PLANNING AND ORGANIZING COMMITTEE 2012

President: Ross Kendall 07 3378 1187

Vice President: John Moss 07 3245 2997

Treasurer: Rob MacSloy 07 3824 4348

Secretary: Jennifer Singfield 07 3869 0359

Magazine: Daphne Bowden (daphne.bowden1 @bigpond.com) 07 3396 6334

Publicity and Library: Lois Hughes 07 3206 6229

Excursion Convenor: Alisha Steward 07 3275 1186

PLANNING AND ORGANIZATION MEETINGS

A quarterly meeting 1s scheduled in order to plan club activities and the magazine.

See BOIC Programme.

CONTACT ADDRESS AND MEMBERSHIP DETAILS

PO Box 2113, Runcorn, Queensland 4113

Membership fees are $30 for individuals, schools and organizations.

AIMS OF ORGANIZATION

e To establish a network of people growing butterfly host plants;

e To hold information meetings about invertebrates;

e To organize excursions around the theme of invertebrates e.g. butterflies,

native bees, ants, dragonflies, beetles, freshwater habitats, and others;

e To promote the conservation of the invertebrate habitat;

e To promote the keeping of invertebrates as alternative pets;

e To promote research into invertebrates;

e To encourage the construction of invertebrate friendly habitats in urban areas.

MAGAZINE DEADLINES

If you want to submit an item for publication the following deadlines apply:

March issue — February Ist June issue — May Ist

September issue — August Ist December issue — November Ist

COVER PAINTING

Underwater predators - left: Giant water bug (Belostomatidae), top: Needle bug

(Nepidae), and right: Water scorpion (Nepidae) — painting by Lois Hughes

BEDS PE DS PF OE Pe PE De De Pe bg Pe be pe Pe bd Pe bg

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 2

FROM THE PRESIDENT

Congratulations to our excursion convenor, Alisha Steward, who recently was

awarded a PhD by Griffith University with the Australian Rivers Institute. Her thesis

was titled: "When the river runs dry: the ecology of dry river beds". I find it

impossible to summarize the detail of her work and will try to persuade her to write a

small article about her findings for a future edition of Metamorphosis. You will find

her contributions on slightly wetter areas 1n this edition.

We also congratulate John Moss who recently received the Redlands City Council

Australia Day 2013 Environmental Award 1n recognition of his extensive work over

many years.

A club member, who wishes to remain anonymous, has donated a sum of money to

meet the membership fees of persons or groups who experience financial hardship

and, as such, are unable to join the Butterfly and Other Invertebrates Club. If you are

aware of any such persons or organizations, please discuss this offer with them with a

view to nominating them for membership.

Each quarter, I find it difficult to adequately acknowledge the various articles

produced by so many interesting writers but again: “Thank you”. You will find that

this edition is slightly larger. Several contributions have been held over for our June

edition!

Best wishes Ross

Creature Feature - Large, predatory water DUS ............cccccesscccessccceseecceeececeeececauscecaeeceeaeeceseeeeeeaees 4

Goliath Stick Insect : a phasmid of colour and Character .............ccccccccssscccessecceeeeceecceeesceeeeeceeeees 9

Moths - The Weird and the Wonderful 00.0... ccc cecccccessececesececeeeecceeeeeceseeeecesaeecessaeeeeseaeeeeaes 13

Observations of aquatic, bug-eating moth fly larvae... eccccsssccceseecceeecceeeeeceeeeeeeaeeeeeeeeeeeas 17

Life history notes on the No-brand Grass-yellow Eurema DriQittd ......cccccccccceseccceecceeneeceaeeeenees 21

CHET IOTGS PUCCISs dines scayastase oeearccavonpaaceuhe rede mete daomaohopkteny eau atone eledcoran ad devs Hieber Wd alersbe Redes had clo ie

Field Notes : Major extentions to the known distribution of the Bright Purple Azure

QV ESD OVINE oct oasesck subs es disPce selves sa Sate dalduSeabo od dsl oa Beeteoba te da’ 26

Book Reviews - A Guide to Australia's freshwater Crayfish ............ccccceseccccseccccesecceceecceeeeeeeees 32

The Moths of Victoria Vol. 4 ......ccccccccccssccccesscccceseececeeeececaeeeceeeeeceeseeceeseeeeeeas 37

Report - The day at Peter Hendry's identifying moths in the family Crambidae. ........................ 37

SV es IIS! TRAY 2s scvansara ave tipste apadesns veut dasugbatessys da tdotouga vedas oeotdata ea rite siya cgh:tdacaagaaina ings ooetdatoaga vnnsy ees 39

In a Corinda Garden - Some butterfly VIOWS .0..........cccecccccssecccessececeesecccaeeececeeeeceeeeecceeeneceeeeneceeeas 39

TFETTOIS: sacssottteeshtetseehcobidetesedebsaetbok itateahstesbd dochdabudabest led EMBeR AoA FAIDAR MOAR Aaa RE PRAR ON AAA. R aL TDRA LOREAL RAIORR Hod FRE 9 Ad 4]

You Asked and BOIC Programme .......... cece cceccccaesceceeeeccaeecccaeecccaencccaeececaueceeaeeeeeaeeeeeanes 43

BEDS REDS PF DE be PE DT De Pe bd Pe be pe Pe bd Pe bd

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 3

CREATURE FEATURE

Large, predatory water bugs — Alisha Steward

Many families in the Order Hemiptera (‘bugs’), Suborder Heteroptera (‘true bugs’),

are aquatic, living in rivers, streams, ponds, lakes and other wetlands — and almost all

aquatic bugs are predatory. This issue's creature feature focuses on two related

families of aquatic Hemiptera — Belostomatidae and Nepidae (Fig. 1). Belostomatids

are known as giant water bugs. Some nepids are known as water scorpions, as they

have a broad, leatf-like body, and others are known as needle bugs, as they have a thin,

stick-like body.

Fig. 1. Water bug body forms and their faces that only a mother could love: a) Giant water bug -

Lethocerus insulanus (Belostomatidae), b) Diplonychus sp. (Belostomatidae), c) and d) water

scorpion - Laccotrephes sp. (Nepidae), e) needle bug - Ranatra dispar (Nepidae), and f) needle

bug (Nepidae). Photograph a) by Peter Chew (www.brisbaneinsects.com) and photograph e) by

the Murray-Darling Freshwater Research Centre (www.mdfrc.org.au/bugguide/index.htm) are

used with permission

BEDS PE DS DE DE PS Pe Re be Pe he bd Oe bd Pe be bd Pe

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 4

Species of both belostomatids and nepids are large (15-75 mm long), and quite scary

bugs at that, as they can inflict a nasty bite! They are differentiated from other aquatic

bugs by their large size, the presence of raptorial forelegs (except in one African

genus), and the presence of paired respiratory processes at the tip of the abdomen

(Andersen and Weir, 2004). In belostomatids the respiratory processes form short,

retractable ‘air straps’, and in nepids they typically form a long siphon.

Belostomatids and nepids spend most, if not all of their life in water, although both

eroups have winged adult stages capable of flying between water bodies and are quick

to colonise new habitats (Gooderham and Tsyrlin, 2002). They can be attracted to

lights at night, and some nepids can even end up in swimming pools! I have observed

nepids in chlorinated swimming pools, presumably breathing through their respiratory

siphon at the end of their abdomen - poked through the surface film and used much

like a snorkel. They don’t seem too fussy about water quality — on the Australian

SIGNAL (Stream Invertebrate Grade Number Average Level) scale used to rate the

tolerance or sensitivity of water bugs to pollution, belostomatids score | out of 10,

meaning that they are extremely tolerant of pollution, and nepids score 3 out of 10,

suggesting that they are moderately tolerant of pollution (Chessman, 2003).

