App 1 Guide to Scale Insect Families

Detection and identification of scale insects families (Hemiptera: Coccoidea)

Chris Malumphy

The Food and Environment Research Agency Department for Environment, Food and Rural Affairs Sand Hutton, York, UK YO41 1LZ

DETECTION AND IDENTIFICATION OF SCALE INSECTS

CONTENTS Page 1. Int roduction 3 1.1 Biology 3 1.2 Dispersal 4 1.3 Economic importance 4 2. Detection of scale insects 5 2. 1 Recognition of scale insect families in the field 8 3. Identification of scale insect families 10 3. 1 Preservation of specimens 10 3. 2 Adult female morphology 14 3. 3 Morph ological key to the scale insect families 14 4. Information sources 20 References 23

1. INTRODUCTION Scale insects are plant-sap feeding insects, closely related to the aphids, whiteflies and jumping plant lice or psyllids. They are among the most highly specialised of all plant parasites and feed on all parts of the plant including the roots, stems, leaves, buds and fruit. Some feed within hollow plant stems or plant galls; others mine beneath bark or live within plant tissue. There are about 7,500 species assigned to 1050 genera, in 28 or more families, in the superfamily Coccoidea. The higher classification is unresolved but here they are placed in the suborder Sternorrhyncha in the order Hemiptera. The purpose of this guide is to provide information that will assist workers in the United Kingdom Overseas Territories (UKOTs) to detect and identify scale insects to family level. This is intended to help develop diagnostic capacity within the UKOTs. The UKOTs are recognised as having a rich biodiversity that is under threat from the introduction of non- native species. Non-native species are a major cause of the loss of biodiversity globally, and island ecosystems typical of UKOTs are particularly vulnerable (Cheesman et al ., 2003; Varnham, 2006). A recent example of an accidental introduction of an invasive pest in a UKOT resulting in an environmental disaster has been the effect of pine tortoise scale (Toumeyella parvicornis ) on the pineyards in the Turks and Caicos Islands (Malumphy et al ., 2012). The scale has killed the majority of the mature pine trees changing the ecosystem. Accurate and rapid species identification for suspect non-native species is fundamental to the enforcement of eradication and quarantine measures to protect biodiversity and agriculture. 1.1 BIOLOGY

Neotenic, larviform adult female coccid Winged adult male scale insect ( Lichtensia (Lichtensia viburni ) viburni ). The caudal wax filaments assist the insect to glide in air currents

Scale insects show a tremendous diversity of biology. Most reproduce sexually, others parthenogenetically, others sexually and parthenogenetically, while a few are true hermaphrodites. The adults exhibit extreme sexual dimorphism. Adult females are larviform and neotenic (sexual mature in the larval stage), whereas adult males are winged (occasionally apterous) and resemble small flies. They have a single pair of forewings; the hind wings are reduced to short ribbon-like structures. They lack functional mouthparts, do not feed and live only from a few hours to a few days. The males of many species are attracted to the females by sex pheromones. Most scale insects lay eggs, some lay eggs that hatch almost immediately and some give birth to first instars. The eggs are protected in a variety of ways, some species enclose their eggs in an ovisac of waxen threads, some keep their eggs beneath the female's body, some beneath the separate scale-like covering of the adult female, some between wax plates secreted from the end of the abdomen and some

inside a ventral abdominal pouch. Female scale insects have two or three nymphal instars; the males have two nymphal instars followed by two non-feeding stages termed prepupa and pupa. 1.2 DISPERSAL

The first nymphal instar, known as the ‘crawler’, is the principal active dispersal stage and seeks out a new feeding site. First instars are also passively dispersed by wind, water, other insects, birds or by man. Subsequent nymphal instars and adult females of most scale insects are sedentary. The widespread dispersal of many species of scale insects has been largely caused by the movement of scale infested plant material around the world by man. In terms of number of species, scale insects are the largest superfamily of arthropods intercepted on imported plants and plant produce in England and Wales. They are also often the most numerous, in terms of the number of individuals, detected on imported plant material. 1.3 ECONOMIC IMPORTANCE

Severe hedge damage in Florida caused by Cycads killed by Aulacaspis yasumatsui in Maconellicoccus hirsutus © 2004-2007 Guam © Anne Brooke, Guam National Florida Department of Agriculture and Wildlife Refuge Consumer Services

Scale insects may attack any part of the plant. They are widely distributed throughout the world with the exception of the cold extremes of the Arctic and Antarctic. Host plant diversity is broad, although scales are not commonly found on ferns or mosses. They damage the plants directly by removing sap which reduces host vigour, and may cause chlorosis, discolouration, pitting of stems and fruit, leaf and shoot distortion, localised necrosis, premature leaf drop, drying out of the foliage and stems, die back, and even death of susceptible plants. They may also induce galls, inject toxic saliva and vector plant pathogenic diseases. Many species, particularly in the families Coccidae and Pseudococcidae, produce copious quantities of honeydew which smoothers the host and serves as a medium for the growth of black sooty moulds. The mould screens light from the leaves and restricts gas exchange, reducing photosynthesis and hence productivity. The market value of ornamental plants and plant produce is lowered by insect feeding damage and the presence of moulds. The small size and cryptic nature of scale insects means that they can easily escape detection during quarantine inspections and are regularly transported in international trade. The

