Invasive Species Compendium

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Datasheet

Aonidomytilus albus
(tapioca scale)

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Datasheet

Aonidomytilus albus (tapioca scale)

Summary

  • Last modified
  • 22 November 2019
  • Datasheet Type(s)
  • Invasive Species
  • Pest
  • Preferred Scientific Name
  • Aonidomytilus albus
  • Preferred Common Name
  • tapioca scale
  • Taxonomic Tree
  • Domain: Eukaryota
  •   Kingdom: Metazoa
  •     Phylum: Arthropoda
  •       Subphylum: Uniramia
  •         Class: Insecta
  • Summary of Invasiveness
  • The main infective stage of A. albus is the first-instar crawler, which is quite short-lived in the absence of a suitable feeding site. Within the field, infestation spreads outwards from infested cuttings, especially down wind, if the planting is no...
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    Compendia
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    OX10 8DE
    UK
    compend@cabi.org
  • Distribution map More information

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Identity

Top of page

Preferred Scientific Name

  • Aonidomytilus albus (Cockerell, 1893)

Preferred Common Name

  • tapioca scale

Other Scientific Names

  • Coccomytilus dispar (Vayssière) Takahashi, 1935
  • Lepidosaphes alba (Cockerell) Fernald, 1903
  • Lepidosaphes cockerelliana Kirkaldy, 1904
  • Lepidosaphes dispar
  • Mytilaspis (Coccomytilus) dispar Vayssière, 1914
  • Mytilaspis albus Cockerell, 1893
  • Mytilococcus dispar (Vayssière) Lindinger, 1943

International Common Names

  • English: cassava scale; cassava stem mussel scale; white mussel scale

Local Common Names

  • Colombia: escama de yuca

EPPO code

  • AONMAL (Aonidomytilus albus)

Summary of Invasiveness

Top of page The main infective stage of A. albus is the first-instar crawler, which is quite short-lived in the absence of a suitable feeding site. Within the field, infestation spreads outwards from infested cuttings, especially down wind, if the planting is not sufficiently widely spaced. However, without human or animal assistance, the capacity of this species to spread over longer distances is limited.

Taxonomic Tree

Top of page
  • Domain: Eukaryota
  •     Kingdom: Metazoa
  •         Phylum: Arthropoda
  •             Subphylum: Uniramia
  •                 Class: Insecta
  •                     Order: Hemiptera
  •                         Suborder: Sternorrhyncha
  •                             Unknown: Coccoidea
  •                                 Family: Diaspididae
  •                                     Genus: Aonidomytilus
  •                                         Species: Aonidomytilus albus

Notes on Taxonomy and Nomenclature

Top of page This species has been described three times under different names (albus Cockerell, 1893; cockerelliana Kirkaldy, 1904; and dispar Vaysière, 1914). These names have been placed in combination with several genera at various times.

Description

Top of page The scale cover of the adult female in life is elongate, mussel-shaped, 1.75-2.5 mm long, straight or curved, whitish to dark-brown, with slightly darker terminal exuviae. Watson (2002) provided colour illustrations of this species in life.

The scale cover of the second-instar male is similar to that of the adult female, but smaller (1.0-1.25 mm long) and narrower, with terminal exuviae (Ferris, 1941; Dekle, 1976). The non-feeding immature stages (pre-pupa and pupa) of the male develop beneath this scale cover.

After moulting to the adult stage, the male rests beneath the scale cover before emerging to seek for females. The adult male of A. albus has a single pair of simple wings, well-developed legs and antennae, and long genitalia.

The slide-mounted body of the adult female is elongate, and membranous except for the pygidium. The pygidium possesses several pairs of marginal lobes. The median lobes are not zygotic; they are well separated, with a pair of gland spines present between their bases, but without any basal scleroses. Perivulvar pores are absent; at least some gland spines are present on the pygidial margins, occurring in groups of two or three. The pygidium has six or seven enlarged marginal macroducts on each side, and the dorsal macroducts are scattered, not arranged in rows. Ferris (1941) and Watson (2002) provided taxonomic illustrations of the adult female of A. albus.

