (e.g., Calhoun et al., 1991a, b; Nematollahi, 2010).Six resistant accessions have been released for commercial production in the USA (Quick, 1994). Two resistant cultivars have been released for commercial production in South Africa (Tolmay and Prinsloo, 1994). Screening of the more than 23,000 accessions in the USDA-ARS National Small Grains Collection (NSGC) has identified 116 accessions with some level of resistance to the United States biotype USA1 (Mornhinweg, 2011). From 40 of these accessions, 4 cultivars and 58 adapted germplasm lines have been developed and released (Mornhinweg et al., 2006, 2007a, b, 2008, 2009, 2011; Bregitzer et al., 2005, 2008). Several studies have examined the genetics of RWA resistance in barley. Two studies indicated that single dominant or semi-dominant genes provided RWA resistance in lines from CIMMYT tested with a Mexican RWA population (Robinson et al., 1992b) and from Iran tested with RWA from Iran (Assad et al., 1999).

Resistance and biotypes problem

Because virulence of D. noxia appears to vary within and between geographic regions (Bush et al., 1989; Puterka et al., 1992, 1993), new biotypes may arise that can overcome the defences of the new cultivars being released. This is especially true for antibiotic forms of resistance because these select for aphids that are not susceptible to antibiosis and may reduce the efficacy of natural enemies (Reed et al., 1992a, b). Resistant cultivars carrying Dn-resistance genes were classically used to control outbreaks of D. noxia (Botha and Venter, 2000). However, this control is increasingly more difficult, with the development of new D. noxia resistance-breaking biotypes putting pressure on farmers to revert to using pesticide applications to increase yields.

Since the occurrence of the first biotype (RWA2) in 2003, biotypes of D. noxia have become a serious threat to the development and deployment of plant resistance in cereals (Haley et al., 2004). Previous to the appearance of RWA2, resistance genes for wheat designated Dn1 to Dn9, Dnx and Dny were found to be effective against the original Russian wheat aphid (RWA1) that invaded the USA in 1986 (Haley et al., 2004). Since the discovery of RWA2, six additional biotypes (RWA3- RWA8) have been described (Burd et al., 2006; Weiland et al., 2008; Randolph et al., 2009).

Biotypic diversity within D. noxia populations has been identified in wheat based on differential reactions associated with the 11 identified genes and QTLs for resistance (Dn1-9, x, y). At least eight biotypes are present in the USA (USA1–8; Burd et al., 2006; Puterka et al., 2006; Weiland et al., 2008). USA1 and USA2 are prevalent in the seven western states most affected by D. noxia (Puterka et al., 2007). In a recent study, Puterka et al. (2014) found that biotypes RWA1, RWA2, RWA6 and RWA8 differed in virulence, while biotypes RWA3, RWA4, RWA5 and RWA7 produced similar virulence profiles. Thus, they consolidated these biotypes as RWA3/7, and indicated that in USA the five main biotypes RWA1, RWA2, RWA3/7, RWA6 and RWA8 can be identified using only four wheat genotypes containing Dn3, Dn4, Dn6 and Dn9.

Jankielsohn (2014) had been provided guidelines for the sampling, identification and designation of D. noxia biotypes in South Africa. There are currently four D. noxia biotypes recorded in South Africa. RWASA1 is the first biotype that was recorded in 1978. RWASA2, virulent against the Dn1 resistant gene in wheat, was recorded in 2005 in wheat-producing areas, especially in the Eastern Free State (Tolmay et al., 2007). RWASA3, virulent against the Dn4 resistant gene in wheat, was recorded in 2009, also predominantly in the Eastern Free State (Jankielsohn, 2014). During 2011 RWASA4, virulent against the Dn5 resistant gene, was recorded near Bethlehem in the Eastern Free State (Jankielsohn, 2014). Addi­tional biotypes have been identified in Hun­gary, Chile, Ethiopia, Argentina and the Czech Repub­lic that differentially affect resistance in wheat (Basky, 2003; Smith et al., 2004; Ricci et al., 2012).

The biotypic diversity in D. noxia has limited useful resistant germplasm to 94M370 (Dn7 gene), CI2401 and STARS 2414-11. In addition, two sources of resistance in barley, STARS 9301B and STARS 9577B, still exhibit strong resistance to all eight biotypes (Puterka et al., 2006). In a recent study, 23 lines resistant to all biotypes, included 10 unadapted germplasm accessions and 13 improved germplasm lines, providing useful germplasm for developing new barley cultivars with resistance to multiple D. noxia biotypes (Daheen et al., 2014).

Resistance genes in wheat appear to function in a biotype-specific manner with single genes providing resistance to one or more biotypes (Puterka et al., 2007; Weiland et al., 2008). In contrast, the two most widely-used barley germplasm lines for breeding resistant cultivars, STARS9301B and STARS9577B, show resistance to all eight United States biotypes, as defined by their differential reactions on wheat (Puterka et al., 2006; Weiland et al., 2008). So it seems that the apparent lack of biotype diversity with respect to resis­tance in barley is probably an artefact of insufficient study (Daheen et al., 2014).

Although new biotypes develop in various parts of the world it appears that there is very limited genetic variation between the different biotypes and also between their endosymbionts (Weiland et al., 2008; Swanevelder et al., 2010).

Integrated Pest Management

Integrated pest management (IPM) based on sampling pest density before deciding whether or not to treat requires cheap and rapid sampling plans for determining pest densities and accurate assessment of economic impact of these densities. Various plans have been developed for sampling D. noxia density (Schaalje et al., 1991; Feng and Nowierski, 1992; Feng et al., 1993, 1994; Nowierski et al., 1994), and for assessing whether D. noxia density exceeds an action threshold (Bechinski and Hohman, 1991; Legg et al., 1991, 1994; Butts and Schaalje, 1994). Legg et al. (1993) developed software for supporting decisions concerning chemical treatments.

Various relationships between D. noxia density and yield loss have been published. Du Toit (1986) reported an economic injury level of 14% infested plants when the ear had emerged from the flag leaf, and 4-7% infested plants when the first node had formed for winter wheat in South Africa. Girma et al. (1993) reported an economic injury level of 1-2 aphids per 7 plants for spring infestations, and 2-4 aphids per 7 plants for autumn infestations for Kansas wheat. Archer and Bynum (1992) found 4.6% yield loss for each 10% increase in damaged tillers and 4.8% yield loss for each 10% increase in infested tillers in dryland winter wheat in Colorado.

It may be possible to improve the level of aphid control by combining plant resistance and biological control (van Emden and Wratten, 1991). Control remains relatively limited when each method is applied separately. The compatibility between these management tactics was studied by Farid et al. (1998) and Brewer et al. (1999). Brewer et al. (1999) showed that the use of resistant barley in areas where the parasitoid, Diaretiella  rapae, is established is justified. In contrast, Reed et al. (1992) found that the development time of D. rapae was prolonged when it developed on D. noxia reared on resistant wheat cultivars with antibiotic factors compared to aphids reared on susceptible cultivars. It seems that plants can have a positive or negative effect on parasitoids, depending on the level and category of resistance (van Emden, 1991; Hare, 1992) and understanding the interactions that may arise from plant-aphid-parasitoid systems can therefore be important in developing a pest management system for these insects.

References

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07/11/15 Impact and Prevention and Control sections updated by:

Mohammad Reza Nematollahi, Assistant Professor of Entomology, Department of Plant Protection, Isfahan Research Center for Agriculture and Natural Resources, Isfahan, Iran.

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