accidental introduction of a single fertilized or parthenogenetic female may initiate a serious infestation, causing serious loss of yield or killing crops. Examples of scale insects introduced to new geographical areas and causing major economic losses include: Phenacoccus manihoti, Rastrococcus invadens, Maconellicoccus hirsutus and Aulacaspis yasumatsui . The cassava mealybug, Phenacoccus manihoti , was accidentally introduced to West Africa from South Amercia and was a serious threat to cassava, a staple crop, until a parasitoid wasp was introduced to control it. The ‘mango mealybug’, Rastrococcus invadens , originates from South East Asia and was inadvertently introduced into Ghana and Togo in 1981-82. It spread rapidly through West Africa where it became a serious economic pest of several crops, reducing mango yields by 50-90%. A parasitoid wasp was introduced into Togo in the late 1980s and has successfully controlled the mealybug in most areas. The pink hibiscus mealybug, Maconellicoccus hirsutus , is a serious pest of many crop and ornamental plants in tropical and subtropical regions, including Africa, Southeast Asia, and northern Australia. It was introduced in to the Caribbean in 1994 and has since spread to the USA and South America. It has caused major economic losses and increased crop production costs and is the subject of a biological control programme. The cycad aulacaspis scale, Aulacaspis yasumatsui , is a cycad pest native to South East Asia. It was introduced to the USA (including Hawaii), Central America, Caribbean, Singapore, Hong Kong, Taiwan, Guam and the Ivory Coast. Populations can build up rapidly and heavily infested plants are killed. For example, 90% of the native cycads in parts of Guam have been killed. Many species of scale insect, however, are beneficial to man. Several species are used to produce dyes for textiles, cosmetics and foods, for example, cochineal red dye is produced from Dactylopius coccus , crimson lake dye from Kermes vermilio , Polish cochineal from Porphyrophora polonica and Armenian cochineal from Prophyrophora hamelii . They have also been used for lacquers (shellac furniture polish from Laccifer lacca ), resins, waterproofing 'fats', medicines, cosmetics and food (Biblical 'manna' came from Trabutina mannipara ). Ericerus pela is cultivated on an enormous scale in China for wax to make candles. Hard-shelled margarodids, known as ground pearls, are used to make jewellery and ornaments. Honeydew produced by Marchalna hellenica on pine trees in Greece is collected by bees and used to make honey. Scale insects have also been used for the biological control of weeds, for example, Dactylopius spp. have been used as biological control agents for Opuntia cacti in Australia, Ceylon, India, Hawaii, and Mauritius. 2. DETECTION OF SCALE INSECTS

Scale insects feed on all parts of the plant, including the leaves, twigs, branches, fruit and roots. Some species are relatively large and highly conspicuous, for example Icerya purchasi , and others are conspicuous as they often occur in large populations. Many produce conspicuous white, waxy ovisacs, particularly some of the Coccidae and Pseudococcidae. Unfortunately, the majority of the scale insects encountered are usually inconspicuous, being small, cryptic and are normally present in low numbers. The presence of scale insects is often detected first by the symptoms caused by feeding damage. For example, chlorotic spotting or streaking on the foliage, uneven-ripening of fruit, galling on the stems and leaves, distortion of the growing tips, premature leaf drop, loss of vigour and stunting of the host plant. In addition, many species, particularly the Coccidae and Pseudococcidae, eliminate excess plant sap as honeydew, which may contaminate the plant. The honeydew serves as a substrate for the growth of black sooty moulds and also attracts ants, wasps and flies, which feed on the sugary substance. White waxy ovisacs, male wax tests or waxy deposits may also betray the presence of scale insects. However, it must

be borne in mind that other plant sap-sucking hemipteran insects, such as aphids and whiteflies, also produce identical symptoms. Some species of diaspid actually burrow beneath the epidermis and bark and are often almost impossible to detect with external examination (for example Andaspis hawaiiensis, Clavaspis herculeana and Howardia biclavis ). Their presence is only indicated by small swellings in the bark, which need to be carefully cut open to reveal the insects beneath. Many scale insects, particularly the pest species, vary considerably in form and colour depending on the host plant species, feeding site, maturity and whether they are parasitized or not. For example, they may be oval and convex on the foliage but elongate and flatter when feeding on stems; or brightly patterned when teneral or a uniform dull colour when mature. Scale insects also exhibit extreme sexual dimorphism. Most scale insects typically exhibit a strongly clumped distribution on the host plants. Therefore, as much plant material as practical should be examined in order to detect them. Scale insects should generally be collected still attached to the plant material, preserved in 70% ethanol, and submitted to the laboratory for identification. Removing scale insects from the host in the field usually damages the specimen making diagnosis more difficult.

Chlorotic spotting caused by Dynaspidiotus Chlorosis and yellow st reaks on a palm frond brittanicus on Ilex caused by Pseudaulacaspis sp.

Chlorotic spotting and streaking on a cycad Uneven ripening of Citrus sinensis fruit caused by Aspidiotus destructor caused by Parlatoria pergandii

Quercus ro bur twig, deeply pitted by The growing tip of Olea europea , exhibiting Asterodiaspis variolosa . The gall on the left is considerable distortion due to Aspidiotus empty, whereas the one on the right still nerii contains the adult female.

Honeydew e xcreted by Coccus hesperidum , Black sooty mould growing on honeydew glistening on the surface of a Citrus leaf excreted by Pulvinaria floccifera on Rhododendron

Ovisacs, eggs and nymphs of Pseudococcus Conspicuous waxy tests of male viburni on Rhapsalis Nipaecoccus nipae on Psidium

Teneral adult female Saissetia neglecta are Mature adult female Saissetia neglecta are reddish brown with distinct dorsal ridges, on black with no obvious dorsal ridges, on an croton unspecified plant

2.1 RECOGNITION OF SCALE INSECT FAMILIES IN THE FIELD

Twelve families are discussed here as they have either been collected in the UKOTs or are found moving in plant trade and may be introduced into the UKOTs in the future: Acleridae, Asterolecaniidae, Coccidae, Conchaspididae, Dactylopiidae, Diaspididae, Eriococcidae, Monophlebiidae, Ortheziidae, Phoenicococcidae, Pseudococcidae and Putoidae. Three families are significantly more speciose and commonly encountered than the other families: Coccidae, Diaspididae and Pseudococcidae.

