Aonidomytilus albus (tapioca scale)

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Adult female

Aonidomytilus albus (tapioca scale); Adult female found on Papaya tree. Vieques, Puerto Rico. July 1972. Collected by Donald Miller.

©Alessandra Rung, Scale Insects, USDA APHIS PPQ/via Bugwood.org - CC BY-NC 3.0

Identity

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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

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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

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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

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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

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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

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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

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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.

Last updated: 23 Apr 2020

Continent/Country/Region	Distribution	Origin	Reference	Notes
Africa
Angola	Present	Introduced	NHM (1978) UK, CAB International (1978) Nakahara (1982) EPPO (2020)
Cabo Verde	Present	Introduced	NHM (1982)
Congo, Democratic Republic of the	Present	Introduced	Nakahara (1982)
Côte d'Ivoire	Present	Introduced	NHM (1942) UK, CAB International (1978) Nakahara (1982) EPPO (2020)
Gambia	Present	Introduced	Nakahara (1982)
Ghana	Present	Introduced	NHM (1937) UK, CAB International (1978) Nakahara (1982) EPPO (2020)
Guinea-Bissau	Present	Introduced	NHM (1931a)
Kenya	Present	Introduced	Bruijn and Guthrie (1982) Nakahara (1982) NHM (1985) EPPO (2020)
Liberia	Present	Introduced	Nakahara (1982)
Madagascar	Present	Introduced	UK, CAB International (1978) Nakahara (1982) Razafindrakoto et al. (1999) EPPO (2020)
Malawi	Present	Introduced	UK, CAB International (1978) Nakahara (1982) NHM (1987) EPPO (2020)
Mauritius	Present	Introduced	Williams and Williams (1988)
Mozambique	Present	Introduced	UK, CAB International (1978) Nakahara (1982) NHM (1982) EPPO (2020)
Nigeria	Present	Introduced	UK, CAB International (1978) Nakahara (1982) NHM (1983) EPPO (2020)
Senegal	Present	Introduced	NHM (1931) UK, CAB International (1978) Nakahara (1982) EPPO (2020)
Somalia	Present	Introduced	De Lotto (1967)
Tanzania	Present	Introduced	Bohlen (1973) UK, CAB International (1978) Nakahara (1982) EPPO (2020)
Uganda	Present	Introduced	NHM (1968) UK, CAB International (1978) Nakahara (1982) EPPO (2020)
Zambia	Present	Introduced	NHM (1984)
Asia
Bahrain	Present		EPPO (2020)
China	Present		CABI (Undated a)	Present based on regional distribution.
-Hainan	Present	Introduced	Tao (1999)
Hong Kong	Present	Introduced	Takagi (1970)
India	Present, Localized	Introduced	Sankaran et al. (1984) APPPC (1987) Williams and Williams (1988) EPPO (2020)
-Andhra Pradesh	Present	Introduced	NHM (1977)
-Karnataka	Present	Introduced	NHM (1982) Pillai et al. (1993)
-Kerala	Present	Introduced	Lal and Pillai (1981) CABI (Undated)
-Tamil Nadu	Present	Introduced	NHM (1954) Subramaniam et al. (1977) EPPO (2020) CABI (Undated)
Indonesia	Present	Introduced	Nakahara (1982)
Malaysia	Present		CABI (Undated a)	Present based on regional distribution.
-Peninsular Malaysia	Present	Introduced	Takahashi (1942)
Sri Lanka	Present	Introduced	Williams and Williams (1988) EPPO (2020)
Taiwan	Present	Introduced	Williams and Williams (1988) Tao (1999) EPPO (2020)
Thailand	Present	Introduced	APPPC (1987) Wongkobrat (1988) Waterhouse (1993)
North America
Antigua and Barbuda	Present	Native	Schotman (1989)
Bahamas	Present		Merrill (1953) Nakahara (1982)
Barbados	Present		UK, CAB International (1978) Bennett and Alam (1985) Schotman (1989) EPPO (2020)
British Virgin Islands	Present	Native	Schotman (1989)
Cuba	Present	Native	Ballou (1923) Merrill (1953) EPPO (2020)
Dominica	Present	Native	Schotman (1989)
Dominican Republic	Present	Native	UK, CAB International (1978) Schotman (1989) EPPO (2020)
Grenada	Present	Native	Schotman (1989)
Guadeloupe	Present	Native	Schotman (1989)
Haiti	Present	Native	UK, CAB International (1978) Schotman (1989) EPPO (2020)
Honduras	Present	Native	Merrill (1953) Nakahara (1982) EPPO (2020)
Jamaica	Present	Native	Merrill (1953) Kondo (2001) EPPO (2020)
Martinique	Present	Native	Schotman (1989)
Mexico	Present	Native	Cockerell (1899) UK, CAB International (1978) EPPO (2020) CABI (Undated)
Montserrat	Present	Native	Schotman (1989)
Puerto Rico	Present	Native	Schotman (1989)
Saint Kitts and Nevis	Present	Native	Schotman (1989)
Saint Lucia	Present	Native	Schotman (1989)
Saint Vincent and the Grenadines	Present	Native	UK, CAB International (1978) Schotman (1989) EPPO (2020)
Trinidad and Tobago	Present	Native	Schotman (1989)
U.S. Virgin Islands	Present	Native	Nakahara (1983)
United States	Present, Localized		Williams and Williams (1988) EPPO (2020)
-Florida	Present		Nakahara (1982) Miller (1996) EPPO (2020)
-New Mexico	Present		Merrill (1953) EPPO (2020)
South America
Argentina	Present	Native	UK, CAB International (1978) Nakahara (1982) Claps et al. (2001) EPPO (2020)
Brazil	Present	Native	Nakahara (1982) Kondo (2001) EPPO (2020)
-Amazonas	Present	Native	Claps et al. (2001)
-Bahia	Present	Native	Silva et al. (1968) Claps et al. (2001)
-Ceara	Present	Native	Bastos et al. (1979)
-Paraiba	Present	Native	Silva et al. (1968) Claps et al. (2001)
-Rio Grande do Sul	Present	Native	Silva et al. (1968) Claps et al. (2001)
Colombia	Present	Native	Mosquera (1976) Lozano et al. (1977) Vargas et al. (1978) Kondo (2001)
French Guiana	Present	Native	NHM (1978) Schotman (1989)
Guyana	Present	Native	NHM (1979) Schotman (1989)
Peru	Present	Native	Nakahara (1982)
Suriname	Present	Native	NHM (1977) UK, CAB International (1978) Nakahara (1982) EPPO (2020)

