Landcare Research - Manaaki Whenua

Landcare-Research -Manaaki Whenua

FNZ 66 - Diaspididae (Insecta: Hemiptera: Coccoidea) - Methods and conventions

Henderson, RC 2011. Diaspididae (Insecta: Hemiptera: Coccoidea). Fauna of New Zealand 66, 275 pages.
( ISSN 0111-5383 (print), ISSN 1179-7193 (online) ; no. 66. ISBN 978-0-478-34726-5 (print), ISBN 978-0-478-34727-2 (online) ). Published 23 May 2011

Methods and conventions

Slide-mounting methods

1. Preparation for maceration. Remove 1 or more specimens from the preserving fluid (e.g., 75% ethanol) and place in a small flat dish; under the binocular microscope:

(a) for diaspidids in general, including nymphs, make a pinprick hole at about the mid-metathorax, or if the body contains many eggs, a mid-lateral slit on one side to allow extraction of the body contents.

(b) for pupillarial female diaspidids, lay the specimen dorsum down, hold in place with a blunt pin, and with a second pin with sharp hooked tip make a running slit inside the ventral margin of the pupillarium from near the posterior to near the head, being careful to avoid slitting the membranous adult female; just make a pinprick through to the adult female body; excess numbers of eggs/neonates can be removed if present, but leave the adult female in situ.

2. Optional first heat treatment: recommended for samples previously stored dried, for those stored in ethanol for a long time, and generally for any where speed is not critical.

Transfer the scales to a heat resistant glass tube (test tube) that has about 1 cm depth of 95% ethanol in it; put a bung of cotton wool in the top; place the tube in a water bath (about 95°C), and heat for 2 minutes.

3. Hot water bath maceration in 10% KOH. Transfer the scales to a clean heat resistant glass tube (test tube) which has about 1 cm depth of 10% KOH in it; put a bung of cotton wool in the top.

Place the tube in a water bath (about 95°C) and heat for between 3 and 5 minutes, or until the scale body becomes pale/clear.

4. Removing body contents. Tip the scales into a clean small dish, and using bent pins or other flattened tools, carefully pump and tease out the remaining body contents.

For pupillarial diaspidids, gently tease out the membranous female from her pupillarium and straighten if needed.

Note: if a specimen is fatty and difficult to pump, and especially if it folds over, stop and do not try to remedy at this stage. Just carry out the transfers with care and wait until the de-waxing stage when it is easier to fix the problem.

All the next stages are in covered staining wells. With a felt-tip pen, write an identifying code name or sample number on the glass cover.

5. Rinse and staining. Transfer to distilled water in a staining well for about 5–10 minutes; then follow one of staining methods below:

Fuchsin stain: add 1–2 drops of acid fuchsin stain (see below for recipe), leave for about 30 minutes or longer until the specimens are a good bright pink colour, then continue to step 6.

McKenzie’s stain: transfer to a staining well containing McKenzie’s stain (see below for recipes) leave for about 60 minutes in a warming oven (40oC), or longer if at room temperature, then continue to step 6.

6. Rinse, dewaxing, and dehydration

–Transfer to a clean staining well with 75% ethanol and leave for a few minutes.

–Transfer to 95% ethanol and leave for a few minutes (=dehydration).

–Transfer to a dewaxing mixture for 2–10 minutes, made up fresh as follows:

–in a small measuring cylinder mix (1 : 1) about 0.5 ml xylene and 0.5 ml absolute Isopropanol

–Transfer to straight absolute Isopropanol and leave for 5–10 minutes (= washing and final dehydration)

–Transfer to 1–2 drops of clove oil and leave from 2 minutes up to 2 days (the specimens may become brittle if left longer than this).

7. Slide-mounting

–Take a clean glass slide and put a small drop of clove oil in the centre

–With a pick-up tool, transfer a scale insect from the staining well of clove oil to the drop of oil on the slide

–Under the microscope arrange the specimen

–With a small piece of paper tissue, very carefully blot away the excess clove oil from the specimen

–Add a very small drop of Canada balsam beside the specimen on the slide, and with a pin gently run the balsam over the specimen (this sticks it in place)

–With a felt pen, write the code label or sample number on the slide and put aside lying flat to set for a few minutes

–Add a larger drop of Canada balsam by gently dropping on top of the specimen

–With forceps, carefully place on a clean cover slip, lowering it at one edge first.

