Bemisia tabaci (tobacco whitefly)
- Summary of Invasiveness
- Taxonomic Tree
- Notes on Taxonomy and Nomenclature
- Distribution Table
- Risk of Introduction
- Hosts/Species Affected
- Host Plants and Other Plants Affected
- Growth Stages
- List of Symptoms/Signs
- Species Vectored
- Biology and Ecology
- Natural enemies
- Notes on Natural Enemies
- Means of Movement and Dispersal
- Impact Summary
- Environmental Impact
- Threatened Species
- Detection and Inspection
- Similarities to Other Species/Conditions
- Prevention and Control
- Distribution Maps
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PicturesTop of page
IdentityTop of page
Preferred Scientific Name
- Bemisia tabaci (Gennadius, 1889)
Preferred Common Name
- tobacco whitefly
Other Scientific Names
- Aleurodes inconspicua Quintance
- Aleurodes tabaci Gennadius
- Bemisia achyranthes Singh
- Bemisia bahiana Bondar
- Bemisia costa-limai Bondar
- Bemisia emiliae Corbett
- Bemisia goldingi Corbett
- Bemisia gossypiperda Misra & Lamba
- Bemisia gossypiperda mosaicivectura Ghesquiere
- Bemisia hibisci Takahashi
- Bemisia inconspicua (Quaintance)
- Bemisia longispina Priesner & Hosny
- Bemisia lonicerae Takahashi
- Bemisia manihotis Frappa
- Bemisia minima Danzig
- Bemisia minuscula Danzig
- Bemisia nigeriensis Corbett
- Bemisia rhodesiaensis Corbett
- Bemisia signata Bondar
- Bemisia vayssieri Frappa
International Common Names
- English: cassava whitefly; cotton whitefly; silver leaf whitefly; sweet potato whitefly
- Spanish: mosca blanca; mosca blanca del algodonero; mosca blanca del camote; mosca blanca del tabaco; mosquita blanca del tabaco
- French: aleurode de la patate douce; aleurode du cotonnier
- Portuguese: mosca branca do feijao
Local Common Names
- Germany: Baumwoll-Mottenschildlaus; Tabak-Mottenschildlaus; Weisse Fliege
- Israel: knimat ash hatabak
- Italy: aleirode delle solanacee; aleurode delle solanacee
- Turkey: beyaz sinek
- BEMIBA (Bemisia bahiana)
- BEMIEM (Bemisia emiliae)
- BEMIGO (Bemisia goldingi)
- BEMIIN (Bemisia inconspicua)
- BEMILO (Bemisia longispina)
- BEMIMA (Bemisia manihotis)
- BEMINI (Bemisia nigeriensis)
- BEMIRH (Bemisia rhodesiaensis)
- BEMITA (Bemisia tabaci)
- BEMIVA (Bemisia vayssieri)
Summary of InvasivenessTop of page
The Bemisia tabaci complex is polyphagous and now attacks many crops, but without significant impact on land use. Any effects on biodiversity would result indirectly from an increased use of insecticides against this pest.
Taxonomic TreeTop of page
- Domain: Eukaryota
- Kingdom: Metazoa
- Phylum: Arthropoda
- Subphylum: Uniramia
- Class: Insecta
- Order: Hemiptera
- Suborder: Sternorrhyncha
- Unknown: Aleyrodoidea
- Family: Aleyrodidae
- Genus: Bemisia
- Species: Bemisia tabaci
Notes on Taxonomy and NomenclatureTop of page
The genus Bemisia contains 37 species and is thought to have originated from Asia (Mound and Halsey, 1978). Bemisia tabaci, being possibly of Indian origin (Fishpool and Burban, 1994), was described under numerous names before its morphological variability was recognised. For full synonyms, see Mound and Halsey (1978). Originally, three distinct groups of B. tabaci were identified by comparing their mitochondrial 16S ribosomal subunits: New World; India/Sudan; and remaining Old World (Frohlich and Brown, 1994). The pest status of B. tabaci insects has now become more complicated and through the comparison of the mitochondrial cytochrome oxidase 1 (mtCO1) gene it is generally accepted that, rather than one complex species, B. tabaci is a complex of 11 genetic groups. These genetic groups are composed of at least 34 morphologically indistinguishable species, which are merely separated by a minimum of 3.5% mtCOI nucleotide divergence (Dinsdale et al., 2010; De Barro et al., 2011; Boykin and De Barro, 2014). First reports of a newly-evolved biotype of B. tabaci, the B biotype (see separate datasheet, now widely accepted, and known as, Middle East-Asia Minor 1 species (MEAM1)), appeared in the mid-1980s (Brown et al., 1995b). This species, commonly referred to as the silverleaf whitefly or poinsettia strain, is highly polyphagous and almost twice as fecund as previously recorded strains, and has been documented as being a separate species, B. argentifolii (Bellows et al., 1994). MEAM1 is able to cause phytotoxic disorders in certain plant species, for example, silverleaf in squashes (Cucurbita sp.) and this is an irrefutable method of identification (Bedford et al., 1992, 1994a). It can also can transfer and infect tomatoes with both Tomato yellow leaf curl virus (TYLCV) and Tomato yellow leaf curl Sardinia virus (TYLCSV) and, depending on the timing of infection, losses can reach 100%.