Acleridae – flat grass scales The body is usually flattened, generally reddish-brown or pink, with the apex of the abdomen heavily sclerotized and dark brown; usually with small amount of white wax ventrally and on the head. Some species produce abundant wax dorsally. They are cryptic, hidden beneath the leaf sheaths or in the crown of plant near soil surface. Asterolecaniidae – pit scales This is a relatively diverse group, but a typical pit scale has the body of the insect set in a depression or pit in the host tissue and a white fringe around the body margin. There generally is a translucent yellow or green test that covers the insect. A few uncommon genera form complete galls.

Coccidae – soft scales This family is diagnosed by the presence of a pair of triangular anal plates. The appearance of soft scales in life is highly variable depending on the group, maturity and host plant. The body shape is frequently round or broadly oval but some are elongate; wax coverings are thin and transparent, filamentous, powdery, thick and opaque, or thin and glassy. Some species produce conspicuous white filamentous waxy ovisacs beneath or behind the female, or that completely envelops the adult.

Conchaspididae – false armoured scales The body is hidden under a thick wax cover, similar to a diaspid cover except that the exuviae are not incorporated. The cover is not attached to the body. The cover of most of the species is whitish and the female body is also usually white.

Dactylop iidae – cochineal scales The body of the adult female is greyish but bright red when crushed; they are often covered by sticky, web-like strands of filamentous wax. They occur on the pads of cacti in clumps in protected areas.

Diaspididae – armoured s cales The body is covered by a separate wax covering with the exuviae of 1 or 2 immature instars incorporated. The body may be elongate or oval; white, yellow, purple, red, or orange. Armoured scales occur on nearly any part of plant, including the roots; some species become buried under plant epidermis. Eriococcidae – felt scales Felt scales produce a white, grey, or yellowish ovisac that encloses the pyriform body of the adult female. Body colour varies from pink or red to purple, green or brown. The posterior end of the sac has a small opening that allows the first instars to escape. Some eriococcids occur under the bark of the host, produce little or no ovisac secretion and are often pink or red. Many species produce plant galls. Monop hlebidae – giant scale insects The body is elongate, oval, large, up to 10 mm or more; legs and antennae conspicuous and dark; usually with wax covering the body, occasionally without wax; often forming an ovisac or with a marsupium. This family is considered part of the Margarodidae by some workers.

Ortheziidae – ensign scales These are among the most easily recognisable of all scale insects in life due to their uniform appearance. The body is adorned with thick wax plates, which may be marginal or cover the body, and the ovisac is attached to the abdomen, rather than the host; the legs and antennae are large and dark.

Pheonicococcidae – palm scales The body is small, about 1.5 mm long, spherical, bright red, embedded or nested in white wax; often occurring on the white tissues at the base of fronds, occasionally on exposed roots and fronds.

Pseudococcidae – mealybugs The adult females are often characterized by a white, mealy or powdery secretion that covers the body. The marginal areas of the body frequently bear a series of protruding lateral wax filaments or tufts, which may occur around the entire body or be confined to the posterior abdominal segments. A filamentous secretion is often produced that encloses the eggs and at least part of the body. Putoidae – giant mealybugs The body is large, 5 mm or longer; covered with thick tufts of mealy white wax; lateral filaments broad and often coalesce; often with a central ridge of wax; with 2 black stripes on dorsal submedial areas when the wax is removed; legs and antennae are large and dark; no definite ovisac is produced in most species. This family is considered part of the Pseudococcidae by some workers.

3. IDENTIFICATION OF SCALE INSECT FAMILIES

For accurate identification scale insects need to be examined microscopically and are mounted on glass microscope slides. In most cases only teneral (newly moulted) adult females are identifiable and this is the stage referred to in this procedure. Before mounting on microscope slides, the specimens are macerated, dewaxed, dehydrated, stained and cleared. Maceration dissolves the body contents, leaving only the exoskeleton. This requires staining to show the microscopic cuticular structures on which the characters for identification are based. Specimens required for future reference are prepared using the permanent method and mounted in Canada balsam (currently regarded as the most stable permanent mounting medium). Other specimens that need to be processed rapidly are prepared using the temporary method and mounted in Heinz mountant. The latter method saves time and reagents but is unsuitable for very waxy specimens. 3.1 PRESERVATION OF SPECIMENS

The preparation technique given below is modified from Martin (1987). The procedure is not rigid and can be readily modified to suit particular samples. Specimens are manipulated and mounted on microscope slides with the aid of a binocular dissecting microscope. Care is necessary to keep watch glasses and glass slides accurately labelled with the reference number throughout the procedure. The glassware must be inscribed with a waterproof marker. The minimum time required for the preparation technique is just over an hour. The temporary preparation technique requires a minimum of 20 minutes. The method requires a

high level of manual dexterity, an understanding of what the reagents are doing and when a specimen is ready for the next treatment. To produce consistently good preparations requires considerable experience and is a highly specialised skill. SCOPE This microscope slide preparation method is suitable for all Coccoidea whether live, dead or preserved specimens. SAFETY Many of the reagents used in this preparation are potentially dangerous and need to be treated with caution (see Table 1). The health and safety legislation regarding the use of these reagents may vary between countries and should be checked before attempting to use this method. In addition the hazards ratings for the reagents are constantly revised and may change from those listed below. The watch glasses containing specimens and solutions are heated, where necessary, on a dry heating block. The entire procedure is carried out in a fume cupboard.