History of Introduction and Spread

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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

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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

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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

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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 name	Family	Context
Atriplex (orach)	Chenopodiaceae	Other
Carica papaya (pawpaw)	Caricaceae	Other
Chrysanthemum (daisy)	Asteraceae	Other
Croton bonplandianus	Euphorbiaceae	Other
Flourensia	Asteraceae	Other
Harrisia	Cactaceae	Other
Jatropha gossypiifolia (bellyache bush)	Euphorbiaceae	Other
Malvastrum americanum (spiked malvastrum (Australia))		Other
Mangifera indica (mango)	Anacardiaceae	Other
Manihot	Euphorbiaceae	Main
Manihot esculenta (cassava)	Euphorbiaceae	Main
Mimosa (sensitive plants)	Fabaceae	Other
Rosa (roses)	Rosaceae	Other
Salvia (sage)	Lamiaceae	Other
Sechium	Cucurbitaceae	Other
Sida	Malvaceae	Other
Solanum (nightshade)	Solanaceae	Other
Suaeda (sea blite)	Chenopodiaceae	Other
Turnera ulmifolia (West Indian holly)	Turneraceae	Other

Growth Stages

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Fruiting stage, Post-harvest, Vegetative growing stage

Symptoms

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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

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Sign	Life Stages	Type
Leaves / abnormal colours
Leaves / abnormal leaf fall
Leaves / external feeding
Leaves / wilting
Stems / external feeding

Biology and Ecology

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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

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Natural enemy	Type	Life stages	Biological control in	Biological 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

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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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Vector	Notes	Long Distance	Local	References
Land vehicles	Vehicles that have recently carried infested cassava.	Yes

Plant Trade

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Plant parts liable to carry the pest in trade/transport	Pest stages	Borne internally	Borne externally	Visibility 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

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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

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Category	Impact
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

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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

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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

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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

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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

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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

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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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CABI, Undated. Compendium record. Wallingford, UK: CABI

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Merrill G B, 1953. [English title not available]. Bulletin of the State Plant Board, Florida. 53.

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

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

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

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

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

NHM, 1942. Specimen record from the collection in the Natural History Museum (London, UK)., London, UK: Natural History Museum (London).

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).

NHM, 1978. Specimen record from the collection in the Natural History Museum (London, UK)., London, UK: Natural History Museum (London).

NHM, 1979. Specimen record from the collection in the Natural History Museum (London, UK)., London, UK: Natural History Museum (London).

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

NHM, 1983. Specimen record from the collection in the Natural History Museum (London, UK)., London, UK: Natural History Museum (London).

NHM, 1984. Specimen record from the collection in the Natural History Museum (London, UK)., London, UK: Natural History Museum (London).

NHM, 1985. Specimen record from the collection in the Natural History Museum (London, UK)., London, UK: Natural History Museum (London).

NHM, 1987. Specimen record from the collection in the Natural History Museum (London, UK)., London, UK: Natural History Museum (London).

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Razafindrakoto C, Ponte J J da, Andrade N C de, Silveira Filho J, Pimentel-Gomes F, 1999. Manipueira and heat treatment for the treatment of cassava cuttings attacked by scale insects. (Manipueira e termoterapia no tratamento de estacas de mandioca atacadas por cochonilhas.). Revista de Agricultura (Piracicaba). 74 (2), 127-136.

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 C Y L, 1989. Plant pests of quarantine importance to the Caribbean. In: RLAC-PROVEG, 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)., 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.

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. In: Insecta Matsumurana, 33 (1) 146 pp.

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). DOI:10.1079/DMPP/20056600081

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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Aonidomytilus albus (tapioca scale)

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