A distinctive, non-specific esterase banding pattern is also helpful in identification (Brown et al., 1995a) but is not infallible (Byrne et al., 1995). A recent study by Rosell et al. (1997) which used SEM to examine the morphological characters documented by Bellows et al. (1994) for identifying the 'B biotype' showed that most Old World populations of B. tabaci were morphologically indistinguishable from the 'B biotype'. These Old World populations did not induce silverleaf disorders or produce similar esterase banding patterns to B. argentifolii. Several other 'biotypes' (up to S) have now been described (Brown et al., 1995b, 1999; Banks et al., 1999; Dinsdale et al., 2010; De Barro et al., 2011; Boykin and De Barro, 2014) which supports the idea of a species complex, rather than of a number of distinct species, such as B. argentifolii. However, within the New World, MEAM1 has been readily accepted as a new species. Even though a recent study has irrefutably shown that MEAM1 can be crossed with a non-B biotype (Mediterranean species (formerly known as biotype Q) from Spain) (Adan et al., 1999).
DescriptionTop of page
Pear shaped with a pedicel spike at the base, approximately 0.2 mm long.
Yellow-white scales, 0.3-0.6 mm long.
Flat, irregular oval shape, 0.7 mm long. On a smooth leaf the puparium lacks enlarged dorsal setae, but if the leaf is hairy, two to eight long dorsal setae are present.
About 1 mm long, the male slightly smaller than the female. The body and both pairs of wings are covered with a powdery, waxy secretion, white to slightly yellowish in colour.
DistributionTop of page
B. tabaci has a global presence. However, certain areas within Europe are still Bemisia free, e.g. Finland, Sweden, Republic of Ireland and the UK (Cuthbertson and Vänninen, 2015).
See also CABI/EPPO (1998, No. 34).
Distribution TableTop of page
The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.Last updated: 28 Jul 2021
Risk of IntroductionTop of page
B. tabaci is regulated by the European Union (EU, 2000) and by other EPPO countries (Belarus, Russia). It is listed in the European Union (EU) Plant Health Directive 2000/29/EC under Annex 1AI (non-European populations) as a harmful organism, whose introduction from non-EU countries into, and spread within, all EU member states shall be banned. Some areas in the EU (British Isles, Nordic countries, parts of Portugal) are maintained as 'protected zones' (Cuthbertson and Vänninen, 2015). B. tabaci also presents a risk to countries in Central America, the Caribbean, Africa and South America. It is already widespread in Asia and most tropical areas. The risk is primarily to the glasshouse industry in northern countries (Bedford et al., 1994b; Cuthbertson, 2013) and mainly concerns MEAM1 species. Since its recent introduction to several of these countries, the pest has proved particularly difficult to combat because of its polyphagy, its resistance to many insecticides and its disruption of biological control programmes (Della Giustina et al., 1989). Very few countries remain free from B. tabaci, illustrating the difficulty of preventing its movement in international trade. Furthermore, it is likely that various species of B. tabaci complex are already present, but unreported, as pests of field crops in other countries. In principle, the introduction of new biotypes into areas where the A biotype has long been present does present a risk, but it is one that is very difficult to manage.
In addition, because B. tabaci is the vector of a number of mainly tropical begomoviruses, temperate areas face the risk that these viruses, of which certain ones are listed, for example, in EU regulations (EU, 2000) will enter with their vector. The EU requires special measures to deal with this additional risk.
Hosts/Species AffectedTop of page
Until recently, B. tabaci was mainly known as a pest of field crops in tropical and sub-tropical countries, on cassava, cotton, sweet potatoes, tobacco and tomatoes. Non MEAM1 B. tabaci populations, in nearly all cases, have a narrow plant host range within the species shown in the tables and may include many obscure indigenous weed species. Some non MEAM1 species have been shown to be monophagous. However, a non-MEAM1 species within a country could have a composite host range of many plant and crop species.
Only MEAM1 species are presently documented as being almost polyphagous, although recent laboratory studies have indicated that only a small number of individuals within some populations are able readily to change hosts. The progeny from these individuals have been shown to be highly polyphagous (Bedford et al., 1996).
Host Plants and Other Plants AffectedTop of page
Growth StagesTop of page
SymptomsTop of page
B. tabaci can acquire and transmit a range of plant viruses (see Economic Impact) which produce a variety of different symptoms on susceptible plant species. Although plants can become infected from migratory feeding of B. tabaci, plants infected with B. tabaci-transmitted viruses are often indicative of B. tabaci colonization.