Leave the slide to dry in a drying oven at about 40°C for 2 weeks, or at room temperature for about 6 weeks. It is permissible to take out of the oven sooner for examination, if care is taken and the slide is stored flat.

The slide must be fully dried before storing on its side, or before examination with oil immersion microscopy.

8. Labelling of slides

Add a printed label with full collection details, starting with country (e.g. NEW ZEALAND) in first line.

Add a second identification label with Family, Genus, and species, if determined; it is helpful to include the number of specimens and their life stages present on each slide.


1. Acid Fuchsin Stain

To make stain from the powder: add 0.35 g acid fuchsin and 25 ml glacial acetic acid to 75 ml distilled water.

2. The suggested stock staining solution is made up of 30 ml Essigs solution with 60 drops of McKenzie’s triple stain added. This has a shelf life of several years under normal storage conditions and with care can be reused many times.

Essig’s Solution:

Lactic acid ……………… 20 parts

Phenol ([liquefied) ………. 2 parts

Glacial Acetic Acid …….   4 parts

Water, distilled ………….  1 part

Mckenzie’s triple stain:

Equal parts of

Acid fuchsin


Lignin pink

Each made up into a 2% aqueous solution.


The key to genera in New Zealand doubles as a key to species for those genera with a single species present. Additional keys to species are provided for genera with more than one species present.


The line drawings of 20 cosmopolitan species and the morphology figures 1–5 are reproduced with kind permission from Miller & Davidson (2005). One line drawing (Poliaspoides leptocarpi adult female) is reproduced with kind permission from Yair Ben-Dov (1976). All the remaining illustrations were prepared by drawing over montaged images of slide-mounted specimens using the software packages Adobe Illustrator and Photoshop. The figures are presented as a map in the customary manner, with dorsal features on the left-hand half and ventral features on the right-hand half, and with detailed vignettes not to scale. Fig. 6–11 of slide-mounted female pygidia and Fig, 24, 26, and 117 were produced using Automontage and Helicon imaging software.

Morphology and terms

Adult female, see Fig. 3, 4, and 5 for general schematic morphology of aspidiotines and diaspidines. The female aspidiotine body is approximately round, ovoid, or turbinate, while that of the diaspidine is often fusiform. Segmentation of the body is not well differentiated, especially the head and thorax (cephalothorax or prosoma), and pygidium (posterior abdomen). The mesothorax is distinguished by the anterior spiracles and the metathorax by the posterior spiracles. Abdominal segments I–IV are the prepygidium (sometimes called free abdominal segments), and V–VIII are the pygidium. Pygidium segmentation is easier to define on the margin by the pair of segmental setae, 1 dorsal, 1 ventral, on each segment (Fig. 5).

Anal opening is on the dorsal surface of the pygidium. The length of the opening and distance from the margin are measured. Length of the anal opening does not include the sclerotised outer ring, and the distance to the pygidium margin is that from the posterior margin of the anal opening to the setae at the base of the median lobes.

Antenna in the 2nd-instar nymphs and adult female is an unsegmented tubercle with 1 or more long setae, short stubs, and coeloconic sensilla. The number and shape of the setae are rather variable in some endemic species (e.g., Poliaspis spp.). Antennae of 1st-instar nymphs are 5- or 6-segmented with fine and fleshy setae.

Basal scleroses arise from lobes, usually from L1 (e.g., on A. nerii and Symeria species), or the medial lobule of L2, (e.g. on Anzaspis angusta and Pseudaulacaspis species) and are not to be confused with Paraphyses.

Bosses (the term preferred here, cicatrices of some authors) are sclerotised and convex round areas on dorsolateral submargins. These are important taxonomically in Lepidosaphes species and are present in nearly all Poliaspis species treated here.