Distribution

Top of page A. albus is a tropical species of New World origin. It has not been recorded from Australia or the Pacific islands.

The distribution map includes records based on specimens of A. albus from the collection in the Natural History Museum (London, UK): dates of collection are noted in the List of countries (NHM, various dates).

Distribution Table

Top of page

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

Last updated: 23 Apr 2020
Continent/Country/Region Distribution Last Reported Origin First Reported Invasive Reference Notes

Africa

AngolaPresentIntroducedNHM (1978); UK, CAB International (1978); Nakahara (1982); EPPO (2020)
Cabo VerdePresentIntroducedNHM (1982)
Congo, Democratic Republic of thePresentIntroducedNakahara (1982)
Côte d'IvoirePresentIntroducedNHM (1942); UK, CAB International (1978); Nakahara (1982); EPPO (2020)
GambiaPresentIntroducedNakahara (1982)
GhanaPresentIntroducedNHM (1937); UK, CAB International (1978); Nakahara (1982); EPPO (2020)
Guinea-BissauPresentIntroducedNHM (1931a)
KenyaPresentIntroducedBruijn and Guthrie (1982); Nakahara (1982); NHM (1985); EPPO (2020)
LiberiaPresentIntroducedNakahara (1982)
MadagascarPresentIntroducedUK, CAB International (1978); Nakahara (1982); Razafindrakoto et al. (1999); EPPO (2020)
MalawiPresentIntroducedUK, CAB International (1978); Nakahara (1982); NHM (1987); EPPO (2020)
MauritiusPresentIntroducedWilliams and Williams (1988)
MozambiquePresentIntroducedUK, CAB International (1978); Nakahara (1982); NHM (1982); EPPO (2020)
NigeriaPresentIntroducedUK, CAB International (1978); Nakahara (1982); NHM (1983); EPPO (2020)
SenegalPresentIntroducedNHM (1931); UK, CAB International (1978); Nakahara (1982); EPPO (2020)
SomaliaPresentIntroducedDe Lotto (1967)
TanzaniaPresentIntroducedBohlen (1973); UK, CAB International (1978); Nakahara (1982); EPPO (2020)
UgandaPresentIntroducedNHM (1968); UK, CAB International (1978); Nakahara (1982); EPPO (2020)
ZambiaPresentIntroducedNHM (1984)

Asia

BahrainPresentEPPO (2020)
ChinaPresentCABI (Undated a)Present based on regional distribution.
-HainanPresentIntroducedTao (1999)
Hong KongPresentIntroducedTakagi (1970)
IndiaPresent, LocalizedIntroducedSankaran et al. (1984); APPPC (1987); Williams and Williams (1988); EPPO (2020)
-Andhra PradeshPresentIntroducedNHM (1977)
-KarnatakaPresentIntroducedNHM (1982); Pillai et al. (1993)
-KeralaPresentIntroducedLal and Pillai (1981); CABI (Undated)
-Tamil NaduPresentIntroducedNHM (1954); Subramaniam et al. (1977); EPPO (2020); CABI (Undated)
IndonesiaPresentIntroducedNakahara (1982)
MalaysiaPresentCABI (Undated a)Present based on regional distribution.
-Peninsular MalaysiaPresentIntroducedTakahashi (1942)
Sri LankaPresentIntroducedWilliams and Williams (1988); EPPO (2020)
TaiwanPresentIntroducedWilliams and Williams (1988); Tao (1999); EPPO (2020)
ThailandPresentIntroducedAPPPC (1987); Wongkobrat (1988); Waterhouse (1993)