Infected plants could exhibit any one or a combination of the following symptoms: vein yellowing, inter-vein yellowing, leaf yellowing, yellow blotching of leaves, yellow mosaic of leaves, leaf curling, leaf crumpling, leaf vein thickening, leaf enations, leaf cupping, stem twisting, plant stunting.
List of Symptoms/SignsTop of page
|Leaves / honeydew or sooty mould|
Species VectoredTop of page
African cassava mosaic virus (African cassava mosaic)
Ageratum enation virus
Bean calico mosaic virus
Bean dwarf mosaic virus
Bean golden mosaic virus (BGMV-type 1)
Bean golden yellow mosaic virus (bean golden yellow mosaic)
Bean yellow disorder virus
Bhendi yellow vein mosaic virus
Cabbage leaf curl virus
Cassava brown streak viruses (cassava brown streak disease)
Chayote yellow mosaic virus
Chino del tomate virus
Cotton leaf curl disease complex (leaf curl disease of cotton)
Cotton leaf curl Gezira virus
Cowpea golden mosaic virus
Cowpea mild mottle virus (angular mosaic of beans)
Croton yellow vein mosaic virus
Cucumber vein yellowing virus (cucumber vein yellowing)
Cucurbit chlorotic yellows virus
Cucurbit yellow stunting disorder virus
Dicliptera yellow mottle virus
Dolichos yellow mosaic virus
East African cassava mosaic Cameroon virus
East African cassava mosaic Malawi virus
East African cassava mosaic virus
East African cassava mosaic Zanzibar virus
Euphorbia leaf curl virus
Euphorbia mosaic virus
Hollyhock leaf crumple virus
Honeysuckle yellow vein virus
Horsegram Yellow Mosaic Virus
Indian cassava mosaic virus (Indian cassava mosaic)
Ipomoea yellow vein virus
Lettuce chorosis virus
Lettuce infectious yellows virus (infectious yellows of lettuce)
Luffa yellow mosaic virus
Macroptilium mosaic Puerto Rico virus
Macroptilium yellow mosaic Florida virus
Macroptilium yellow mosaic virus
Malvastrum yellow vein virus
Melon chlorotic leaf curl virus
Melon yellowing-associated virus
Mungbean yellow mosaic India virus
Mungbean yellow mosaic virus
Okra yellow vein mosaic virus
Papaya leaf curl China virus
Papaya leaf curl Guandong virus
Papaya leaf curl virus
Pepper golden mosaic virus
Pepper huasteco yellow vein virus
Pepper leaf curl Bangladesh virus
Pepper leaf curl virus
Pepper yellow vein Mali virus
Potato yellow mosaic Panama virus
Potato yellow mosaic virus
Radish leaf curl virus
Rhynchosia golden mosaic virus
Sida golden mosaic Costa Rica virus
Sida golden mosaic Florida virus
Sida golden mosaic Honduras virus
Sida golden mosaic virus
Sida golden yellow vein virus
Sida micrantha mosaic virus
Sida mottle virus
Sida yellow mosaic virus
Sida yellow vein virus
South African cassava mosaic virus
Soybean crinkle leaf virus
Squash leaf curl China virus
Squash leaf curl Philippines virus
Squash leaf curl virus (leaf curl of squash)
Squash leaf curl Yunnan virus
Squash mild leaf curl virus
Squash vein yellowing virus
Sri Lankan cassava mosaic virus
Stachytarpheta leaf curl virus
Sweet potato chlorotic stunt virus
Sweet potato leaf curl Georgia virus
Sweet potato leaf curl virus
Sweet potato mild mottle virus (mild mottle of sweet potato)
Tobacco curly shoot virus
Tobacco leaf curl Japan virus
Tobacco leaf curl Yunnan virus
Tobacco leaf curl Zimbabwe virus
Tomato chino La Paz virus
Tomato chlorosis virus (yellow leaf disorder of tomato)
Tomato chlorotic mottle virus
Tomato curly stunt virus
Tomato golden mosaic virus
Tomato golden mottle virus
Tomato leaf curl Bangalore virus
Tomato leaf curl Bangladesh virus
Tomato leaf curl China virus
Tomato leaf curl Gujarat virus
Tomato leaf curl Karnataka virus
Tomato leaf curl Laos virus
Tomato leaf curl Malaysia virus
Tomato leaf curl Mali virus
Tomato leaf curl New Delhi virus (Tomato New Delhi virus)
Tomato leaf curl Philippines virus
Tomato leaf curl Sinaloa virus
Tomato leaf curl Sri Lanka virus
Tomato leaf curl Sudan virus
Tomato leaf curl Taiwan virus
Tomato leaf curl Vietnam virus
Tomato mild mottle virus
Tomato mosaic Havana virus
Tomato mottle virus
Tomato rugose mosaic virus
Tomato severe leaf curl virus
Tomato severe rugose virus
Tomato torrado virus
Tomato yellow leaf curl China virus
Tomato yellow leaf curl Kanchanaburi virus
Tomato yellow leaf curl Malaga virus
Tomato yellow leaf curl Mali virus
Tomato yellow leaf curl Sardinia virus (Tomato yellow leaf curl virus - European strain)
Tomato yellow leaf curl Thailand virus
Tomato yellow leaf curl virus (leaf curl)
Tomato yellow vein streak virus
Watermelon chlorotic stunt virus
Biology and EcologyTop of page
Eggs are laid usually in circular groups, on the undersides of leaves, with the broad end touching the surface and the long axis perpendicular to the leaf. They are anchored by a pedicel inserted into a fine slit made by the female, and not into stomata as in the case of many other aleyrodids. Eggs are whitish in colour when first laid, but gradually turn brown. Each female lays up to 160 eggs. Hatching occurs after 5-9 days at 30°C depending on host species, temperature and humidity.