Table 1 List of reagents used for making slide-preparations and their hazards Reagent Hazard Acid fuchsin Carcinogen. Toxic and Corrosive. Stains. Canada balsam Irritating to eyes and skin. Flammable. Vapour is narcotic in high concentration. Chloral hydrate Toxic by ingestion. Irritating to skin and eyes. Harmful vapour. Clove Oil May cause irritation to skin and eyes. D-(+)-Glucose May cause irritation to skin and eyes. Ethanol Causes severe eye irritation and moderate skin irritation. Highly flammable. Glacial acetic acid Harmful by skin contact and ingestion. Causes severe burns. Flammable, corrosive. Lactic acid Corrosive. Causes severe burns to eyes and skin. Phenol Very toxic by skin contact ingestion and inhalation. Causes burns. Wash skin immediately with plenty of polyethylene glycol 300 and seek medical advice. Polyvinyl alcohol Carcinogen. Causes irritation by inhalation and ingestion. Also Irritating to the eyes. Potassium hydroxide Causes severe irritation and burns to eyes and skin. If ingested causes severe internal irritation and damage. Xylene Toxic by skin contact and ingestion. Causes burns. Harmful if inhaled. Wash skin with plenty of water and seek medical advice. Highly flammable.

MATERIALS Potassium hydroxide (KOH) 10% Glacial acetic acid Chloral phenol (recipe is given in the Appendix) Acid fuchsin stain (recipe is given in the Appendix)

Ethanol 70% Clove oil Canada balsam Heinz mountant (recipe is given in the Appendix) Xylene Dry heating block Safelabs Systems Arone 1000 GS Safety cabinet. AG/0131 Leica Mz Microscope Light Source Schott KL 1500 LCD Watch glasses Glass squares Spatulas Glass Pasteur pipettes with teats Glass Jar with lid for waste solutions Needles and mounts Forceps Glass microscope slides Lens tissue Tissue paper Glass cover slips 13 mm-19 mm Bristol board Indian ink Mapping pen Scissors Permanent marker Glue

PROCEDURE FOR PERMANENT PREPARATION

1. Place specimens into 70% ethanol in a watch glass, cover with a glass square and heat gently to simmering point for a few minutes. Fixation in hot alcohol makes specimens less fragile, so they lose fewer setae during mounting. Pipette off the alcohol using a fine glass teat pipette, taking care not to accidentally suck up the specimens. 2. Add approximately 1ml of 10% potassium hydroxide (KOH) and heat on Dry block set at 70 °C to simmering point for approximately 5-10 minutes, or until the specimens lose most of their body colour. Make a small dorsal incision above and between the antennal bases to help the process of maceration. The length of time required varies considerable (depending on the species, body size, and thickness of wax, how long the specimens have been preserved in alcohol, the particular instar and their maturity). Diaspididae take less time than Pseudococcidae and Coccidae. Early instars require less time than adults. Large, heavily sclerotized adult females take much longer. An alternative procedure is to leave mature specimens of Pseudococcidae or Coccidae in cold KOH for about 24 hours; the latter method is also better for delicate species. 3. Examine the specimens under a binocular microscope. Where necessary, tease away the wax or scale covering from the specimens using fine needles. With Diaspididae, the body of the insect has to be separated from the protein and wax covering. With 'pupillarial' Diaspididae, the adult is encased within the second instar exuvia (cast skin) and has to be dissected out. Expel the liquefied body contents through the dorsal incision using two fine spatulas. With Pseudococcidae and Coccidae, the main tracheal branches are teased from the spiracles and removed through the dorsal incision, using

DETECTION OF SCALE INSECT FAMILIES

mounted micropins with curved tips. Parasitoid larvae and pupal cases and fungal hyphae are also removed. Parasitoid larvae are retained with the host specimens. Pipette off the excess KOH. 4. Soak the specimens in about 2 mls of 70% ethanol for a minimum of 10 minutes. This rinses out the remaining KOH. Pipette off the liquid. 5. Rinse the specimens in about 2 mls cold glacial acetic acid, which is then pipetted off. 6. Add a few drops of choral phenol, a wax solvent, to the watch glass. Gently heat for 10- 15 minutes on Dry block set at 80 °c, depending upon how waxy the specimens are. Waxier specimens require longer. Females with ovisacs are often associated with large quantities of wax. In such cases, the choral phenol has to be changed one or more times, until all the wax has dissolved. Pipette off the choral-phenol. In cases where the specimens are exceptionally small and easily lost they may be stained (stage 7) prior to adding chloral phenol and stained again afterwards. 7. Add fresh glacial acetic acid, together with a drop of acid fuchsin stain. Agitate the watch glass until the acid is a uniform pink colour. Leave the stain for about twenty seconds until the specimens acquire a pink tinge. Pipette off the liquid. If the specimen is over-stained, it is soaked in 70 % ethanol until the excess stain dissolves. It is then returned to glacial acetic acid. 8. Rinse the specimens with fresh glacial acetic acid to remove any remaining stain. Pipette off the liquid. 9 Add fresh glacial acetic acid to the specimens and leave for at least 5 minutes to completely dehydrate. This is again pipetted off. 10. Add a few drops of clove oil, enough to allow the specimens to float freely, and leave for at least 10 minutes while the specimens clear. Remove any remaining body contents in the same way as described in stage 3 above. Care has to be taken, however, not to accidentally remove setae after the specimens have been dehydrated. 11. Using a fine spatula transfer a single specimen with its dorsal surface upwards onto a clean glass slide. Parasitoid larvae are usually mounted with their host. Always mount the specimens separately unless it is absolutely certain that they are the same species, in which case, mount up to six individuals on the same slide. Space the specimens evenly on the slide to avoid the coverslip tilting when mounted. Spread out each scale insect body and arrange the appendages. Absorb excess clove oil with the rolled corner of a tissue. Take care not to leave fibres from the tissue on the slide. Carefully apply a drop of dilute Canada balsam to the specimens on the slide. Rest one edge of a 13mm or 19 mm diameter coverslip on the slide holding the opposite edge with a needle. Gently lower the coverslip with the needle onto the droplet of balsam covering the specimen. Take care to ensure that air is excluded and that the meniscus spreads outwards to the edge of the coverslip. Allow the coverslip to settle under its own weight. 12. Label (all essential data) using Bristol board squares before placing in the collection to dry. Drying can take two months or more to complete. When dry scrape off excess balsam that has spread out from beneath the coverslip using a razor blade.