Habitat and diet

Belostomatids and nepids occur throughout Australia. They prefer still or slow-

moving water, and are usually found at the edge of streams or ponds where aquatic

plants are present. Both types of bugs are brownish in colour and are well

camouflaged amongst mud, leaves and twigs. They are typically ambush predators

that sit and wait for prey. Prey includes other invertebrates, tadpoles, small frogs and

even fish, all of which are captured with their forelegs. As a result of their dietary

preferences, belostomatids have been found to be a nuisance in fish farms

’ (Carver et al., 1991).

In preparation for this article

and its accompanying cover

drawing by Lois Hughes, I

visited Lois’ property at Mt

Cotton to look for our artist’s

models. The creek yielded

many water bugs, but no

belostomatids or nepids. Next,

we visited a dam on the

property, which was shallow

bs s-4 and had a great deal of leaf

Fig, 2. Typical predatory water bug habitat in 1 south- east . litter on the bottom and

Queensland — a shallow dam containing leaf litter. around the edges (Fig. 2). A

few scoops of the dip net

BIDS REDS PF OE be PE DS De Pe bs Pe be Pe Pe bd Pe be

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 5

revealed almost a dozen water scorpions! See Fig. 3 for a photograph of some that we

collected. Predatory water bugs are often the last aquatic invertebrates to be seen in

drying waters, so it was no surprise to see them in this shallow dam. They prey on any

remaining aquatic invertebrates, and then fly off when the water dries up. Lois kept a

few of the water scorpions

for observation, and in order

to feed them offered them a

erasshopper. The

grasshopper was quickly

grasped and greedily

devoured.

Life cycle

Males in both families can

produce sound to attract

females for mating, and |

often initiate mating by Fig. 3. Spot the water bugs! There are two water scorpions

‘display-pumping’ or (Nepidae: Laccotrephes sp.) in this photograph. Note their

rhythmical push-up-like leaf-like appearance.

body movements near the

water’s surface. Female belostomatids from the genus Diplonychus lay their eggs on

the back of the male for protection (Fig. 4), whereas females of the genus Lethocerus

lay eggs above the water on emergent vegetation that are defended by males in the

water below the egg mass (Andersen and Weir, 2004). Nepid eggs possess respiratory

horns, and are deposited at the water's edge or within plant tissue in a way that the

horns are exposed to the air for respiration. Eggs of both families hatch into nymphs,

which are similar to the adults — only smaller and usually lacking wings. Nymphs

reach the adult stage in 5 moults (Gooderham and Tsyrlin, 2002).

. :

eal

Fig. 4. Male Diplonychus sp. (Belostomatidae) with eggs.

a Od Dg Od OG Og DO OF OG DO Od Od OG Od Od Od DO OF

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 6

Interesting facts!

I observed one of the water scorpions “playing dead’ after 1t was touched — it rolled

over, drew its legs against its body, and stopped moving (Fig. 5). I imagine that this

response is to confuse and hide from potential predators, as once they stop moving

they blend in well with leaf litter and other debris.

Giant water bugs are consumed throughout many Asian countries. There are many

photographs of these bugs prepared as food — usually fried — on the web. Here 1s a

link to an example (there are many more if you search):

http://www. flickr.com/photos/cibergaita/56687803/

*. 4 ‘7 *

+" +

2 = rv

Fig. 5. This water scorpion (Nepidae: Laccotrephes sp.) appeared to ‘play dead’ when touched.

Conclusion

Andersen and Weir (2004) state that the biology of many Australian species 1s poorly

known. This 1s in stark contrast to our knowledge of many other invertebrate groups,

particularly butterflies, where we know much more about the biology of individual

species (and even sub-species) — such as what plants they lay their eggs on, what time

of year they are on the wing, how long the pupal stage lasts, and even information

about caterpillar parasitoids. The lack of species-specific information 1s the reason

that this article features a group of taxa, rather than one species. Aquatic Hemiptera,

particularly belostomatids and nepids, are common in Australia, can be found in farm

dams and backyard ponds, and can be reared in captivity, making them easy to study.

These interesting water bugs are certainly worthy of more detailed research.

Classification and species checklist

Belostomatids and nepids belong to the superfamily Nepoidea. The family

Belostomatidae is recognised in Australia by only 2 genera: Diplonychus and

Lethocerus. The family Nepidae is represented in Australia by 5 genera — Austronepa,

Cercotmetus, Goondnomdanepa, Laccotrephes and Ranatra. Austronepa and

Goondnomdanepa are endemic to Australia.

Australian species checklist (based on Hawking et al., 2009):

Order Hemiptera

Suborder Heteroptera

Infraorder Nepomorpha

Superfamily Nepoidea

BEDS PE DS PF DE be PE DT pe Pe bd Pe be pe Pe bd be bg

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 7

Family Belostomatidae — giant water bugs

Diplonychus eques Dufour

Diplonychus rusticus Fabricius

Lethocerus distinctifemur Menke

Lethocerus insulanus Montandon

Family Nepidae

Subfamily Nepinae — water scorpions

Laccotrephes tristis Stal

Subfamily Ranatrinae — needle bugs

Austronepa angusta Hale (endemic)

Cercotmetus brevipes Lansbury

Goondnomdanepa brittoni Lansbury (endemic)

Goondnomdanepa prominens Lansbury (endemic)

Goondnomdanepa weiri Lansbury (endemic)

Ranatra dispar Montandon

Ranatra diminuta Montandon

Ranatra ocidentalis Lansbury

References

Andersen, N.M. & Weir, T.A. (2004). Australian Water Bugs: Their Biology and

Identification (Hemiptera-Heteroptera, Gerromorpha & Nepomorpha).

Apollo Books, CSIRO Publishing: Collingwood, Victoria.

Carver, M., Gross, G.F. & Woodward, T.E. (1991). Hemiptera. In: ‘The Insects of

Australia: A Textbook for Students and Research Workers.’ Chapter 30, 2"°

Edition, Volume 1. Melbourne University Press. Pp. 429-509.

Chessman, B.C. (2003). New sensitivity grades for Australian river macroinvertebrates.

Marine and Freshwater Research, Volume 54: 95-103.

Gooderham, J. & Tsyrlin, E. (2002). The Waterbug Book: A Guide to the Freshwater

Macroinvertebrates of Temperate Australia. CSIRO Publishing: Collingwood,

Victoria.

Hawking, J.H., Smith, L.M., Le Busque, K. (editors) (2009). Identification and

Ecology of Australian Freshwater Invertebrates. http://www.mdfirc.org.au/bugguide

Version January 2009. Accessed 20/01/2013.

Further reading

Online identification key to distinguish between the families Belostomatidae and

Nepidae (the same website can be used to identify other aquatic invertebrates) —

http://www.mdirc.org.au/bugguide/display.asp?type=4&class=17&Subclass=& Order

=3&Family=& genus=&species=&couplet=14&fromcouplet=13

Peter Chew’s website to insects and spiders of Brisbane, including aquatic

invertebrates: www.brisbaneinsects.com

BEDS REDS PF DE be PE DT pe Pe bs Pe he pe Pe bd Pe bg

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 8

Solution to Fig. 3:

Photos, except where otherwise

credited, Alisha Steward

ITEMS OF INTEREST

The Goliath Stick Insect: a phasmid of colour and character —

Densey Clyne

Generally speaking I don't like to keep insects in cages. Most insects need a lot of

space, especially flying ones that will simply batter themselves to death trying to

escape. However there are some insects that take very well to life in a cage, or even

free in the house if properly fed. Phasmids for instance make rewarding house pets.