On hatching, the first instar or 'crawler' is flat, oval and scale-like, and is the only mobile larval stage. It moves to a suitable feeding location on the lower leaf surface where it moults and becomes sessile throughout the remaining nymphal stages. The first three nymphal stages last 2-4 days each (depending on temperature). The fourth nymphal stage is termed the puparium, and is approximately 0.7 mm long. True pupation within the whitefly life-cycle is debatable as it does not occur in other Homopterous families, although the last stage of the fourth nymphal instar after apolysis has been completed is typically referred to as a pupa. Pupation lasts for about 6 days and within the latter period, the metamorphosis to adult occurs.
The adult emerges through a 'T'-shaped rupture in the puparium and expands its wings before powdering itself with wax from glands on the abdomen. Copulation begins 12-20 hours after emergence and takes place several times throughout the life of the adult. A female may live for 60 days, although the life of the male is generally much shorter, being between 9 to 17 days. Some 11 to 15 generations can occur within one year.
Natural enemiesTop of page
|Natural enemy||Type||Life stages||Specificity||References||Biological control in||Biological control on|
|Agistemus exsertus||Predator||Adults; Arthropods|Nymphs|
|Amblyseius aleyrodis||Predator||Adults; Arthropods|Nymphs|
|Amblyseius limonicus||Predator||Arthropods|Nymphs||Cuthbertson (2014)||UK||poinsettia plants|
|Amblyseius swirskii||Predator||Arthropods|Nymphs||Cuthbertson (2014)||UK||poinsettia plants|
|Aschersonia aleyrodes||Pathogen||Adults; Arthropods|Nymphs|
|Bacillus thuringiensis kurstaki||Pathogen||Adults; Arthropods|Nymphs|
|Bacillus thuringiensis thuringiensis||Pathogen||Adults; Arthropods|Nymphs|
|Beauveria bassiana||Pathogen||Cuthbertson et al. (2012)||UK||poinsettia plants|
|Campylomma nicolasi||Predator||Adults; Arthropods|Nymphs|
|Chrysoperla carnea||Predator||Adults; Arthropods|Nymphs|
|Chrysoperla rufilabris||Predator||Adults; Arthropods|Nymphs|
|Coccinella septempunctata||Predator||Adults; Arthropods|Nymphs|
|Coccinella undecimpunctata||Predator||Adults; Arthropods|Nymphs|
|Coenosia attenuata||Predator||Adults; Arthropods|Nymphs|
|Collops vittatus||Predator||Adults; Arthropods|Nymphs|
|Cybocephalus micans||Predator||Adults; Arthropods|Nymphs|
|Delphastus pusillus||Predator||Adults; Arthropods|Nymphs||California|
|Deraeocoris pallens||Predator||Adults; Arthropods|Nymphs|
|Encarsia adrianae||Parasite||Arthropods|Nymphs||Pakistan||beans; Lantana camara|
|Encarsia aleurochitonis||Parasite||Adults; Arthropods|Nymphs|
|Encarsia cibcensis||Parasite||Arthropods|Nymphs||Pakistan||beans; Lantana camara|
|Encarsia formosa||Parasite||Arthropods|Nymphs||Israel; New Zealand; Norway||ornamental plants|
|Encarsia lutea||Parasite||Arthropods|Nymphs||Egypt||soyabeans; tomatoes|
|Encarsia mohyuddini||Parasite||Adults; Arthropods|Nymphs|
|Encarsia nigricephala||Parasite||Adults; Arthropods|Nymphs|
|Encarsia pergandiella||Parasite||Adults; Arthropods|Nymphs|
|Encarsia reticulata||Parasite||Adults; Arthropods|Nymphs|
|Encarsia transvena||Parasite||Adults; Arthropods|Nymphs|
|Encarsia tricolor||Parasite||Adults; Arthropods|Nymphs|
|Eretmocerus aligarhensis||Parasite||Adults; Arthropods|Nymphs|
|Eretmocerus corni||Parasite||Adults; Arthropods|Nymphs; Arthropods|Pupae||Paraguay||cotton|
|Eretmocerus diversiciliatus||Parasite||Adults; Arthropods|Nymphs|
|Eretmocerus eremicus||Parasite||Adults; Arthropods|Nymphs|
|Eretmocerus haldemani||Parasite||Adults; Arthropods|Nymphs|