DETECTION OF SCALE INSECT FAMILIES

PROCEDURE FOR TEMPORARY PREPARATION

The temporary preparation method is similar to the permanent preparation method but the specimens are not dewaxed and are mounted in Heinz mountant. This method is only suitable for non-waxy specimens. Only an outline of the method is given, as the details are the same as those above. 1. Simmer specimens in 10% KOH for approximately 5-10 minutes, or until the specimens lose most of their body colour. 2. Examine the specimens under the microscope. Expel the body contents by making a small dorsal incision above and between the antennal bases, and pumping the liquefied body contents out, using two fine spatulas. Pipette off the excess KOH. 3. Soak the specimens in cold 70% ethanol for two minutes. Pipette off the liquid. 4. Mount the specimens in Heinz mountant on a glass microscope slide.

3.2 ADULT FEMALE MORPHOLOGY

Illustrations showing the main morphological characters of the three main families of Coccoidea are presented below. There are many easily available texts, which describe the morphology in detail. For the Pseudococcidae see McKenzie (1967), Williams (2004), Williams & Watson (1988a) and Williams & Granara de Willink (1992). For the Coccidae see Gill (1988), Hamon & Williams (1984) and Hodgson (1994). For the Diaspididae see Ferris (1937) and Williams & Watson (1988). The following site includes excellent morphological illustrations of all the families: http://www.sel.barc.usda.gov/scalekeys/scalefamilies/index.html

3.3 MORPHOLOGICAL KEY TO SCALE INSECT FAMILIES

Scale insects are characterized by having a single claw, neotenic adult females, winged and non-feeding adult males, and an unusual form of metamorphosis that normally includes a prepupa and pupa in the adult male. Generally there are three or four instars in the female and five instars in the male. Classification of the Coccoidea is based mainly on the morphology of the adult female. There is disagreement on the number of families within the Coccoidea and the system used here follows ScaleNet (2015). A key is provided to 20 families, including all those that have been encountered in the UKOTs. The Margarodidae and closely related families key out as the ‘Margarodidae Group’.

DETECTION OF SCALE INSECT FAMILIES

Key to families of scale insect based on adult female morphology

1. Abdominal spiracles present, numbering 2 -8 pairs ...... 2

Icerya purchasi abdominal spiracle - Abdominal spiracles absent …… ...... 3 2. Antennae each with 3 -8 segments, the terminal segment with a stout apical spine-like seta. Anal ring present with 6 setae and a band of pores. Eyes often stalked. Ovisac band of spines present ..... ORTHEZIIDAE

Nipponorthezia ardisiae antennae - Antennae without an apical spine, varying from flat plates to segmented. Up to 11 segments present. Anal opening simple, never surrounded by a ring of pores and setae, but sometimes an internal tube present with the inner end surrounded by a band of pores. Anus often distinctly dorsal in position. Ovisac band, if present, composed of pores...... Icerya seychellarum MARGARODIDAE GROUP (including MONOPHLEBIDAE) antennae 3. Anterior thoracic spiracles conspicuously larger than posterior pair. Brachia and brachial plates present on thorax. Dorsal spine present near centre of abdomen. Posterior of abdomen prolonged into an anal tubercle, bearing an anal ring with 10 setae surrounded by a ring of fimbriations, lobes or setae ...... KERRIIDAE Tachardiella fulgens © ScaleNet - Anterior and poster ior thoracic spiracles of similar size. Brachia, brachial plates and dorsal spine absent. Anal opening external or at the base of a small tube, never with an anal fringe at the apex of an anal tubercle bearing fimbriations, lobes or setae ...... 4 4. Anal cleft present, with a pair of dorsal triangular or rounded anal plates situated at its base; inner margins of the plates usually contiguous, forming an operculum ...... COCCIDAE

Saissetia coffeae anal plates

DETECTION OF SCALE INSECT FAMILIES

- Anal cleft, if present, without paired plates at its base, although there may be a single plate in this position ...... 5 5. Furrows and ridges present on posterior margin. Short anal cleft present, with a single triangular or oval anal plate situated at its base. Anus leading to a telescoping anal tube with an anal ring bearing about 10 setae at its inner end ...... ACLERDIDAE

Aclerda berlesii posterior margin - Posterior margin lacking furrows and ridges. Without a single, oval anal plate, although a small or divided plate may be present at base of anal lobes ………………….....… 6 6. Anus oval, heavily sclerotised, situated ne ar middle of dorsum ...... STICTOCOCCIDAE

Stictococcus sjostedti anal plates - Anal opening at posterior of body ...... 7 7. 8-shaped pores present somewhere on body ...... 8

Asterolecanium pustulans 8-shaped pores - 8-shaped pores absent, although tubular ducts, each with single opening and the duct 8-shaped in cross section, may be present ...... 11 8. 8-shaped pores present on dorsum only. Ventral tubular ducts present in a submarginal band around body. Antennae each with 5 segments. On Quercus and occasionally closely related plants …………………………...... KERMESIDAE (in part) - 8-shape d pores present on dorsum and on venter in a submarginal zone. Ventral tubular ducts scattered. Antennae each with 1-9 segments. On various host plants ...... 9