They are harmless, peaceful - indeed lethargic - and easy to feed. As a bonus their

droppings make convenient fertiliser pellets for indoor pot-plants.

One of my favourite ‘pet’ insects 1s the Goliath Stick insect (Eurycnema goliath), the

most colourful Australian phasmid and one of the biggest. The bulky females can

reach up to 25 cm long, but they can still be hard to locate up 1n the tree-tops where

they prefer to feed. The young stages are brown and stick-like, and the mainly leaf-

ereen adults while being well camouflaged among the leaves are surprisingly

colourful close up.

2"° instar nymph Wings expanding after moult

BEES PE DS PE De PS Pe Re be Pe he be Oe bs Pe be bd Pd

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 9

Newly moulted female with exuviae Mature female

Underside colour pattern of female Female feeding on eucalyptus leaf

In captivity it is easy to follow and photograph the life of this charming insect, to

record the colour changes through each ecdysis as the wings unfold, and to get a close

look at that strange little face. But what intrigues me most is a remarkable aspect of

the Goliath stick insect’s biology that it shares not only with many other phasmids but

surprisingly with some plants. It’s a mutualistic association with ants.

BEDS PADS PE DE Pe PE De Pe Pe bg Pe he be Pe bg be bd

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 10

Female face Male and female mating

Many plant species have an arrangement with ants to distribute their seeds called

myrmecochory (Gk. myrmeco = ant and chory = dispersal). The seeds of these plants

have attached to them a small body called an elaiosome (Greek for ‘oil-body’ ) that is

attractive to ants. A foraging ant will pick up a fallen seed, return home and feed this

nutritious morsel to its young siblings, the ant larvae. The hard seed remains intact

and is discarded or added to the ants’ “midden’. If conditions are otherwise right it

will germinate and flourish there.

Now comes the phasmid connection. It involves the phenomenon of convergent

evolution that occurs within both the fauna and the flora. The term 1s used when two

unrelated organisms, often separated geographically, independently develop similar

characteristics in response to similar environmental pressures. Think of hedgehogs

and echidnas (spiky bristles for defence); ducks and platypuses (broad bills for

foraging); mantids and mantispids (raptorial legs for predation); and of course birds,

bats and insects (wings for flight). Best known among plants are American cacti and

African succulents (water storage for desert survival).

PEPE PERS DE DE PS Pa RT be Pe he be Oe bd Pe be bd Pd

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 11

There’s another example of convergent evolution that I find quite amazing because it

involves structures and functions that cross the borderline between flora and fauna. It

relates to the dispersal of insect eggs on the one hand and plant seeds on the other,

that is performed in the same way, by the same agents - ants.

The eggs of many phasmids, including the Goliath Stick insect’s, actually look like

seeds. They have a knoblike attachment on them called a capitulum, analogous to the

elaiosome of a myrmecochorous plant. Like the seed’s elaiosome, the egg’s

capitulum tempts a foraging ant to carry the egg into the nest. As with the seed’s

elaiosome the egg’s capitulum is fed to the ant larvae, leaving the egg intact.

But first the eggs must get to the ground. In Image No. 9 the female’s scoop-like rear

appendage (the operculum) can be seen holding an egg in place. The operculum is

used like a bat (or pe/ota) to toss the egg out and away from the female’s own food

plant. She can only toss it so far (though I’ve seen one throw her eggs right across my

kitchen!) but then the ants take over, completing the eggs’ dispersal to fresh fields.

Female operculum with egg Goliath (Eurycnema goliath) eggs

The advantage? It is in the future phasmids’ interests to be taken to a possibly-

untouched food source. Perhaps in the female’s best interests too, reducing

competition for food by her offspring. Also, with aggressive ants in their vicinity the

eggs may have the benefit of protection from predators and parasites. And being

underground 1s no problem — it seems the newly hatched phasmid nymphs can dig

themselves out from as deep as 6 cm. Now those vulnerable nymphs that escape

attack by their former benefactors can go scurrying up into the treetops for safety!

So there’s more to the Goliath stick insect than just a funny face. Her story gives us

yet another reminder of the interdependence of all life on this remarkable planet of

ours.

Reference

Hughes L .and Westoby M., 1992: Capitula on stick insect eggs and elaiosomes on

seeds; convergent adaptations for burial by ants. Functional Ecology 19926, 642-648

Photos Densey Clyne

PEPE PE DS PF OE be PE De De PE be Pe he pe Pe bd Pe bg

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 12

Moths -The Weird and the

Wonderful - Graham J. McDonald

Introduction

The world of very small and unusual moths

often goes unnoticed. Their interesting

colours, shapes and patterns only become

apparent after taking their images using a

good macro lens and DSLR camera.

Knowing where to look for them, how to

attract them and getting to know their habits

and distribution all help to make this pastime

very rewarding. All the images shown here

were taken using a hand-held Canon 450D

DSLR camera and Canon macro {2.8 100

mm lens. All moths were live unpinned

specimens with a few in their natural habitat.

Weird Moths

Fig. 1 COSMOPTERIGIDAE:

Pyroderces terminella (Walk.)

Tallebudgera Creek Estuary, Gold Coast

in Casuarina glauca forest

The larvae of this moth invade deserted

Polistes wasp nests. They have also been

reared from larvae eating dead leaves,

remains of dead insects and galled flower

buds of Acacia binervata. They have also

been found infesting the egg-sacs of the

golden orb-weaver spider (Nephila edulis)

and feeding on the eggs.

Note: There were many orb-weavers

where the photo was taken.

BEDS PERS PF OE be PE DT De PE be Pe he be Pe bd be bg

Fig. 2 COSMOPTERIGIDAE:

Cosmopterix heliactis

Wongi Waterholes, Maryborough -

attracted to light

Wing span approx. 9 mm (Length 6 mm)

This genus contains about 10 Australian

species. The tiny adults are superficially

similar to each other. Larvae are leaf and

stem miners of grasses and sedges.

Fig. 3 COLEOPHORIDAE:

Coleophora alcyonipennella

Mudgeeraba, Gold Coast - attracted to light

Wing span approx. 10 mm

This brilliant metallic moth is an exotic

species which is naturalised in areas of

Australia where the food plants are found.

Larvae feed on the flowers and seeds of

Trifolium spp. (clovers) and Medicago spp.

(medics).

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 13

Fig. 4 TINEODIDAE: F ig. 5 DEPRESSARIIDAE:

Peritornenta circulatella

North Burleigh Headland, Gold Coast on

Cupaniopsis anacardioides (tuckeroo)

Wing span approx. 20 mm

There are six species of Peritornenta in

Australia, all in northern areas. They are

found in rainforest, soft wood scrubs and

Cenoloba obliteralis

Tallebudgera Creek Estuary, Gold Coast

in mangrove community

Wing span approx. 8 mm

Moths of this family are rarely seen. They

are tropical and sub-tropical in

distribution. This one occurs from |

Innisfail, Qld to Grafton, NSW. Larvae woodland. P. circulatella occurs in the NT

and from north Qld to Ballina, NSW, inland

as well as coastal. This moth resembles a

damaged Smilax leaf and was photographed

in habitat where the larvae join together

leaves of C. anacardioides. They feed

while hidden in the shelter so formed.

When mature, larvae leave the shelter and

pupate exposed on a leaf or other object.

Pupae are also cryptically coloured.

feed in the live seeds of Avicenna marina

(Grey Mangrove). Cenoloba obliteralis

resembles moths of the family

Pterophoridae, the plume moths which

also have deeply incised wings.