North America

Antigua and BarbudaPresentNativeSchotman (1989)
BahamasPresentMerrill (1953); Nakahara (1982)
BarbadosPresentUK, CAB International (1978); Bennett and Alam (1985); Schotman (1989); EPPO (2020)
British Virgin IslandsPresentNativeSchotman (1989)
CubaPresentNativeBallou (1923); Merrill (1953); EPPO (2020)
DominicaPresentNativeSchotman (1989)
Dominican RepublicPresentNativeUK, CAB International (1978); Schotman (1989); EPPO (2020)
GrenadaPresentNativeSchotman (1989)
GuadeloupePresentNativeSchotman (1989)
HaitiPresentNativeUK, CAB International (1978); Schotman (1989); EPPO (2020)
HondurasPresentNativeMerrill (1953); Nakahara (1982); EPPO (2020)
JamaicaPresentNativeMerrill (1953); Kondo (2001); EPPO (2020)
MartiniquePresentNativeSchotman (1989)
MexicoPresentNativeCockerell (1899); UK, CAB International (1978); EPPO (2020); CABI (Undated)
MontserratPresentNativeSchotman (1989)
Puerto RicoPresentNativeSchotman (1989)
Saint Kitts and NevisPresentNativeSchotman (1989)
Saint LuciaPresentNativeSchotman (1989)
Saint Vincent and the GrenadinesPresentNativeUK, CAB International (1978); Schotman (1989); EPPO (2020)
Trinidad and TobagoPresentNativeSchotman (1989)
U.S. Virgin IslandsPresentNativeNakahara (1983)
United StatesPresent, LocalizedWilliams and Williams (1988); EPPO (2020)
-FloridaPresentNakahara (1982); Miller (1996); EPPO (2020)
-New MexicoPresentMerrill (1953); EPPO (2020)

South America

ArgentinaPresentNativeUK, CAB International (1978); Nakahara (1982); Claps et al. (2001); EPPO (2020)
BrazilPresentNativeNakahara (1982); Kondo (2001); EPPO (2020)
-AmazonasPresentNativeClaps et al. (2001)
-BahiaPresentNativeSilva et al. (1968); Claps et al. (2001)
-CearaPresentNativeBastos et al. (1979)
-ParaibaPresentNativeSilva et al. (1968); Claps et al. (2001)
-Rio Grande do SulPresentNativeSilva et al. (1968); Claps et al. (2001)
ColombiaPresentNativeMosquera (1976); Lozano et al. (1977); Vargas et al. (1978); Kondo (2001)
French GuianaPresentNativeNHM (1978); Schotman (1989)
GuyanaPresentNativeNHM (1979); Schotman (1989)
PeruPresentNativeNakahara (1982)
SurinamePresentNativeNHM (1977); UK, CAB International (1978); Nakahara (1982); EPPO (2020)

History of Introduction and Spread

Top of page A. albus is a tropical species of New World origin. There is no mention in the literature of the history of its spread, but it has undoubtedly reached countries outside the New World as a result of human transport of infested planting sticks of cassava.

Risk of Introduction

Top of page Transport of infested planting sticks of cassava, and stored cassava, is the main risk of transporting A. albus to new territory. Transport of infested material through fields planted with cassava also risks spread of the infestation, as crawlers may drop (or be blown) off the harvested material onto uninfested plants still in the field.

Habitat

Top of page A. albus feeds on the lower stems and, in heavy infestations, on the side shoots, petioles and ventral surfaces of the leaves.

Hosts/Species Affected

Top of page The preferred hosts of A. albus are species of Manihot, but this insect has been recorded feeding on a variety of hosts, including several species of Solanum.