|Eretmocerus mundus||Parasite||Adults; Arthropods|Nymphs; Arthropods|Pupae||Egypt; Mali||cotton; soyabeans; tomatoes|
|Eupeodes corollae||Predator||Adults; Arthropods|Nymphs|
|Euseius hibisci||Predator||Adults; Arthropods|Nymphs|
|Euseius scutalis||Predator||Adults; Arthropods|Nymphs||Morocco||Citrus|
|Franklinothrips vespiformis||Predator||Adults; Arthropods|Nymphs||Paraguay||cotton|
|Geocoris ochropterus||Predator||Adults; Arthropods|Nymphs|
|Geocoris punctipes||Predator||Adults; Arthropods|Nymphs|
|Hippodamia convergens||Predator||Adults; Arthropods|Nymphs|
|Labidura riparia||Predator||Adults; Arthropods|Nymphs|
|Laius venustus||Predator||Adults; Arthropods|Nymphs||Sudan||cotton|
|Lecanicillium lecanii||Pathogen||Adults; Arthropods|Nymphs|
|Mallada boninensis||Predator||Adults; Arthropods|Nymphs|
|Metaseiulus occidentalis||Predator||Adults; Arthropods|Nymphs|
|Microlestes discoidalis||Predator||Adults; Arthropods|Nymphs||Sudan||cotton|
|Nabis alternatus||Predator||Adults; Arthropods|Nymphs|
|Nabis capsiformis||Predator||Adults; Arthropods|Nymphs||Sudan||cotton|
|Nephaspis maesi||Predator||Adults; Arthropods|Nymphs||Nicaragua||Citrus; pawpaws|
|Orius albidipennis||Predator||Adults; Arthropods|Larvae; Arthropods|Nymphs||Sudan||cotton|
|Orius tristicolor||Predator||Adults; Arthropods|Nymphs|
|Paecilomyces farinosus||Pathogen||Adults; Arthropods|Nymphs|
|Paederus alfierii||Predator||Adults; Arthropods|Nymphs|
|Paragus compeditus||Predator||Adults; Arthropods|Nymphs|
|Phidippus audax||Predator||Adults; Arthropods|Nymphs|
|Scymnus syriacus||Predator||Adults; Arthropods|Nymphs|
|Serangium parcesetosum||Predator||Eggs; Arthropods|Larvae; Arthropods|Pupae|
|Sinea confusa||Predator||Adults; Arthropods|Nymphs|
|Sphaerophoria rueppellii||Predator||Adults; Arthropods|Nymphs|
|Transeius montdorensis||Predator||Arthropods|Nymphs||Cuthbertson (2014)||UK||poinsettia plants|
|Typhlodromus athiasae||Predator||Adults; Arthropods|Nymphs|
|Typhlodromus sudanicus||Predator||Adults; Arthropods|Nymphs|
|Zelus renardii||Predator||Adults; Arthropods|Nymphs|
Notes on Natural EnemiesTop of page
The species of Encarsia recorded as parasitoids of B. tabaci were revised by Polaszek et al. (1992), and also summarized by Cock (1993). They recognised 18 species, one or more of which are usually found parasitizing B. tabaci wherever natural enemies have been studied. Four additional Encarsia spp. parasitoids were described by Evans and Polaszek (1997).
The other important parasitoids attacking B. tabaci belong to the genus Eretmocerus. In each region one or more species of each of these two genera cause heavy mortality. There are also numerous records of generalist predators of Homoptera recorded as attacking B. tabaci (Cock, 1986, 1993). However, the combined impact of these natural enemies is insufficient to prevent virus transmission, but may be adequate to prevent losses where direct feeding damage is important. These natural enemies are all susceptible to insecticides and injudicious application has caused devastating resurgence, notably on cotton, for example, in the Sudan (Eveleens, 1983).
An isolate of the parasitoid Isaria fumosorosea has shown potential to be further developed as a biopesticide for controlling B. tabaci (Rahim Eslamizadeh et al., 2013).
Various species of predatory mites have also been shown to be effective in reducing B. tabaci populations, including Amblyseius limonicus, A. swirskii and Transeius montdorensis (Cuthbertson, 2014). A large range of natural enemies of B. tabaci have been recorded in China (Li et al., 2011).