Fig. 6 DECOPHORIDAE:

OECOPHORINAE: Wingia psittacodes They are boldly coloured in shades of

Pine Ridge Conservation Park, Gold Coast orange, pink and cream and have bent

in dry coastal heath wing tips. Larvae of some species tie

Wing span approx. 22 mm leaves of eucalypts together with silk and

Moths of this genus usually have upturned _ hide and feed inside. This species

palpi, the base of which is covered in hairy probably ties the leaves of Leptospermum

scales. spp. together.

BEDS PE DS PF OE be PE Pe De PE bd Pe be pe Pe bd be bg

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 14

Wonderful Moths

Fig. 7 AGANAIDAE: Agape chloropyga

Mudgeeraba, Gold Coast in planted

rainforest

Wing span approx. 48 mm

This large spectacular moth 1s active

during the day but also comes to light at

night. Its ability to mimic a falling

senescent yellow fig leaf protects 1t from

predators. The moth’s distribution 1s from

Coen on Cape York Peninsula, Qld to

Lismore, NSW. The larvae feed on a

variety of Ficus spp. (fig trees).

se |

Fig. 9 NOCTUIDAE: CATOCALINAE:

Donuca orbigera

Wongi Waterholes, Maryborough in

eucalypt woodland

Wing span approx. 55 mm

Common in tropical eucalypt woodland

from Cape York Peninsula, QLD to

Sydney, NSW. Its food plant and life cycle

are unknown.

Fig. 8 GEOMETRIDAE: GEOMETRINAE:

Pea a?®

. i :

. .

«, n _

' 4

Uliocnemis partita

Mudgeeraba, Gold Coast - attracted to light

Wing span 38 mm

This beautiful emerald geometrid moth lives

in forest where its cryptic colours hide it

from predators as it rests on lichen encrusted

tree trunks. Larvae feed on Acacia leiocalyx

(early flowering black wattle) and Acacia

disparrima (hickory wattle). They disguise

themselves with buds and flower parts of the

host plant. They are pinkish brown with a

dark dorsal line and have flanges along the

back to which the plant parts are attached.

Pupae are also covered by plant detritus.

Fig. 10 GEOMETRIDAE:

GEOMETRINAE: Anisozyga insperata

Mudgeeraba, Gold Coast - attracted to light

Wing span 30 mm

Another beautiful emerald geometrid from

rainforest habitats.

BEDS PE DS PF OE be PE DS pe Pe be Pe be pe Pe bd be bg

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 15

Fig. 11 NOCTUIDAE: CATOCALINAE: Fig. 12 ARCTITDAE: LITHOSHNAE:

Eudocima salaminia Schistophleps albida

Mudgeeraba, Gold Coast in moist forest Mudgeeraba, Gold Coast

types Wing span 12 mm

Wing span 75 mm This small elegant moth often flies

The larvae of this fruit-piercing moth chew during daylight but it is also attracted to

leaves of vines of the Menispermaceae light. It seems to be common in areas

family. Among these are Stephania where lichens grow.

Japonica, Tinospora spp. and Legnophora

moorei. Adult moths pierce ripe fruit and

suck the sugary fluids.

References

Common, I. F. B. (1990). Moths of Australia, Melbourne University Press

Herbison-Evans, Don and Crossley, Stella http://lepidoptera.butterflyhouse.com.au

Richardson, B. (2008). Mothology: Discover the Magic, LeapfrogOz

Zborowski, P. and Edwards, E. D. (2007). A Guide to Australian Moths,

CSIRO Publishing

Acknowledgements

Ted Edwards (ANIC - CSIRO) for identifying some of the moth species and

providing accurate life cycle data

Peter Hendry who assisted in identification and checking of the manuscript

John Moss for his attention to detail in fine tuning the manuscript

Photos Graham McDonald

BEDS Pa De Pe a he be Pe Pe Pe he bd be be be Pe bd bg

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 16

Observations of aquatic, bug-eating moth fly larvae! —

Alisha Steward

Moth flies belong to the fly family Psychodidae. The adults are sometimes also called

‘drain flies' because they are often found close to sluggish or stagnant water in which

their aquatic larvae live, including household drains. I see their larvae now and then in

invertebrate samples from rivers and waterholes that I collect for my work as a

freshwater scientist, but otherwise I have never had a close encounter with them —

until recently. I discovered that Psychodidae larvae can consume dead bugs in

contaminated water! Here 1s the story...

Occasionally, native citrus bugs (Musgraveia sulciventris), also known as bronze

orange bugs, feed on my citrus trees. I usually tolerate them. However, 1n the summer

of 2012/2013 they increased greatly in numbers into an infestation and were severely

affecting the trees — I needed to control them. Not only do they damage citrus trees,

they can also squirt nasty caustic fluid at predators, such as me, which burns the skin

and can stain it yellowish-brown for days! Instead of spraying the trees with

insecticide, I removed the bugs individually with tongs and dropped them into a

bucket of soapy water — soapy to break the surface tension so they would sink and

drown. I forgot about my haul of evil citrus bugs, and they remained in a bucket under

the house for a few weeks. On discovering the rancid bucket of dead bugs, I noticed

life! There was movement in the water. Moth fly larvae had colonised the bucket, and

they were feeding on the dead bug bodies, apparently undeterred by the presence of

washing up detergent!

Some moth fly larvae are aquatic, and are reasonably tolerant of pollution (Chessman,

2003), and some species are known to tolerate low oxygen levels and eutrophic

(nutrient-rich) waters (Hawking et al. 2009). My larvae were able to survive in the

soapy water of the bug bucket, although I did use biodegradable dishwashing liquid,

which may have degraded during the preceding weeks. Interestingly, the water was

not colonised by mosquito larvae until I topped up the bucket with rainwater. Moth

fly larvae can be distinguished morphologically from other aquatic fly larvae, such as

mosquito wrigglers, by having the following combination of characters: sclerotised

(hardened) dorsal sections on the thorax and abdomen (Fig. 1); no prolegs; and a

complete head capsule (some dipteran larvae have retractile or incomplete head

capsules) (Williams, 1980, Hawking et al., 2009). The larvae I observed were less

than 10 mm in length. Other features I noticed include a posterior respiratory tube

(Fig. 1), and hairs covering the thorax and abdomen.

BEDS PE DS PF OE be PE DT De Pe bd Pe be pe Pe bd Pe hd

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 17

sclerotised

plates

respiratory

tube

a) larva — lateral view b) larva — ventral view

Fig. 1. Psychodidae larva

Over the following days of observation I noticed aquatic pupae in the bucket, and

adult moth flies resting above the water on the sides of the bucket. The pupa I

photographed had a pair of anterior respiratory horns (Fig. 2), lateral and ventral

spines on its abdomen, and it did not retain the dorsal sclerotised plates characteristic

of the larvae.

b) pupa —respiratory horn

Fig. 2. Psychodidae pupa

You can see from the photographs of the grey-coloured adults (Fig. 3) that they

acquired their name 'moth flies' from the fluffy appearance of their heads, wings, and

bodies. I think that they are quite cute little flies! They weren't easily disturbed, and

settled nicely on the walls of a glass photography tank so that I could take some snaps

of them. The adult moth fly that I photographed kept perfectly still, until I was

finished. The larvae were another matter - they were full of beans! I observed them

crawling around on the bottom of the photography tank, and they were not keen to

cooperate as nicely as the adults for their portrait. I observed the larvae crawling

underwater, much like a caterpillar, and at a rapid pace. This resulted in numerous

out-of-focus pictures! I assume that the larvae were heading for the shelter of leaf

litter or other debris, to reduce the risk of predation by being exposed. I used a

microscope with a camera attachment so I could take some better shots.