Host Plants and Other Plants Affected

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Plant nameFamilyContext
Atriplex (orach)ChenopodiaceaeOther
Carica papaya (pawpaw)CaricaceaeOther
Chrysanthemum (daisy)AsteraceaeOther
Croton bonplandianusEuphorbiaceaeOther
FlourensiaAsteraceaeOther
HarrisiaCactaceaeOther
Jatropha gossypiifolia (bellyache bush)EuphorbiaceaeOther
Malvastrum americanum (spiked malvastrum (Australia))Other
Mangifera indica (mango)AnacardiaceaeOther
ManihotEuphorbiaceaeMain
Manihot esculenta (cassava)EuphorbiaceaeMain
Mimosa (sensitive plants)FabaceaeOther
Rosa (roses)RosaceaeOther
Salvia (sage)LamiaceaeOther
SechiumCucurbitaceaeOther
SidaMalvaceaeOther
Solanum (nightshade)SolanaceaeOther
Suaeda (sea blite)ChenopodiaceaeOther
Turnera ulmifolia (West Indian holly)TurneraceaeOther

Growth Stages

Top of page Fruiting stage, Post-harvest, Vegetative growing stage

Symptoms

Top of page On cassava, A. albus coats the stems, side shoots and even sometimes the leaf petioles and leaf undersides. Infestation in the field occurs in patches around a cutting that was infested at planting. Heavy infestation causes desiccation of the stems, making them become thin and weak so that they often break in the wind; death of the plant may result. The breakage of stems leads to profuse branching so infested plants often appear bushy. Root development in infested plants is poor, and the roots become unpalatable (Lal and Pillai, 1981).

List of Symptoms/Signs

Top of page
SignLife StagesType
Leaves / abnormal colours
Leaves / abnormal leaf fall
Leaves / external feeding
Leaves / wilting
Stems / external feeding

Biology and Ecology

Top of page Genetics

The genetics and karyotype of A. albus have not been studied.

Physiology and Phenology

Adult female A. albus feed throughout their lives and, once adult, live for several months. The adult male lacks mouthparts, so cannot feed and lives only a few days.

The eggs hatch in 3-4 days; in 20-25 days the immature stages are fully grown (Lal and Pillai, 1981). The first-instar crawlers are the primary dispersal stage and walk to new areas of the plant or are dispersed by wind or animal contact. Mortality due to abiotic factors is high in this stage. There are two immature instars in the female and four in the male (including non-feeding pre-pupal and pupal stages).

Reproductive Biology

Reproduction is sexual. The sessile females mate with winged males, and begin to lay eggs approximately 2 days after reaching maturity (Anantanarayanan et al., 1957).

Environmental Requirements

Dry conditions may make plants more susceptible to attack, and favour dispersal of the crawlers, which are vulnerable to drowning and being swept off the host in heavy rain and high winds.

Natural enemies

Top of page
Natural enemyTypeLife stagesSpecificityReferencesBiological control inBiological control on
Aphytis chrysomphali Parasite Eggs/Larvae/Nymphs
Aphytis diaspidis Parasite Eggs/Larvae/Nymphs
Aphytis lingnanensis Parasite Eggs/Larvae/Nymphs India Jatropha gossypiifolia
Aschersonia sp. Pathogen
Chilocorus distigma Predator
Cybocephalus Predator
Cybocephalus nitens Predator Cape Verde cassava
Encarsia aurantii Parasite
Nectria coccophila Pathogen
Pharoscymnus horni Predator
Pharoscymnus tomeensis Predator Cape Verde cassava

Means of Movement and Dispersal

Top of page Natural Dispersal

The first-instar crawlers are the dispersal stage and move across quite short distances to new parts of the host-plant or to adjacent plants. Dispersal over longer distances is only possible with the assistance of wind or animals/humans. Mortality due to abiotic factors is high during dispersal.

Movement in Trade

Dispersal of the sessile adults and immature stages between countries occurs through human transport of infested plant material, mainly on planting sticks rather than on stored tubers.