Means of Movement and DispersalTop of page
Adults of B. tabaci do not fly very efficiently but, once airborne, can be transported quite long distances by the wind. All stages of the pest are liable to be carried on planting material and cut flowers of host species. The international trade in poinsettia is considered to have been a major means of dissemination of MEAM1 species of B. tabaci within the EPPO region (Cuthbertson, 2013).
Impact SummaryTop of page
|Fisheries / aquaculture||None|
ImpactTop of page
The pest status of B. tabaci insects is complicated and through the comparison of mitochondrial cytochrome oxidase 1 (mtCO1) gene it is generally accepted that, rather than one complex species, B. tabaci is a complex of 11 genetic groups. These genetic groups are composed of at least 34 morphologically indistinguishable species, which are merely separated by a minimum of 3.5% mtCOI nucleotide divergence (Dinsdale et al., 2010; De Barro et al., 2011; Boykin and De Barro, 2014). Within the B. tabaci complex, the Middle East-Asia Minor 1 (MEAM1) cryptic species, formerly referred to as 'B' and Mediterranean (MED) cryptic species, formerly referred to as 'Q' biotype that are the two most widely distributed, and as a result, best known of the species. These two species present the greatest threat to glasshouse crops worldwide (Bethke et al., 2009). The damaging MEAM1 is described as an aggressive coloniser and is an effective vector of many viruses, whereas the MED characteristically shows strong resistance to novel insecticides (Jones et al., 2008; McKenzie et al., 2009).
B. tabaci has been recorded as a minor pest of cotton and other tropical or semi-tropical crops within the warmer parts of the world and, until recently has been successfully managed with a range of insecticides.
A few biotypes from certain areas have become major pests, often within large mono-cropping areas where they are regularly exposed to insecticides. In these cases, the biotypes have rapidly evolved resistance to almost all currently available insecticides (Cahill et al., 1996; Mushtaq Ahmad et al., 2002; Luo et al., 2010; Wang et al., 2010). Exposure to sustained insecticide treatments may have promoted other characteristics of these 'pest' biotypes, such as increased fecundity and host adaptability. Populations of the cosmopolitan MEAM1 species [see datasheet on B. tabaci (MEAM1)], the Pakistan K biotype and the Mediterranean species are currently within this group. Other presently uncharacterized biotypes within Africa appear specifically adapted to cassava, causing severe losses to this important subsistence crop (Maruthi et al., 2000).
The feeding of B. tabaci adults and nymphs causes chlorotic spots to appear on the surface of the leaves. Depending on the level of infestation, these spots may coalesce until the whole of the leaf is yellow, apart from the area immediately around the veins. Such leaves are later shed. The honeydew produced by the feeding of the nymphs covers the surface of the leaves and can cause a reduction in photosynthetic potential when colonized by moulds. Honeydew can also disfigure flowers and, in the case of cotton, can cause problems in processing the lint. With heavy infestations, plant height, number of internodes and quality and quantity of yield can be affected (for example, in cotton).
Most biotypes of B. tabaci can vector over 60 plant viruses in the genera Geminivirus, Closterovirus, Nepovirus, Carlavirus, Potyvirus and a rod-shaped DNA virus (Markham et al., 1994; Alegbejo, 2000). Those biotypes that are poor vectors, appear so, due to their inability to feed on alternative host plant species (Bedford et al., 1994b). Whitefly-transmitted geminiviruses, now called begomoviruses, are by far the most important agriculturally, causing yield losses to crops of between 20 and 100% (Brown and Bird, 1992; Cathrin and Ghanim, 2014). Begomoviruses cause a range of different symptoms that include yellow mosaics, yellow veining, leaf curling, stunting and vein thickening. Presently a million ha of cotton is being decimated in Pakistan by cotton leaf curl disease (CLCuV) (Mansoor et al., 1993) and important African subsistence crops such as cassava are affected by begomoviruses such as African cassava mosaic virus (ACMV). Tomato crops throughout the world are particularly susceptible to many different begomoviruses, and in most cases exhibit yellow leaf curl symptoms. This has caused their initial characterization as Tomato yellow leaf curl virus (TYLCV). TYLCV has also recently been recorded in the New World, as well as several other begomoviruses such as Tomato mottle virus (EPPO/CABI, 1996), Tobacco leaf curl virus (TLCV), Sida golden mosaic virus (SiGMV), Squash leaf curl virus (SLCV), Cotton leaf crumple virus (CLCV) and Bean golden mosaic virus (BGMV) some of which cause heavy yield losses in their respective hosts. Dual infections have also been shown to occur (Bedford et al., 1994c).