BEDS PED PE DE PS Pe Re be Pe De be Oe bd Pe be bd Pe

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 18

Fig. 3. Psychodidae: a) larva ventral and lateral views; b) pupa ventral and dorsal views;

and c) adult ventral and dorsal views

Over the following weeks the moth fly larvae consumed much of the dead citrus bug

material. I read stories about them on the internet, and 1t seems that people consider

them a nuisance, asking for advice on how to get rid of them. On Wikipedia, they

suggest removing “these pests” with “boiling water, bleach, or drain cleaner”

(Wikipedia, 2013). In contrast, I found that the moth flies were providing a service -

cleaning up the dead citrus bugs that I forgot about. I imagine that, in their ‘native’

(non-plastic bucket) habitat such as a pond, they may occasionally perform the same

function, cleaning up dead insects - as their 'normal' food is decomposing organic

material. Interestingly, the adults generally do not feed, and most only live for a day

or two (Colless and McAlpine, 1991). I suggest that we leave these moth flies to live

BEES PF DE DT be PE DT be be Pe bd Pe Re bd Pe Pe bd bd

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 19

in their natural habitat, and to keep our drains clean so that there is no need to kill off

these cute little flies 1n the first place.

Although our moth flies appear to be mostly harmless, I should note that one

subfamily of moth flies, the Phleobotominae, do feed as adults and are blood-suckers

(Colless and McAlpine, 1991). Some of these species - found outside of Australia -

transmit diseases, including leishmaniasis.

Australian taxonomic checklist (from Hawking et al. 2009):

Order Diptera — family Psychodidae

Subfamily Bruchomyiinae

Nemapalpus australiensis Alexander

Subfamily Psychodinae

Brunettia 4 species

Clogmia 2 species

Didicrum 2 species

Epacretron pulchrum Duckhouse

Notiocharis 3 species

Paratelmatoscopus 5 species

Peripsychoda 7 species

Psychoda 21 species

Rotundopteryx 15 species (formerly Pericoma)

Telmatoscopus 10 species

Threticus 3 species

Trichopsychoda montana Satchell

Subfamily Sycoracinae

Sycorax australis Duckhouse

Subfamily Trichomyiinae

Trichomyia 19 species

References

Chessman, B.C. (2003). New sensitivity grades for Australian river macroinvertebrates.

Marine and Freshwater Research, Volume 54: 95-103.

Colless, D.H., McAlpine, D.K. (1991). Diptera. In: ‘The Insects of Australia: A

textbook for students and research workers.’ Chapter 39, 2™ Edition, Volume 2.

Melbourne University Press and Cornell University Press. Pp. 717-786.

Hawking, J.H., Smith, L.M., Le Busque, K. (editors) (2009). Identification and

Ecology of Australian Freshwater Invertebrates.

http://www.imdfrc.org.au/bugguide Version January 2009, Accessed 20/01/2013.

Wikipedia (2013). Psychodidae. http://en.wikipedia.org/wiki/Psychodidae

Accessed 20/01/2013

Williams, W.D. (1980). Australian Freshwater Life: The Invertebrates of Australian Inland

Waters. 2" edition. The Macmillian Company of Australia, South Melbourne.

Photos Alisha Steward

PEPE REDS PF OE Pe PE DT De Pe bd be be pe Pe bd Pe be

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 20

Life history notes on the No-brand Grass-yellow, Eurema brigitta

australis (Stoll, 1780) Lepidoptera: Pieridae - Wesley Jenkinson

The No-brand Grass-yellow is encountered in the Northern Territory, and coastal and

sub-coastal regions from north-eastern Queensland into southern New South Wales.

Migration of this species occurs throughout its range depending on regional rainfall,

temperature and suitable availability of its host plants.

In Queensland the species is encountered in a variety of habitats where the host plants

are established. This includes grasslands, woodland, eucalypt open forest and

occasionally in suburban gardens where suitable habitat 1s nearby.

The adults fly close to the ground amongst low growing herbs and grasses. When

disturbed they can fly quite rapidly and can be difficult to follow. During cloudy

conditions they settle on low growing shrubs and ground cover and resume flight

when sunny conditions return. Both sexes occasionally feed from a variety of small

native and introduced flowers.

Whilst in flight, the adults can be very easily confused with

other species of the Eurema genus, particularly E. herla and E.

laeta. Voucher specimens are best for correct identification.

The males of this species do not have a sex brand (as the name

implies). In fresh specimens the adults can be separated from

other Eurema spp. by the presence of two very faint pale

yellow (or pinkish yellow) streaks along the costal margin

towards the forewing apex as pictured. The species also has wet and dry season forms.

The sexes are quite similar in appearance. In comparison to the males, the females are

slightly paler yellow with more extensive black scaling across the wings. The black

margin along the hind wing 1s also generally broken and 1s less extensive towards the

tormmus.

The average wingspan for the adults pictured is 38mm for both sexes.

Eurema brigitta australis (No-brand Grass-yellow)

Images left to right: male, female, male underside, female underside

PEPE REDS PF DE be PE DS pe Pe bs Pe be pe Pe bd Pe bd

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 21

In January 2004 at Beaudesert, in south-eastern Queensland, a female was observed

slowly fluttering around a host plant. Settling on the host plant with her wings closed,

she curled her abdomen onto the upperside of a leaf and oviposited a single egg. This

egg was kept for life history studies. Subsequently the larva was raised on a Fishbone

Cassia (Chamaecrista sp.). The three confirmed host plants for this species are C.

concinna, C. maritima and C. nomame (Moss, 2010).

This egg was pale yellow, spindle shaped with fine longitudinal ribs,

approximately 0.5 mm wide x 1.5 mm high.

Freshly laid egg

When the first instar larva emerged the eggshell was consumed shortly afterwards. It

was observed feeding during daylight hours and resting on the upperside of the host

plant leaves. The later instars rested along stems of the host plant. The larva raised

completed five instars and attained a length of 27mm.

oe |

" tw 4s |

PPPoA hte,

1“ instar larva 2" instar larva

4" instar larva

tithe aol . r | 2 ee Sin Cre fas

Pe OREN STNNE STEUER TTT DP IL : ESET TE DENN PRNIDHE OCP ST WPERS

‘ . ~ to htt

FE et ST ci) UT I eh gh

5" instar larva dorsal view

Pupa lateral view Pupa lateral view

BEDS PED DE De PS Pa Re be Pe De be Oe bd Pe be bd Pd

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 22

In captivity the pupa, measuring 16mm in length, was located below a stem of the

host plant. It was attached with silk by the cremaster and a central girdle. The pupae

occasionally have reddish-brown spots and markings as pictured.

The total time from egg to adult was three weeks, with egg duration of 3 days, larval

duration |1 days and pupal duration of 7 days.

Within the new boundary of the Scenic Rim Regional Shire south of Brisbane, I have

records of adults from November to April and one record for June. In this region the

adults appear to be more numerous during the summer and autumn months. However,

this relates to the timing of local rainfall triggering fresh growth of the host plants.

References

Braby, M.F., 2000. Butterflies of Australia — Their Identification, Biology and Distribution.

vol 1. CSIRO Publishing, Melbourne.

Moss, J.T. 2010. Butterfly Host Plants of south-east Queensland and northern New South

Wales. 3" edition, BOIC.