Pathway Vectors

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VectorNotesLong DistanceLocalReferences
Land vehiclesVehicles that have recently carried infested cassava. Yes

Plant Trade

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Plant parts liable to carry the pest in trade/transportPest stagesBorne internallyBorne externallyVisibility of pest or symptoms
Bulbs/Tubers/Corms/Rhizomes adults; eggs; larvae; nymphs; pupae Yes Pest or symptoms not visible to the naked eye but usually visible under light microscope
Stems (above ground)/Shoots/Trunks/Branches adults; eggs; larvae; nymphs; pupae Yes Pest or symptoms not visible to the naked eye but usually visible under light microscope
Plant parts not known to carry the pest in trade/transport
Bark
Flowers/Inflorescences/Cones/Calyx
Fruits (inc. pods)
Growing medium accompanying plants
Leaves
Roots
Seedlings/Micropropagated plants
True seeds (inc. grain)
Wood

Wood Packaging

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

Impact Summary

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

Impact

Top of page A. albus is only an occasional problem in the field; most often, it is a pest of cassava stems stored for later planting. Infested cuttings often do not root, and use of infested cuttings at planting can result in rooting failure of up to 80% (Lal and Pillai, 1981). Heavy infestation causes desiccation of the stems; in the field, this causes them to become thin and weak, and to break in the wind; death of the plant may result. Breakage of stems leads to profuse branching, and infested plants often appear bushy. The severity of attack becomes worse in drought conditions, aggravating drought stress (Lal and Pillai, 1981). The socio-economic impact of this can be considerable, as cassava is an important staple crop during drought, e.g. in Africa.

A. albus is a more or less serious pest of cassava in East and West Africa, Argentina, Brazil, Colombia, India, Madagascar, Mexico, Taiwan, Thailand, West Indies and USA (Florida) (Simmonds, 1960; Subramaniam et al., 1977; Anon., 1978; Vargas et al., 1978; Lal and Pillai, 1981; Wongkobrat, 1988). In Brazil, this species is a pest on Manihot and Solanum spp. (Foldi, 1988), and was regarded with potential pest status on Manihot spp. (source of Ceara rubber) by Bastos et al. (1979). It can cause serious damage locally in Kenya (Bruijn and Guthrie, 1982). Severe attacks on cassava cuttings kept for planting can lead to losses (Lal and Pillai, 1981; Chua and Wood, 1990); it is a field pest less often (Lal and Pillai, 1981).

Social Impact

Top of page The severity of attack by A. albus becomes worse in drought conditions, aggravating drought stress (Lal and Pillai, 1981). The socio-economic impact of this can be considerable, as cassava is an important staple crop during drought in Africa.

Diagnosis

Top of page Microscopic examination of slide-mounted adult females is required for authoritative identification to species. Ferris (1942, 1943) and Watson (2002) give identification keys that include A. albus.

Detection and Inspection

Top of page In the cassava field, look for bushy growth. In the field or in the cuttings store, examine stems of cassava closely (especially the bases) for pale to dark-brown, mussel-shaped scale covers.

Similarities to Other Species/Conditions

Top of page A. albus resembles Aonidomytilus solidaginis, but is readily recognisable when slide-mounted female specimens are examined at high power. In A. albus there are no perivulvar pores and the pygidial lobes are low; in A. solidaginis, perivulvar pores are present and the pygidial lobes are prominent (Ferris, 1941).

Prevention and Control

Top 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.

Phytosanitary Measures

Planting sticks of cassava, and stored cassava, should be thoroughly inspected for A. albus scales before export, as there is a risk of their dissemination on such material (Lozano et al., 1977). Imported planting material of cassava should also be thoroughly inspected before planting and treated if necessary, to kill any scale insects present.

Cultural Control and Sanitary Methods

Crowded planting encourages the development and spread of A. albus infestations; use of clean planting material, and well-spaced planting, reduces the risk of serious infestation. Spacing cuttings out avoids creation of a microclimate that favours the spread of any infestation. Use of infested cuttings should be avoided as they often do not root (Lal and Pillai, 1981).

The main infective stage of A. albus is the first-instar crawler, which is quite short-lived in the absence of food. Infected fields should therefore be completely cleared of cassava and left empty for at least 3 days before re-planting with cassava; crop rotation is also a valuable insurance against the carry-over of any infestation.