DETECTION OF SCALE INSECT FAMILIES

9. Anal plate well deve loped, triangular or butterfly - shaped; cribriform plates present at least on dorsum of abdomen; tubular ducts present, each with a large terminal filament ...... 10

Cribriform plates © ScaleNet - Anal pla te poorly developed or absent; cribriform plates absent; tubular ducts each without a terminal filament ...... ASTEROLECANIIDAE 10. Antennae each 1 -segmented, with a group of quinquelocular pores present near the base. Anal plate triangular and shield-like. Venter of abdomen with transverse rows of 8-shaped pores present ……………...... CEROCOCCIDAE - Antennae each with 7 -9 segments, without quinquelocular pores present near base. Anal plate butterfly-shaped, usually formed of 2 triangular plates linked by a narrow yoke; a small, arched plate also present, situated anterior to anal ring. Venter without transverse rows of 8-shaped pores ……………………………...... LECANODIASPIDIDAE 11. Dorsum with scattered clusters of 3 -, 4 - or 5 -locular pores, each cluster with a single common duct. Thick truncate setae present, each with the sides almost parallel ...... DACTYLOPIIDAE

Dactylopius coccus - Dorsum without single -ducted clusters of 3 -, 4 - or 5 - locular pores. If truncate setae are present, their sides are conical ...... 12 12. Abdomen with posterior segments fused into a pygidium or pygidium-like area ...... 13

Chrysomphalus aonidum pygidium - Abdomen with posterior segments not fused into a pygidium ...... 15

DETECTION OF SCALE INSECT FAMILIES

13. Legs and antennae usually present. Scale covering not incorporating exuviae of previous instars. Multilocular disc pores often ‘flower-‘ or star-shaped ……………………… ...... CONCHASPIDIDAE

Conchaspis angraeci multilocular pores - Legs and antennae reduced or absent. Scale covering, if present, incorporating exuviae of previous instars. Multilocular disc pores absent; disc pores, if present, each with a maximum of 5 loculi ...... 14 14. Pygidium with dorsal ducts and a marginal fringe of plates or gland-spines and lobes. Usually with a scale cover incorporating the exuviae of previous instars but sometimes adult female remains inside the exuviae of previous stage (pupillarial) ...... DIASPIDIDAE

Parlatoria © ScaleNet - Pygidium simple, without plates, gland -spines or lobes. Adult female pupillarial, always remaining inside the exuviae of second instar, the posterior end of which forms a flat anal plate or operculum surrounded by a sclerotised rim ...... HALIMOCOCCIDAE

Colobopyga australiensis © ScaleNet 15. Adult female pupillarial (enclosed in exuviae of previous stage). Thoracic spiracles displaced towards posterior end of abdomen. Antennae and legs absent ……………… ...... BEESONIIDAE

DETECTION OF SCALE INSECT FAMILIES

16. Anal ring simple, with 0 -2 setae and no pores. Tubular ducts 8-shaped in cross-section ... PHOENICOCOCCIDAE

Phoenicococcus marlatti simple anal ring - Anal ring variable. Tubular ducts never 8 -shaped in cross-section ...... 17 17. With a combination of dorsal ostioles, ventral circuli, triocular pores and usually 1-18 pairs of cerarii. Anal ring usually with outer and inner rows of pores. Tubular ducts not cupped at inner end ……………………………….. 18 - Without dorsal ostioles, ventral circuli, triocular pores and cerarii. Anal ring variable. Macrotubular tubular ducts each with inner end invaginated to form a cup. Microtubular ducts present or absent ...... …. 19 18. Trochanter with 3 -4 pores ...... PUTOIDAE

Puto sp. trochanter with 3 pores - Trochanter with 2 pores …………… .... PSEUDOCOCCIDAE 19. Anal ring with setae; anal lobes usually protruding but sometimes absent. Macrotubular ducts, when present, scattered over venter. Microtubular ducts present ……… ...... ERIOCOCCIDAE

Eriococcus buxi anal lobes - Anal ring simple, without pores or setae; anal lobes not distinctly protruding. Macrotubular ducts present in a ventral submarginal zone. Microtubular ducts absent ...... KERMESIDAE (in part)