Photos Wesley Jenkinson

TK OK OK OK OK OK OK OK OK OK OK OK

Getting more pixels - Malcolm Tattersall

In the last Metamorphosis of 2012, | compared image sizes and camera sensor sizes

and concluded that most modern digital cameras will produce images of most insects

which are satisfactory for most purposes. This time I will look at extending our range

downwards, to get bigger images of tiny invertebrates close to us, and 1n the next

issue I will talk about how to get better photos of larger but more distant subjects.

Wedding photographers have it easy: their subjects are all one to two metres tall and

all one to ten metres away. By contrast, our invertebrate subjects can range from a

tiny spider to an adult stick insect, or less than 0.2mm to more than 200mm in length,

and 10cm to 10m or more away. That’s an enormous range to cover - a size ratio of

1000:1 and a distance ratio of 100:1 - and our solutions this time have to be hardware

rather than software.

Small and close

Let’s start with the small, close subjects. Suppose you have seen a 5mm bug sitting on

a leafa metre away. Your photo from that distance with a general-purpose camera and

lens combination will show it as a dot in the middle of the frame (50-1 00px wide in a

S000px image). Obviously, your first decision 1s to move closer, but there are two

kinds of limits:

(1) The bug may take fright and jump away. Let’s call this its startle distance. If the

bug doesn’t take fright, you keep moving in until ...

BEDS REDS PF DE bs PE De De Pe be Pe be pe Pe bd Pe bg

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 23

(2) The camera can’t focus because it 1s too close to the subject, 1.e. closer than its

minimum focus distance. This will typically be 25—50cm from the sensor to the

subject with everyday lenses. The distance from the front of the lens to the

subject is called the working distance, and is another useful figure to keep 1n

mind.

At the minimum focus distance you are at the point of maximum magnification of the

lens and the image 1s as big as possible with your set-up. Maximum magnification 1s

usually between 0.13 and 0.30. That is, the image on your sensor is 0.13 — 0.30 times

the size of your subject, with 0.20 being about average. A rough calculation tells us —

or a bit of experimenting shows us — that our 5mm bug becomes a 200px image.

That’s still not much use, so what can we do?

If you have a camera which doesn’t take interchangeable lenses, you may have to go

shopping for a new camera. If you don’t want to go all the way to the price and bulk

of a SLR, then look at point-and-click cameras with a large optical zoom range or the

newish bridge category (http://en.wikipedia.org/wiki/Digital_camera will introduce

you to the differences). However, some point-and-click cameras do accept close-up

filters (see below) or clip-on macro lenses, so before you go too far with your

research, ask about accessories for your present camera at your camera shop. Or you

may be able to improvise, like an online friend of mine who gets great macro shots

with a Panasonic DMC-FZ50. It has a 10x optical zoom to start with but he has

extended its capabilities by fitting a philatelic magnifying glass into the frame of an

old UV filter to make a +10 close-up filter — see

http://www.flickr.com/photos/72842252(@N04/ for some results.

This is an appropriate time to mention the difference between optical zoom and digital

zoom. The former 1s a function of the lens and changes the size of the image on the

sensor, and is extremely useful. The latter 1s a software function exactly analogous to

cropping the image on your computer at home and, on the camera, it does nothing

much except fool you into thinking you have a bigger image. See

http://www.cambridgeincolour.com/tutorials/image-interpolation.htm for more on this

subject.

If you have a SLR, then a macro lens 1s the best solution. Macro lenses should have a

maximum magnification of 1.0 and Canon’s three macros, of 60, 100 and 180mm,

achieve that at their minimum focus distances of 0.2, 0.31 and 0.48m respectively.

(This neatly illustrates the advantage of the longer lenses: you have less need to get

within the startle distance. However, longer lenses are affected more by camera shake,

so image stabilisation becomes particularly useful.) Some manufacturers’ macros may

have less magnification, e.g. 0.5, but will still provide a bigger image than a non-

macro lens.

BEDS PE DS PF DE be PE De pe Pe bg Pe be be Pe bd Pe bg

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 24

Orange Bush-brown, Mycalesis

terminus, and cake, cropped from

3888 x 2592px to 1240 x 1240 for

effect. Canon EOS 1000D, 100mm

macro lens without extension tubes

or CU filter. The butterfly was

enjoying the Christmas cake so much

that it let me get right down to

minimum focus distance and take a

dozen shots, even with the flash.

Alternatively, we can put a close-up filter on the front of a normal lens or insert

extension tubes between the lens and the camera body to increase its maximum

magnification. We can even do both, to achieve a maximum magnification up to

perhaps 1.10. But be warned: they only offer their benefit at very short working

distances, so many bugs will move off because you are too close. Also, they prevent

the lens from focusing at long distances so you need to remove them for normal

shooting.

Paper wasps, Ropalidia revolutionalis (about 8mm long), on nest. Canon EOS

600D, 15-85mm zoom lens at 85mm with a +2 close-up filter; 5184 x 3456px

cropped to 2100 x 2800 to show wasps on nest, then cropped again to 480 x 600px

to show (without taking up a whole page of the magazine) the level of detail

achieved. A +4 close-up filter would have given me more magnification but I may

have been stung for it.

BT PS PERS DE DE be Pe Re be Pe De be Oe bg Pe be bd Pe

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 25

Golden Mosquito,

Coquillettidia xanthogaster.

Canon 600D, 100mm macro

lens with +2 CU filter, 5184 x

3456px cropped to 1330 x

1000.

Finally, we can use close-up filters

and/or extension tubes with a macro

lens for a maximum magnification of

1.5 or a little more. With any of this equipment, our 5mm bug is 800px or more wide

— avast improvement on the 200px we started with — so we can achieve satisfactory

shots of close subjects as small as 3-5mm (jumping spiders, small flies and the like)

and great detail with 10-20mm subjects. But other technical limits already begin to

intrude in the sub-10mm range and more specialised gear is needed for the tiniest

critters so I will stop here.

Photos Malcolm Tattersall

TK OK OK oR oR OK OK OK OK OK

Field Notes: Major extensions to the known distribution of the

Bright Purple Azure, Ogyris barnardi (Miskin 1890) in Queensland

(Lepidoptera: Lycaenidae) - Kelvyn L. Dunn

E-mail: kelvyn_ dunn@yahoo.com

During spring of 2011 and 2012, I investigated the distribution of the Bright Purple

Azure butterfly, Ogyris barnardi in Queensland, 1n order to confirm the westernmost

limits. I found that 1t extended much farther inland than currently believed. Prior to

this investigation there had been relatively little survey of the butterfly fauna of

western Queensland (see Dunn & Franklin 2010, online appendix), which by

corollary raises the question as to the completeness of the limited distribution

historically attributed to this species. Nonetheless, over many decades keen

enthusiasts have regularly sought Ogyris (as a group) by determined searching, a

selective process which might offset that inland knowledge deficit (for butterflies in

general) to an extent. Indeed, given their general scarcity (and hard work required to

obtain them) enhanced too by their brilliant hues, Ogyris butterflies have earned a

status as gems among the Australian lycaenidae, with many rare species coveted and

those reared for the cabinet proudly treasured. That the species, O. barnardi, was

actually of limited and disjunct distribution in northern Australia, as generally

thought, had hinged on published knowledge: Braby (2000) earlier presented a

BEDS PERT PE DE PS Pa Re be Pe De be Oe bd Pe be bd be

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 26

synoptic range-fill map based on available records in museums and literature sources

up to that time. All contributory factors considered then, the synoptic depiction was

supposed to be reasonably accurate and thus likely inclusive of most of its

distribution, approximating to the outer western limit of its occurrence.