Eye, nearly always inconspicuous (except in 1st-instar nymphs) but sometimes modified into a spur, which may point anteriorly or posteriorly (e.g., Abgrallaspis cyanophylli, Lepidosaphes pinnaeformis).

Exuvium (plural exuvia), the cast 1st and 2nd nymphal skins that are incorporated into the diaspidid scale cover with sheets of wax between them. The 1st-exuvium is the skin of the 1st-instar nymph, present on both female and male 2nd-instar nymph covers. The 2nd-exuvium is part of the adult female cover and not present on the male cover. Note: the terms exuvium/exuvia follow the precedence set by Takagi (1969), and are preferred to emphasise the distinctions between 1st and 2nd nymphal skins for their taxonomic importance.

Gland spines are usually simple, elongate, marginal projections with an associated microduct (Fig. 5), mostly located on the pygidium, and common in diaspidines.

Gland tubercles are short and broad versions of gland spines that occur on the submargins of prepygidial segments and thoracic areas, always with an associated microduct. It is convenient to distinguish them from gland spines here although some authors consider them one and the same (Miller & Davidson 2005).

Lobes occur on the pygidial margin and are said to act like “trowels” for spreading of wax in scale cover formation. The median lobes (L1) may be notched or serrate on their margins: important diagnostic characters are their shape and size, if yoked (also known as zygotic, meaning joined at the base, see Fig. 5), and with/without basal scleroses. Second (L2) and third (L3) lobes are also diagnostic; they may be single or bilobed (divided), or lacking, or mere points. Length of lobes on the pygidium includes the basal sclerosis when present. Presence/absence of an obvious pair of setae between the median lobes is important.

Macroducts or dorsal ducts are cylindrical wax glands, either 1-barred or 2-barred (Fig. 5). One-barred ducts may be very long and often their openings are arranged in linear furrows on the pygidium. Two-barred ducts may be barrel-shaped and in the diaspidines are generally arranged in segmental rows on the margins and submargins of the dorsal pygidium and prepygidium, they may, however, be widely scattered over the dorsum, as well as present on the ventral submargins of the anterior abdomen and thorax. Larger macroducts on the pygidium margin are known as marginal macroducts. On 1st-instar nymphs/1st exuvia the presence/ absence of a pair of dorsal cephalic ducts (head ducts) is taxonomically important.

Microducts are thin tubular wax glands most often present on the venter, associated with gland spines, gland tubercles, plates, and grouped near the spiracles. Their presence/absence may also be diagnostic on the thoracic dorsum of 1st-instar nymphs/1st-exuvia.

Paraphyses (Fig. 5) are sclerotised rods on the pygidium that usually arise from the lateral margins of the lobes but occasionally also from the interlobular spaces (e.g., Lindingaspis rossi). They are thickened areas in interlobular spaces at the ends of duct furrows on Hemiberlesia species (Fig. 10–11) and absent there on Abgrallaspis cyanophylli and Aspidiotus nerii (Fig. 6–7), and thus a useful diagnostic aid. They are a very distinctive shape in Anoplaspis metrosideri.

Perispiracular pores are wax discpores grouped by the spiracles; whether present/absent, and the number of pores and loculi are diagnostic. In this volume, Anoplaspis and Parlatoria species have 5-locular pores, whereas all the rest have 3-locular pores (when present).

Perivulvar pores are 5-locular discpores clustered around the vulva, producing short c-shaped wax strands that adhere to laid eggs and prevent them sticking to surfaces or each other.

Plates are flat dermal projections on the pygidium margin that usually contain microducts and are usually fringed (fimbriate), more so in spaces between the lobes, and become simple anteriorly on the pygidium (further forward on the abdomen in Parlatoria species).

Pupillarium (plural pupillaria) is the sclerotised 2nd-instar exuvium that completely encloses the membranous 3rd-instar (adult) female in pupillarial species (see Henderson et al. 2010, p. 4). As the diminutive adjective pupillarial is the accepted term for this diaspidid lifestyle, the term pupillarium follows logically for this female exuvium. Previous authors have used the term puparia which is common to Lepidoptera and Diptera for the larval moult enclosing a metamorphosing male or female insect before emergence.