DETECTION OF SCALE INSECT FAMILIES

4. INFORMATION SOURCES

Internet There are some outstanding, up-to-date, freely available information sources on the internet. Some of the most useful include the following: ScaleNet http://www.sel.barc.usda.gov/scalenet/scalenet.htm This is an excellent site, the objective of which is to provide comprehensive information on the scale insects of the world, including queriable information on their classification, nomenclatural history, distribution, hosts and literature. Keys to Scale Families and important species http://www.sel.barc.usda.gov/ScaleKeys/index.html This is a LUCID expert system that will help make identifications of the extant Coccoidea families; and species of Coccidae, Pseudococcidae and some of the other families intercepted at US ports-of-entry. Selected literature published as hard copy WORLD FAUNA Ben-Dov, Y. 1981. A catalogue of the Conchaspididae (Insecta, Homoptera, Coccoidea) of the world. Annales de la Société Entomologique de France 17 , 143-156. Ben-Dov, Y. 1993. A systematic catalogue of the soft scale insects of the world (Homoptera: Coccoidea: Coccidae) with data on geographical distribution, host plants, biology and economic importance . Flora & Fauna Handbook, No. 9. Sandhill Crane Press, Gainesville, FL. 536 pp. Ben-Dov, Y. 1994. A systematic catalogue of the mealybugs of the world (Insecta: Homoptera: Coccoidea: Pseudococcidae and Putoidae) with data on geographical distribution, host plants, biology and economic importance. Intercept Limited, Andover, UK. 686 pp. Ben-Dov, Y. 2005. A Systematic Catalogue of the Scale Insect Family Margarodidae (Hemiptera: Coccoidea) of the World . Intercept Ltd., Wimborne, U.K.. 400 pp. Ben-Dov, Y. & German, V. 2003. A Systematic Catalogue of the Diaspididae (Armoured Scale Insects) of the World, Subfamilies Aspidiotinae, Comstockiellinae and Odonaspidinae. Intercept, Andover, Hants, U.K.. 1112 pp. Fernald, M.E. 1903b. A catalogue of the Coccidae of the World. Bulletin of the Hatch Experiment Station of the Massachusetts Agricultural College 88 , 1-360. Hodgson, C.J. 1994. The scale insect family Coccidae: an identification manual to genera. CAB International, Wallingford, Oxon, UK. 639 pp. Kozár, F. 2004. Ortheziidae of the World . Plant Protection Institute, Hungarian Academy of Sciences, Budapest, Hungary. 525 pp. Kozár, F. & Miller, D.R. 2000. World revision of Ortheziola Šulc (Homoptera: Coccoidea: Ortheziidae) with descriptions of eleven new species. Systematic Entomology 25 , 15-45. Miller, D.R. & Williams, D.J. 1995 (1993). Systematic revision of the Family Micrococcidae (Homoptera: Coccoidea), with a discussion of its relationships, and a description of a gynandromorph. Bollettino del Laboratorio di Entomologia Agraria 'Filippo Silvestri'. Portici 50 ,199-247.

DETECTION OF SCALE INSECT FAMILIES

PALAEARCTIC Ben-Dov, Y. 1971. An annotated list of the soft scale insects (Homoptera: Coccidae) of Israel. Israel Journal of Entomology 6, 23-34. Danzig, E.M. 1986. Coccids of the Far-Eastern USSR (Homoptera: Coccinea). Phylogenetic analysis of coccids in the World fauna. Amerind Publishing Co., New Delhi, India. 450 pp. Ferris, G.F. 1950. Report upon scale insects collected in China (Homoptera: Coccoidea). Part I. (Contribution no. 66). Microentomology 15 , 1-34. Ferris, G.F. 1950a. Report upon scale insects collected in China (Homoptera: Coccoidea). Part II. (Contribution no. 68). Microentomology 15 , 69-97. Kosztarab, M. & Kozár, F. 1978. Scale insects - Coccoidea. (In Hungarian; Summary In English). Fauna Hungariae, Akadémiai Kiadó, Budapest 17 , 1-192. Kosztarab, M. & Kozár, F. 1988. Scale Insects of Central Europe. Akademiai Kiado, Budapest. 456 pp. Kozár, F. (ed.) (1998): Catalogue of Palaearctic Coccoidea. Plant Protection Institute, Hungarian Academy of Sciences, Budapest, 1-526. Williams, D.J. 1962. The British Pseudococcidae (Homoptera: Coccoidea). Bulletin of the British Museum (Natural History) Entomology 12 , 1-79. Williams, D.J. 1985. The British and some other European Eriococcidae (Homoptera: Coccoidea). Bulletin of the British Museum (Natural History) Entomology Ser. 51 , 347- 393. NEARCTIC Ferris, G.F. 1937. Atlas of the scale insects of North America. Stanford University Press, Palo Alto, California. Ferris, G.F. 1937a. Contributions to the knowledge of the Coccoidea (Homoptera). IV. (Contribution no. 5). Microentomology 2, 1-45. Ferris, G.F. 1938. Atlas of the scale insects of North America. Series 2. Stanford University Press, Palo Alto, California. Ferris, G.F. 1942. Atlas of the scale insects of North America. Series 4. Stanford University Press, Palo Alto, California. Ferris, G.F. 1950b. Atlas of the Scale Insects of North America . Series 5. The Pseudococcidae (Part I). Stanford University Press, Palo Alto, California. 278 pp. Ferris, G.F. 1953. Atlas of the Scale Insects of North America, v. 6, The Pseudococcidae (Part II). Stanford University Press, Palo Alto, California. 506 pp. Ferris, G.F. 1955. Atlas of the Scale Insects of North America, v. 7, the Families Aclerdidae, Asterolecaniidae, Conchaspididae Dactylopiidae and Lacciferidae. iii. Stanford University Press, Palo Alto, California. 233 pp. Gill, R.J. 1988. The Scale Insects of California: Part 1. The Soft Scales (Homoptera : Coccoidea : Coccidae). California Dept. of Food & Agriculture, Sacramento, CA. 132 pp. Gill, R.J., Nakahara, S. & Williams, M.L. 1977. A review of the genus Coccus Linnaeus in America north of Panama (Homoptera: Coccoidea: Coccidae). Occasional Papers in Entomology, State of California, Department of Food and Agriculture 24 , 44. Hamon, A.B. & Williams, M.L. 1984. The soft scale insects of Florida (Homoptera: Coccoidea: Coccidae). Arthropods of Florida and Neighboring Land Areas. Fla. Dept. of Agric. & Consumer Serv. Div. Plant Ind., Gainesville. 194 pp. Howell, J.O. & Williams, M.L. 1976. An annotated key to the families of scale insects (Homoptera: Coccoidea) of America, North of Mexico, based on characteristics of the adult female. Annals of the Entomological Society of America 69 , 181-189. Kosztarab, M. 1996. Scale insects of Northeastern North America. Identification, biology, and distribution . Virginia Museum of Natural History, Martinsburg, Virginia. 650 pp.