Europe presently has five known begomoviruses. Three have been shown to no longer be transmissible by B. tabaci: Honeysuckle yellow vein mosaic (also known as Tobacco leaf curl virus), Abutilon mosaic virus (Bedford et al., 1994a) and Ipomea yellow vein virus (Banks et al., 1999), possibly through many years of vegetative propagation of their ornamental host plants. The others are two different transmissible TYLCVs that are causing major crop losses within the tomato industries of Spain, Portugal, Italy and the Canary Islands. Indigenous weed species such as Solanum nigrum and Datura stramonium have also been shown as field reservoirs for these tomato viruses (Bedford et al., 1998) and may be the source of others yet to be identified within Europe. Two B. tabaci-transmitted closteroviruses are also now affecting European crops, including those in the Canary Islands. Cucurbit yellow stunting disorder, is causing severe damage to cucumbers and melons in southern Europe (Celix et al., 1996), along with Tomato chlorosis virus (Navas-Castillo et al., 2000). There are also reports of a third closterovirus, Tomato infectious chlorosis virus, in Europe (Duffus et al., 1996) although this virus currently appears not to be of economic significance. However, a Bemisia-transmitted potyvirus, Cucumber yellow vein virus, appeared in cucumber crops in southern Spain for the first time in 2000 (Cuadrado et al., 2001). Despite a crop destruction programme to eradicate this virus, it has recently spread to melon crops in the region. Protected Zones (e.g. UK and Finland) within Europe remain free from damaging begomoviruses (Cuthbertson and Vänninen, 2015).
In Pakistan, the K biotype is responsible for the spread of a decimating viral disease of cotton, cotton leaf curl disease (CLCuD) (Briddon and Markham, 2000). This disease first became a serious problem in the early 1990s, rapidly affecting a million ha of cotton, which comprises 60% of the country's foreign exchange (Mansoor et al., 1993).
Mediterranean species (Biotype Q)
The Mediterranean species (formerly known as Q biotype) is found throughout the Iberian peninsula, around the Mediterranean basin (including Israel) and in the Canary Islands. It is widely thought that this is the indigenous biotype to these regions, although it co-exists with MEAM1 species in Israel, Italy and the Canary Islands. A population of MEAM 1 was recorded within a Mediterranean species population around Almeria in southern Spain in 1995. It appears that this population failed to become established since recent surveys have only identified Mediterranean species. Mediterranean species was first recorded entering Guatemala in 2009 (Bethke et al., 2009) and the UK in 2012 (Powell et al., 2012). Mediterranean species has, over recent years, been exposed to extensive insecticide applications and within areas of intensive agriculture exhibits a high level of resistance (Dennehy et al., 2010). The use of IPM control programmes is presently restricted where crops are susceptible to viruses. This is particularly the case with Tomato yellow leaf curl viruses which are transmitted very efficiently by B. tabaci. Because of insecticide resistance, large numbers of Mediterranean species often infest crops within southern Europe, resulting in rapid spread of viruses to newly planted crops. Field grown tomato crops in areas of southern Spain and Morocco have recently suffered 100% losses and TYLCV has spread to Phaseolus vulgaris and Capsicum annuum crops.
Environmental ImpactTop of page
The impact of the B. tabaci multi-species complex has mainly been in glasshouses in temperate countries, where Trialeurodes vaporariorum already presented problems. Any additional problems caused by B. tabaci, in terms of changes in crops cultivated or in the use of new control measures, have been essentially in this protected environment and cannot be said to impact the natural environment.
B. tabaci has also proliferated on outdoor crops in warmer countries. There is no particular indication that it has changed the crops cultivated or land use, but its control with insecticides has added to the general pesticide load on the environment.
Threatened SpeciesTop of page
|Threatened Species||Conservation Status||Where Threatened||Mechanism||References||Notes|
|Allactaga alaster||No Details|
Detection and InspectionTop of page
Numerous chlorotic spots develop on the leaves of affected plants, which may also be disfigured by honeydew and associated sooty moulds. Leaf curling, yellowing, mosaics or yellow veining could also indicate the presence of whitefly-transmitted viruses.
Close observation of the undersides of the leaves will show the tiny, yellow/white larval scales and in severe infestations, when the plant is shaken, numerous small, white adult whiteflies will flutter out and quickly resettle. These symptoms do not differ appreciably from those of Trialeurodes vaporariorum, the glasshouse whitefly, which is common throughout Europe and also occurs elsewhere.
Similarities to Other Species/ConditionsTop of page
B. tabaci is now widely regarded to be a multi-species complex. Consisting of as many as 34 species that are morphologically indistinguishable from each other (De Barro et al., 2011; Boykin and De Barro, 2014). They can however, be distinguished molecularly. Differentiation of B. tabaci from other whitefly species by means of the adults is often difficult, although close observation of adult eye morphology may often show differences in ommatidial arrangements between some species. At rest, B. tabaci has wings more closely pressed to the body than Trialeurodes vaporariorum (greenhouse whitefly), which is a larger whitefly and more triangular in appearance.