Transport of infested material through fields planted with cassava also risks transfer of the infestation, as crawlers may drop (or be blown) off the harvested material onto plants still in the field. If possible, it is better to transport the harvest by a route that avoids newly planted cassava fields.

Host-Plant Resistance

The cultivar 'Butter stick' has been reported to be highly susceptible to attack by A. albus (Lal and Pillai, 1981).

Biological Control

No mention of use of natural enemies against A. albus has been found in the literature.

Chemical Control

Lozano et al. (1977) recommended a 5-minute dip of planting sticks in 200 ppm malathion or diazinin to kill any infestation. Lal and Pillai (1981) found that vertical storage of stems reduced infestation, and spraying of infested stems with 0.1% malathion or methyl demeton before planting minimised subsequent infestation problems in the field. Pillai et al. (1993) recommended the use of dimethoate and methyl demeton for control of A. albus.

In Madagascar, it was found that a 60-minute immersion in liquid extract from cassava roots (manipueira) would kill A. albus before the cuttings were used for planting; this was more effective than immersion in hot water (Razafindrakoto et al., 1999).
 

References

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Anantanarayanan KP, Subramanian TR, Muthukrishnan TS, 1957. A note on the tapioca scale (Aonidomytilus albus Cockerell). Madras Agricultural Journal, 44(7):281-286.

APPPC, 1987. Insect pests of economic significance affecting major crops of the countries in Asia and the Pacific region. Technical Document No. 135. Bangkok, Thailand: Regional Office for Asia and the Pacific region (RAPA).

Ballou CH, 1923. Nota sobre coccidos Cubanos. Memorias de Sociedad Cubana de Historia Natural "Felipe Poey", 2/4:85-87.

Bastos JAM, Flechtman CHW, Figueiredo RWde, 1979. Contribution to knowledge of the pests of Ceara rubber. Fitossanidade, 3(1/2):45-46

Bennett FD, Alam MM, 1985. An annotated check-list of the insects and allied terrestrial arthropods of Barbados. Bridgetown, Barbados; Caribbean Agricultural Research & Development Institute, vi + 81 pp.

Bohlen E, 1973. Crop pests in Tanzania and their control. Berlin, Germany: Verlag Paul Parey.

Bruijn GH de, Guthrie EJ, 1982. Kenya. Root crops in eastern Africa. Proceedings of a workshop held at Kigali, Rwanda, 23-27 November 1980 International Development Research Centre Ottawa Canada, 95-98

Chua TH, Wood BJ, 1990. Other Tropical Fruit Trees and Shrubs. In: Rosen D, ed. Armoured Scale Insects, their Biology, Natural Enemies and Control. Vol. B. Amsterdam, Netherlands: Elsevier, 543-552.

Claps LE, Wolff VRS, González RH, 2001. Catálogo de las Diaspididae (Hemiptera: Coccoidea) exóticas de la Argentina, Brasil y Chile. Revista de la Socieded Entomológica Argentina, 60:9-34.

Cockerell TDA, 1893. The West Indian species of Mytilaspis and Pinnaspis. Entomologist’s Monthly Magazine, 29:155-158.

Cockerell TDA, 1899. Rhynchota, Hemiptera - Homoptera. [Aleurodidae and Coccidae]. Biologia Centrali Americana, 2:1-37.

Commonwealth Institute of Entomology, 1978. Distribution Maps of Pests, Series A (Agricultural), Map No. 81 (revised). Wallingford, UK: CAB International.

Dale PS, Maddison PA, 1984. Transport services as an aid to insect dispersal in the South Pacific. In: Laird M, ed. Commerce and the spread of pests and disease vectors. New York, USA: Prpger Publishers, 225-256

De Lotto G, 1967. A contribution to the knowledge of the African Coccoidea (Homoptera). Journal of the Entomological Society of southern Africa, 29:109-120.

Dekle GW, 1976. Florida armored scale insects. In: Arthropods of Florida and Neighboring Land Areas. Gainesville, Florida: Florida Department of Agriculture and Consumer Services, Division of Plant Industry, 3:1-345.