DETECTION OF SCALE INSECT FAMILIES

McKenzie, H.L. 1967. Mealybugs of California with taxonomy, biology, and control of North American species (Homoptera: Cooccoidea: Pseudococcidae). Univ. Calif. Press, Berkeley. 526 pp. Miller, G.L. & Williams, M.L. 1990. Tests of male soft scale insects (Homoptera: Coccidae) from America north of Mexico, including a key to the species. Systematic Entomology 15 , 339-358. Miller, G.L. & Williams, M.L. 2002. Systematics of the adult male soft scales from America north of Mexico (Hemiptera: Coccidae). Contributions on Entomology, International 5 (2): 53-126. Williams, M.L. & Kosztarab, M. 1972. Morphology and systematics of the Coccidae of Virginia with notes on their biology (Homoptera: Coccoidea). Research Division Bulletin, Virginia Polytechnic Institute and State University 74 , 1-215. AFRICA Ben-Dov, Y. 1974. On the species of Conchaspididae (Homoptera: Coccoidea) from Africa and Madagascar with description of a new species. Revue de Zoologie Africaine 88 , 363- 373. Van Harten, A., Cox, J.M. & Williams, D.J. 1990. Scale insects of the Cape Verde Islands (Homoptera: Coccoidea). Courier Forschungsinstitut Senckenberg 129 , 131-137. ORIENTAL REGION Williams, D.J. 2004. Mealybugs of southern Asia. The Natural History Museum, London. 896 pp. AUSTRALASIAN REGION Hodgson, C.J. & Henderson, R.C. 2000. Coccidae (Insecta: Hemiptera: Coccoidea). Manaaki Whenua Press, Lincoln, Canterbury, NZ. 259 pp. Williams, D.J. 1985. Australian mealybugs (Special Publication No. 953). British Museum (Natural Hist.), Longon. 431 pp. Williams, D.J. & Watson, G.W. 1988. The Scale Insects of the Tropical South Pacific Region. Pt. 1. The Armoured Scales (Diaspididae). CAB International Institute of Entomology, London. 290 pp. Williams, D.J. & Watson, G.W. 1988a. The Scale Insects of the Tropical South Pacific Region. Pt. 2: The Mealybugs (Pseudococcidae). CAB International Institute of Entomology, London. 260 pp. Williams, D.J. & Watson, G.W. 1990. The scale insects of the tropical South Pacific region. Pt. 3: The soft scales (Coccidae) and other families. CAB International Institute of Entomology, London. 267 pp. NEOTROPICAL REGION Williams, D.J. & Granara de Willink, M.C. 1992. Mealybugs of Central and South America. CAB International, London, England. 635 pp.

DETECTION OF SCALE INSECT FAMILIES

REFERENCES

Cheesman, O.D., Clubbe, C., Glasspool, A.F. & Varnham, K. 2003 Dealing with invasive species: sharing knowledge and experience. In: M. Pienkowski ed. A Sense of Direction: a conference on conservation in the UK Overseas Territories and other small island communities. Over Norton: UK Overseas Territories Conservation Forum, 257-272. Available from: http://www.ukotcf.org . Ferris, G. F. 1937. Atlas of the scale insects of North America . Stanford University Press, Palo Alto, California. Gill, R. J. 1988. The Scale Insects of California: Part 1. The Soft Scales (Homoptera: Coccoidea: Coccidae). California Dept. of Food & Agriculture, Sacramento, CA. 132 pp. Hamon, A. B. & Williams, M. L. 1984. The soft scale insects of Florida (Homoptera: Coccoidea: Coccidae). Arthropods of Florida and Neighboring Land Areas. Fla. Dept. of Agric. & Consumer Serv. Div. Plant Ind., Gainesville. 194 pp. Hodgson, C. J. 1994. The scale insect family Coccidae: an identification manual to genera. CAB International, Wallingford, Oxon, UK. 639 pp. McKenzie, H. L. 1967. Mealybugs of California with taxonomy, biology, and control of North American species (Homoptera: Cooccoidea: Pseudococcidae). Univ. Calif. Press, Berkeley. 526 pp. Malumphy, C., Hamilton, M.A., Manco, B.N., Green, P.W.C., Sanchez, M.D., Corcoran, M. & Salamanca, E. 2012. Toumeyella parvicornis (Hemiptera: Coccidae), causing severe decline of Pinus caribaea var. bahamensis in the Turks and Caicos Islands. Florida Entomologist 95, 113-119. Martin, J. 1987. An identification guide to common whitefly pest species of the world (Homoptera, Aleyrodidae). Tropical Pest Management , 33: 298-322. ScaleNet. 2015. http://www.sel.barc.usda.gov/scalenet/scalenet.htm Varnham, K. 2006. Non-native species in UK Overseas Territories: a review. JNCC Report 372 . Peterborough: United Kingdom. Williams, D. J. 2004. Mealybugs of southern Asia . The Natural History Museum, London. 896 pp. Williams, D. J. & Watson, G. W. 1988. The Scale Insects of the Tropical South Pacific Region. Pt. 1. The Armoured Scales (Diaspididae). CAB International Institute of Entomology, London. 290 pp. Williams, D. J. & Watson, G. W. 1988a. The Scale Insects of the Tropical South Pacific Region. Pt. 2: The Mealybugs (Pseudococcidae). CAB International Institute of Entomology, London. 260 pp. Williams, D.J. & Watson, G.W. 1990. The scale insects of the tropical South Pacific region. Pt. 3: The soft scales (Coccidae) and other families. CAB International Institute of Entomology, London. 267 pp. Williams, D.J. & Granara de Willink, M.C. 1992. Mealybugs of Central and South America. CAB International, London, England. 635 pp.