The fourth instar or puparium can also be used to distinguish B. tabaci from T. vaporariorum as glasshouse pests. T. vaporariorum is 'pork-pie shaped', regularly ovoid, has straight sides (viewed laterally) and in most instances, 12 large, waxy setae. In B. tabaci, the puparium has an irregular, 'pancake-like', oval shape, with oblique sides and shorter, finer setae. Although the number of enlarged setae in B. tabaci and T. vaporariorum can vary according to host plant morphology, the two caudal setae are always stout and nearly always as long as the vasiform orifice in B. tabaci.
For more information on the identification of B. tabaci from slide-mounted pupae, see Martin (1987).
Prevention and ControlTop of page
Due to the variable regulations around (de)registration of pesticides, your national list of registered pesticides or relevant authority should be consulted to determine which products are legally allowed for use in your country when considering chemical control. Pesticides should always be used in a lawful manner, consistent with the product's label.
Intercropping practices using non-hosts have been used in many countries aiming to reduce numbers of whiteflies on specific crops. However, intercropping with susceptible crops can promote whitefly populations, by offering more leaf area for feeding.
Weed species play an important role in harbouring whiteflies between crop plantings and attention should be paid to removing these in advance of planting susceptible crops. Weeds also often harbour whitefly-transmitted viruses (Bedford et al., 1998) and may be a major source of crop virus epidemics.
Conservation of natural enemies is important in field crops where feeding damage is the cause of losses, rather than virus transmission, for example, on cotton. Under these circumstances, attempts have been made in Israel to enhance natural enemy action on cotton by introduction of additional, hopefully more efficient parasitoids (Rivany and Gerling, 1987). This effort resulted in the establishment of two species from the USA, Encarsia luteola and a species of Eretmocerus. Similarly, parasitoids are being introduced in Florida, USA, from other regions for the control of B. tabaci on vegetables and ornamentals (Rosen et al., 1994). Predatory mites have been shown to be efficient against Mediterranean species (Cuthbertson, 2014). Entomopathogenic agents such as nematodes (Cuthbertson et al., 2003a, 2007a,b) and fungi (Cuthbertson et al., 2005a, 2012; Cuthbertson and Walters, 2005) have also been shown to be important biological tools in the control/eradication of B. tabaci.
Plant and crop species that exhibit a high level of resistance to both vector and virus must also be considered when designing an IPM system. The development of transgenic resistant plant and crop species through genetic engineering must be considered and accepted as a future method of control where whitefly-transmitted viruses are already endemic and causing severe crop losses.
B. tabaci appears to develop resistance to all groups of pesticides that have been developed for its control. A rotation of insecticides that offer no cross-resistance must therefore be used to control B. tabaci infestations (Cuthbertson et al., 2012).
Active ingredients that have already been reported to have an effect in controlling B. tabaci worldwide include bifenthrin, buprofezin, imidacloprid, fenpropathrin, amitraz, fenoxycarb, deltamethrin, azidirachtin and pymetrozine. However, development of resistance to the products is a continual problem (Dennehy et al., 2010).
Integrated Pest Management
Until recently, B. tabaci was readily controlled with insecticides in field and glasshouse situations. However, problems with its effective control on many crops are now being experienced worldwide due to insecticide resistance. It appears that no single control treatment can be used on a long-term basis against this pest, and that the integration of a number of different control agents needs implementing for an effective level of control.
IPM appears to offer the best option for controlling B. tabaci without causing contamination of the environment: beneficial insects are used alongside chemicals that offer a high level of selectivity such as insect growth regulators. However, the use of biological control agents alone, such as Encarsia formosa and Lecanicillium lecanii, although moderately successful (Nedstam, 1992), can never bring infestation levels down to a level that stops virus transmission, as B. tabaci is such an efficient virus vector. Cuthbertson et al. (2012) developed a series of chemical control programmes, including Beauveria bassiana (Naturalis-L) that offered complete control of Mediterranean species under laboratory conditions. Nematodes and fungi have also been shown to be successfully tank-mixed with several chemical products for use in eradication schemes against what is now known to be MEAM1 species (Cuthbertson et al., 2003b, 2005b, 2012; Cuthbertson and Collins, 2015).
In countries where B. tabaci is not already present, the enforcement of strict phytosanitary regulations should help reduce the risk of this whitefly becoming established (Cuthbertson and Vänninen, 2015) . Because of the difficulty of detecting low levels of infestation in consignments, it is best to ensure that either the area or the place of production is free from the pest (OEPP/EPPO, 1990). If this cannot be obtained, a detailed treatment and inspection regime can be used to ensure that traded plants are free from the pest. Particular attention is needed for consignments from countries where certain B. tabaci-vectored viruses, now on the EPPO A1 or A2 quarantine lists, are present (see Risk of Introduction) (Cuthbertson and Vänninen, 2015).
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25/07/15 Reviewed by:
Andrew Cuthbertson, Food and Environment Research Agency, Sand Hutton, York, UK
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