EPPO, 2014. PQR database. Paris, France: European and Mediterranean Plant Protection Organization. http://www.eppo.int/DATABASES/pqr/pqr.htm

Ferris GF, 1941. Atlas of the Scale Insects of North America. Series 3. Palo Alto, California: Stanford University Press.

Ferris GF, 1942. Atlas of the Scale Insects of North America. Series IV. The Diaspididae (Part IV). Stanford, California, USA: Stanford University Press.

Ferris GF, 1943. Additions to the knowledge of the Diaspididae (Homoptera: Coccoidea). (Contribution no. 41). Microentomology, 8:58-79.

Foldi I, 1988. New contribution to the study of scale insects from Brazilian Amazonia (Homoptera: Coccoidea). Annales de la Societe Entomologique de France, 24(1):77-87

Greathead DJ, 1990. Crawler behaviour and dispersal. 1.4.3. In: Rosen D, ed. Armoured Scales, their Biology, Natural Enemies and Control. World Crop Pests, Volume 4A. Amsterdam, The Netherlands: Elsevier, 305-308.

Kondo T, 2001. Las cochinillas de Colombia (Hemiptera: Coccoidea). Biota Colombiana, 2(1):31-48.

Lal SS, Pillai KS, 1981. Cassava pests and their control in southern India. Tropical Pest Management, 27(4):480-491

Lozano JC, Toro JC, Castro A, Bellotti AC, 1977. Production of cassava planting material. Series GE, CIAT, No.17:28pp.

Merrill GB, 1953. Bulletin of the State Plant Board, Florida, 1:53.

Miller DR, 1996. Checklist of the scale insects (Coccoidea: Homoptera) of Mexico. Proceedings of the Entomological Society of Washington, 98(1):68-86; 33 ref.

Mosquera PF, 1976. Escamas protegidas más frecuentes en Colombia. Boletín Técnico, Ministerio de Agrícola Instituto Colombiano Agropecuario, División de Sanidad Vegetal, 38:1-103.

Nakahara S, 1982. Checklist of the Armored Scales (Homoptera: Diapididae) of the Conterminous United States. Washington, USA: USDA, Animal and Plant Health Inspection Service, Plant Protection and Quarantine, 110 pp.

Nakahara S, 1983. List of the Coccoidea species (Homoptera) of the United States Virgin Islands. United States Department of Agriculture, Plant Protection and Quarantine, APHIS [Mimeograph], 8142:1-21.

Pillai KS, Palaniswami MS, Rajamma P, Mohandas C, Jayaprakas CA, 1993. Pest management in tuber crops. Indian Horticulture, 38(3):20-23

Rao YRVJ, Pillai KS, 1972. Effect of some insecticides on the scale insect, Aonidomytilus albus Ckll., infesting tapioca stems in storage. Indian Journal of Entomology, 34(2):181-182

Razafindrakoto C, Ponte JJda, Andrade NCde, Silveira Filho J, Pimentel-Gomes F, 1999. Manipueira and heat treatment for the treatment of cassava cuttings attacked by scale insects. Revista de Agricultura (Piracicaba), 74(2):127-136; 7 ref.

Sankaran T, Nagaraja H, Narasimham AU, 1984. On some South Indian armoured scales and their natural enemies. Proceedings of the 10th International Symposium of Central European Entomofaunistics, Budapest, 15-20 August 1983, 409-411.

Schotman CYL, 1989. Plant pests of quarantine importance to the Caribbean. RLAC-PROVEG, No. 21:80 pp.

Silva AGA, Gontalves CR, Galvpo DM, Gontalves AGL, Gomes J, Silva MN, Simoni L, 1968. Quarto catálogo dos insetos que vivem nas plantas do Brasil. Parte II, Tomo 1: Insetos, hospedeiros, inimigos naturais. Rio de Janeiro, Brazil: MinistTrio da Agricultura, 622 pp.