Pygidium is the more or less fused and sclerotised posterior abdomen including segments V–VIII. Dorsal features are the anal opening, with presence/absence of dorsal ducts, pre-anal scars, and paraphyses. Ventral features are the vulva, with presence/absence of perivulvar pores and microducts. Marginal features are the lobes, opening of marginal macroducts, with presence/absence of gland spines and/or plates, and various setae as important features.

Spiracles are not important diagnostically except for their associated perispiracular pores and for defining the mesothorax and metathorax.

Female 2nd-instar nymphs resemble a small adult female of their particular species, except lacking a vulva and perivulvar pores, and with fewer ducts and glands.

Male 2nd-instar nymphs often have a different body shape to the female nymph, either rounder or more elongate, and generally are more glandiferous. Male nymphs may have modified dorsal ducts. For example, Pseudaulacaspis brimblecombei (Fig. 207) and Ps. eugeniae (Fig. 211) both have remarkable communal duct clusters on the posterior abdomen, whereas Trullifiorinia acaciae has so-called ‘frame ducts’ as marginal macroducts (Fig. 236) (Howell & Tippins 1990). All male 2nd-instar nymphs have a line of 3 pairs of setae on the ventral head between the anterior margin and mouthparts, although these are less obvious on Symeria species. Dorsal ducts are often present on both dorsum and venter, and may form a band along the lateral margins where the body is rounded rather than dorsoventrally flattened. Marginal macroducts on the pygidium usually open slightly more ventrally than dorsally. Lobes on the pygidial margin may be digitate and divided or plate-like (e.g., Poliaspis lactea (Maskell), fig. 187; P. media when on Hebe, fig. 194). Marginal gland spines may be present (e.g., in Anzaspis, Fusilaspis) while in other genera they may be plate-like or absent on the pygidium but with gland tubercles present further forward (e.g., in Poliaspis). Faint vestigial leg patches may be seen on some Poliaspis (Fig. 193, 198) and Pseudaulacaspis (Fig. 207) species.

1st-instar nymphs are generally membranous except for appendages, pygidial lobes, and sometimes sclerotisation on the head (e.g., Poliaspoides, Fig. 204; Trullifiorinia , Fig. 237). They possess 3 pairs of 5-segmented legs where the tarsus is much longer than the tibia, and 5- or 6-segmented antennae. On the pygidium are at least 1 pair of lobes (although barely noticeable in Symeria), usually 1 or 2 pairs of gland spines, and a pair of very long apical setae. Always present is a line of 3 pairs of setae on the ventral head between the anterior margin and mouthparts (as on the 2nd-instar male nymphs). There is always a lateral microduct on each side of the abdominal segments, except in Anoplaspis species where the posterior 4 pairs are replaced by large macroducts (Fig. 128). Microducts on the dorsal thorax may be diagnostic, as is the presence/absence of a pair of large dorsal cephalic ducts near the anterior head margin (these are not 8-shaped as described by some authors).


Under ‘material examined’, the data given for each species are those on the slide label, categorised under the area codes of Crosby et al. (1998), followed by the NZAC accession number and the number of slides and specimens studied, including their life stages. For example: #09-073a–e [5]: 3 F, 2 f2nd, 2 m2nd, 4 1st denotes 5 consecutive slides were made (a–e) with a total of 3 adult females, 2 2nd-instar female nymphs, 2 2nd-instar male nymphs, and 4 1st-instar nymphs. Other life stage codes are: fpl (2nd-instar female pupillarium), exuv1 (1st-exuvium), exuv2 (2nd-exuvium), pp (prepupa), p (pupa), M (adult male).

A total of 9782 diaspidid specimens, including type specimens, were examined in this study. The number of NZAC specimens examined in addition to type material is listed under each species and in Appendix A.

Biota codes: A = adventive (exotic); E = endemic to New Zealand; N = native (Australasian) are indicated in bold font following each valid species name in the taxonomic section.

Type depositories

ANIC      Australian National Insect Collection, Canberra

BMNH    The Natural History Museum, London

NZAC     New Zealand Arthropod Collection, Auckland

QMBA    Queensland Museum, Brisbane

Purchase this publication