Ogyris barnardi (Figure 1) inhabits mixed woodlands where the Grey Mistletoe,

Amyema quandang (Loranthaceae) on which its larvae regularly feed, parasitises

particular species of wattles (Acacia) (see Braby 2000 and references therein). The

usual means of obtaining specimens of this uncommonly seen butterfly has been to

rear adults from the juvenile stages (usually older larvae or pupae, found secreted in

borer holes or under loose bark). Determined searching to this end has been

responsible for most museum specimens collected to date and so less information is

available about its seasonal flight period and the times of day that adults are active in

those areas where it occurs. Encounters with the high-flying adults are often fleeting

(making field identifications very difficult at best) and the regularity of their sighting

in the field is a little unpredictable; on the balance then, those adults netted

opportunistically are often in poor condition and so rendering them less suitable for

retention. For this reason too, field sampling of adults 1s an infrequent practice as the

lengthy time spent and labouring efforts made 1n the physically trying outback

climate, where it lives, are not particularly rewarding to collectors.

In October and November, during the butterfly’s spring appearance I systematically

sampled mixed woodlands for evidence of the species whilst en route to the Gulf of

Carpentaria in northern Australia, where I planned to seek out one or more other

species. The butterfly is not readily found by random sampling: I examined numerous

sites for various butterflies (not just this species) on my two trips (Figure 2), and of

those inspected (Table 1), 22 locations (a rather small proportion by comparison)

provided evidence of adults of O. barnardi. Where achievable, capture of one or more

individuals was the means to confirm field identifications with certainty. That

evidence obtained (Table 1) suggests that O. barnardi is widespread 1n the inland,

albeit rather patchy 1n occurrence and confined to, and localised within, suitable

habitat in western, central and northern Queensland. Indeed, the butterfly could

extend farther west into the Northern Territory where Grey Mistletoe-infested

woodland patches exist, and so, the remote border area could be the focus of those

who wish to expand upon my study.

The butterfly looks drab in flight and, at such times, can be confused easily with other

members of that genus. In general appearance, it closely resembles the Broad-

margined Azure, O. olane, a species that 1s similarly sized and coloured, but one that

seems absent from the acacia-dominated woodlands 1n the outback where O. barnardi

exists. Importantly, O. olane does not utilise the Grey Mistletoe as a larval host in the

wild, although a captive larva, proffered foliage, consumed this 1n one trial (Dunn

1997) and so an ancestral tolerance of a broader diet remains. Indeed, current

evidence suggests the two species are host (and habitat) segregated, an adaptive

BEDS PE DS PF OE be PED pe PE bg Pe be pe Pe bd Pe bg

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 27

strategy that reduces competition for resources and one that probably drove their

speciation eons ago. Yet, this divergence from its closest ally does not assure sole use

of the larval host by that particular member of the genus. In fact, there 1s another

species of Ogyris, but one more distantly related (unlike these two species, it has a

differently shaped antennal club), that shares the same woodland habitat with O.

barnardi and, at times, shares too that same mistletoe species as a host (but perhaps

not the same individual plant). Hence, in some places in western Queensland, the

species 1s sympatric with the Satin Azure, O. amaryllis, and where this situation

arises, 1t represents an additional factor that confounds field identifications, short of

handling. The latter species is a more widespread butterfly (its broader host range has

enabled this) and one that looks very similar in flight (when in silhouette), albeit often

(but not always) slightly larger. Obviously, the glistening azure blue upper-wings of

O. amaryllis readily distinguish it from others but this telltale sign 1s rarely seen from

below (the observers’ usual position), and small males, when perched in shade with

closed wings (the usual stance adopted), could easily be mistaken for an O. barnardi

on prima facie evidence.

A helpful clue 1s the fact that adults of O. barnardi differ slightly in behaviour

compared with O. amaryllis, which the trained eye may detect. They tend to patrol

more locally and tenaciously perch on older wattle trees — especially favoured are

those with many dead branches projecting into the canopy. Disturbing these may

reveal a settled male, which otherwise could remain undetected during a passing

inspection of habitat. From these high vantage points, the males dart out briefly to

defend an aerial flight space, particularly one that overlooks one or more larval host

plants. At such times, they may rapidly encircle those Grey Mistletoes growing

nearby to seek out newly emerged females or to intercept visiting females inspecting

the larval hosts for egg laying purposes and, particularly at certain times of day — late

afternoon seems favoured — may pause to feed briefly at the mistletoe’s flowers (when

available and nectar enriched). During feeding bouts, which seemed loosely

synchronised, I observed that several adults 1n succession would frequent one or two

individual flowers, leaving other flowers close by on the same plant unattended. Much

of the time though, adults did not feed but perched for lengthy periods, or routinely

patrolled their haunts when intruding adults strayed into their airspace. At such times,

a flurry of activity involving three or more adults could result and which often drew

my attention to the presence of members of the genus at a location.

Provision of high quality identifications 1s paramount for rigour, yet at some

locations, all adults evaded capture. Common reasons for this were (1) their sustained

height in the canopy, (2) their rapid and evasive flight, and (3) the regularity of

sightings at individual locations. In particular, a low abundance at the time or hour of

my visit(s) sometimes made encounters infrequent and reduced opportunities to net

passing adults. At the best of times, when adults were common and opportunities to

net were duly increased, the quantity of snags (dead branches) on the older wattles

BEDS PE DS PF DE be PE DT De Pe bs Pe be pe Pe bd Pe bg

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 28

(the butterfly particularly favours these for perch sites) and festoons of creeping

Capparis, equipped with net-hitching spines, often hindered the successful

manoeuvring of a deep hoop-net attached to long poles. Those who have attempted to

capture adults of Ogyris on a regular basis (not just occasionally) will recall that this

is a time consuming task and, indeed, often a difficult one — particularly during

midday hours when the sun is directly overhead — hence, an expectation of success at

all sites 1s unrealistic. The inclusion of observations then (see Table 1) may raise

doubt in the minds of some, as the species in this group can look very similar to the

novice or advanced observer, even when equipped with field glasses. Nonetheless, I

am experienced with this and other related species in Queensland, largely from survey

work conducted on the Darling Downs as part of employment in the early 1990s (see

fieldwork detailed in Dunn & Kitching 1994). On this accreditation, those field

identifications deemed reliable comprise ‘expert opinion’ which is categorically

‘almost certain’ and a legitimate (albeit arguable) substitute, short of capture. These

are marked “Obs-only’ to distinguish them from those verified (evidence-based)

records (captures); I appreciate too that others may wish to gather more evidence at

those sites concerned (where specimens were not obtained) and would encourage this.

The 22 locations where I found the species on my two most recent field trips are listed

from north to south (Table 1); those marked with an asterisk (*) are considerably

beyond the range indicated by Braby (2000) and so now enrich the knowledge base.

All distances were measures by road from the Post Office of the nearest township, and

were calculated by vehicle odometer usually from the closest road marker (where

these were available) to reduce instrumental error, and so are considered precise but

variably accurate. GPS coordinates (and elevations) were obtained at each site with a

hand-held trekking device, and the measured road distances to each were checked

later on Google Earth (www.google.com/earth/index.html) to ensure agreement (that

is, to within a kilometre) — there was minor disagreement for some though where

large road distances were involved. Most locations were widely separated, often by

hundreds of kilometres, but occasionally juxtaposed sites (variably within one

kilometre of the other) received attention as well. For each of the juxtaposed locations

the habitat was continuous and likely, the butterfly too. The GPS coordinates to the

nearest minute are included (Table |), but in all cases, the specimens are labelled

more finely, whether that is to seconds or to one decimal place of a minute, as

accuracy needs for each location dictated. Finally, the elevations inhabited by the

butterfly ranged from about 70m to 540m, and all specimens examined corresponded

with the geographically closest populations, namely the nominate subspecies to the

east.