Simmonds FJ, 1960. Report on a tour of Commonwealth countries in Africa, March-June 1960. 1960. pp. 98. Commonwealth Agricultural Bureaux, Farnham Royal, Bucks., England.

Subramaniam TR, David BV, Thangavel P, Abraham EV, 1977. Insect pest problems of tuber crops in Tamil Nadu. Journal of Root Crops, 3(1):43-50.

Takagi S, 1970. Diaspididae of Taiwan based on material collected in connection with the Japan-U.S. Co-operative Science Programme, 1965. (Homoptera:Coccoidae). Part II. Insecta Matsumurana, 33(1):146 pp.

Takahashi R, 1942. Some Coccidae from Malaya and Hongkong (Homoptera). Transactions of the Formosa Natural History Society, 32:63-68.

Tao C, 1999. List of Coccoidea (Homoptera) of China. Taichung, Taiwan: Taiwan Agricultural Research Institute, Wufeng, 1-176.

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Williams JR, Williams DJ, 1988. Homoptera of the Mascarene Islands - an annotated catalogue. Entomology Memoir, Department of Agriculture and Water Supply, Republic of South Africa, No. 72, 98 pp.

Wongkobrat A, 1988. Insect pests of cassava in Thailand. Cassava Newsletter, 12(1):5-7

Distribution References

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NHM, 1931. Specimen record from the collection in the Natural History Museum (London, UK)., London, UK: Natural History Museum (London).

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NHM, 1937. Specimen record from the collection in the Natural History Museum (London, UK)., London, UK: Natural History Museum (London).

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NHM, 1954. Specimen record from the collection in the Natural History Museum (London, UK)., London, UK: Natural History Museum (London).

NHM, 1968. Specimen record from the collection in the Natural History Museum (London, UK)., London, UK: Natural History Museum (London).

NHM, 1977. Specimen record from the collection in the Natural History Museum (London, UK)., London, UK: Natural History Museum (London).

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NHM, 1982. Specimen record from the collection in the Natural History Museum (London, UK)., London, UK: Natural History Museum (London).

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Silva AGA, Gontalves CR, Galvpo DM, Gontalves AGL, Gomes J, Silva MN, Simoni L, 1968. (Quarto catálogo dos insetos que vivem nas plantas do Brasil. Parte II, Tomo 1: Insetos, hospedeiros, inimigos naturais)., 1 (II) Rio de Janeiro, Brazil: MinistTrio da Agricultura. 622 pp.

Subramaniam T R, David B V, Thangavel P, Abraham E V, 1977. Insect pest problems of tuber crops in Tamil Nadu. Journal of Root Crops. 3 (1), 43-50.

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Takahashi R, 1942. Some Coccidae from Malaya and Hongkong (Homoptera). Transactions of the Formosa Natural History Society. 63-68.

Tao C C, 1999. List of Coccoidea (Homoptera) of China. Wufeng, Taichung, Taiwan: Taiwan Agricultural Research Institute. 176 pp.

UK, CAB International, 1978. Aonidomytilus albus. [Distribution map]. In: Distribution Maps of Plant Pests, Wallingford, UK: CAB International. Map 81 (Revised).

Vargas HO, Brekelbaum T, Bellotti A, Lozano JC, 1978. The white scale (Aonidomytilus albus Ckll.) on cassava. [Proceedings cassava protection workshop CIAT, Cali, Colombia, 7-12 November, 1977], [ed. by Carlos Lozano J]. 199-202.

Waterhouse D F, 1993. The major arthropod pests and weeds of agriculture in Southeast Asia. Canberra, Australia: ACIAR. v + 141 pp.

Williams J R, Williams D J, 1988. Homoptera of the Mascarene Islands - an annotated catalogue. In: Entomology Memoir, Department of Agriculture and Water Supply, Republic of South Africa, iii + 98pp.

Wongkobrat A, 1988. Insect pests of cassava in Thailand. Cassava Newsletter. 12 (1), 5-7.

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