References

Braby, M.F. 2000. Butterflies of Australia: their identification, biology and distribution.

CSIRO Publishing, Collingwood Vic.

Dunn, K.L. 1997. Biological notes on some eastern Australian Butterflies, Part II. Victorian

Entomologist 27(6): 114-118.

BEDS PADS PF OE be PE DS pe Pe bs Pe be pe Pe bd be bg

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 29

Dunn, K.L & Franklin, D.C. 2010. Exploring the adequacy of representation of butterfly

species’ distributions in a more accessible portion of northern Australia. Northern

Territory Naturalist 22: 88-94 (& online appendix:

http://sites.google.com/site/ntfieldnaturalists/journal ).

Dunn, K. L. & Kitching, R. L. 1994. Distribution, status and management of the Piceatus Jewel

butterfly on the Darling Downs, Queensland. A report to the Queensland Department

of Environment and Heritage, Conservation Strategy Branch. (May 1994). 46 pp +

tables and appendices.

Figure 1. O. barnardi male at mistletoe flowers of Amyema quandang, 25km NW

of Tambo, 02 Oct 2012. The arrangement of the eight black-edged, often smoky

brown-tinged segments of the post-median band (which extends from the dorsum to

the costa of the hind-wing, and which comprises five joined segments, with one

(variably disjunct) segment at the dorsum and two more-or-less upright (disjunct)

segments towards the costa) suggests the species involved. (In contrast, three or four

slanted markings, situated increasingly distally along the hind-wing costa towards the

apex, more abruptly end the continuation of the five joined segments of the post-

median band in O. olane, the species it 1s most similar to). Although I offer this means

as a useful guideline to help observers identify this species from field-gained

photographs (and with a good probability of correctness too), I remind that the

subsequent capture and examination of this adult confirmed its identification (K LDC).

Photo Kelvyn Dunn

PEPE PERT DE OE PS Pe Re be Pe he be Oe bd Pe be bd Pe

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 30

(AJUO SQQ) M34

AURA

(AJUO SQQ) 2U0D

AUR

MO]

(AJUO SQQ) AURA

AURIA]

(AJUO SQQ) 90

alt @)

(AJUO SQQ) UO

ouO

(AJUO SQQ) MOH

MO.

(AJUO SqQ) 9UQ

(AJUO SQQ) 9UD

AURTA

MOJ

alte)

(ATUO SqQ) Mo

MO]

(AJUO SqQ) Md]

AURTA]

sduBpuNngy

YOCEL-OFCI

YOLTL-OLol

YOCOT-O¢S I

YUSe9l-OC9T

YOSE T-OScl

USCFL-OOFI

YOLTT-0¢60

YOOCL-O¢O0!

YOCLeL-O0L1

USSFI-StFl

USFII-SIILI

USFII-SITI

YOOS L-O0F I

YOLTT-OOLT

USLOl-SPsl

YUSOOT-SS60

USTEL-O0LT

YOO’ [-00CI

Used I-ss9]

YOse tO!

YOF9OT-OFS |

USTFIL-OLF I

YOO9T-S cS I

YOOCI-S cI

YORCL-OCI

YSFOL-OS60

(LSAV)

oul |

[10¢ AON FI

CLOC WO CO

CLO? WO 10

CLO0C WO [0

CLOC PO CO

CLOT BO CO

[10¢ PO 92

CLO~ BO £0

C10C WO £0

[1L0@ PO 97

[LO¢ AON OL

L10c AON OT

[TOC AON 60

ClOC PO 8C

[10 SON FO

L10c AON CO

I10c AON CO

CLOT PO 61

CLOC WO 07

CLOC PO 61

CLO~ PO VI

CLOT PO OI

CLO? WO 07

CLO WO 0C

CLOT PO 07

CLOT PO 97

aug

x + & *

+ + £€ & HH &

(tu) *AoTy

‘xoiddy

UONBVULIOJUL pIzJBIIOSse pu puLjsudINe) UI

A.8606F1 S.8FoST

A.9TIFL SS. T00ST

A.9TIFT “S.1T0087

A.9009F LS. SPoF TC

A.909F LS. S FoF 7Z

ACSF S.LTorC

AStottl S.9loFC

ALGSPL S.10orC

A.9FoLFEL S.EL0kC

A.PPoLFL S. Lote

A.flof FL S.9S007

A.Leoh Pl S.C 007

A.fLoS PL Sit hoOC

A.6£06E1 “S. TFT

A.1LO6L1T S.Flo0C

A.LSo8ET S.FT0C

A.C LOFL S$. £0007

A.LEoRECT “S.0000T

A.FEOET “S.00007

A.OLo8ET “S.6L061

A.TLo8ET S.8f061

4.0C0T1 S$. 1.61

9p0d095y

WIOOIR T, JO S AG MS ULYRZ

OquIe |,

Jo JS urycZ °.dduei uo jutod ysoysiH,

Pa UONEIS

SOYURAT Je “OQUIR], JO FS WYPZ

OquUIET JO MAN UDIST

OQUIP | JO MN UD?

eyoryg JO S Aq A UrygT

PIOJSIS] JO FJ WAGE

eye JO A\ AQ N uryre

oOLyeuy JO 4 Ure

Py yoouyH ye “omyddes jo s wyy

uopusysny

JO FAQ § UAT] “Peoy 1OyTPA\ IW

SILI JON Aq J WYZ

INOYOO'] IsukY BLING JO ASA UD]

US] YA JO Ff OUYQ] “YOOID 9d10D

BST WAN JO AAN WDLPS8

LS]

JUNOWY JO MN WYNG “OAR Aoppng

ALINDUOTD JO ANN UNTER

jROMOOUIR JO S Ag FWY

[BOMOOUIB) JO S AG FY UWYAR

JLOMOOWIR) JO FN UlyLS

[POMOOUIBD JO FN URS

ISNOYPROY SILA

W WANG ASS WYO'E “YIU [OZPH

udIeI0'T

IPADUADG “(—) AOY SUOTFYIO] AOYING 77 IY "[ AIQuy

PEPE PADS Pe DE PS Pe he be Pe he bd Oe bd pe be bd Pe?

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 31

Figure 2. Author’s butterfly survey sites in Queensland, spring 2011 and 2012

KLD Survey Sites #4011 & #2012

~ Ales stick depne | ) T:

BOOK REVIEWS

McCormack, Robert B. 2012. A guide to

~ Australia's spiny freshwater crayfish. CSIRO

| Publishing, Collingwood. Octavo, paperback, 235

pp. colour photographs, line drawings.

$59.95 from the publisher.

Reviewed by Jonathan Marshall

eel «L have long held a fascination and admiration for freshwater

crayfish, particularly spiny crayfish from the genus Euastacus

covered by Robert McCormack’s new book. Australia is a

hot-spot for world freshwater crayfish biodiversity and yet this diverse and endemic

fauna 1s not recognised or appreciated by most Australians. The main reason for this,

I am sure, 1s that there has been a conspicuous absence of natural history books about

these animals to inform and intrigue the nation’s many nature lovers. Until now,

BEDS PED Pe DE PS Pa Re be Pe De bd Oe bs Pe be bd Pd

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 32

much knowledge about Australian crayfish has been locked away in dry taxonomic

papers in scientific journals, inaccessible and uninteresting to most outside a small

community of freshwater scientists and amateur enthusiasts. This excellent new book

from CSIRO Publishing makes a great leap forward towards introducing these

beautiful and significant invertebrates to Australians and in doing so, I hope, igniting

community conscience for the need to better conserve this unique biodiversity 1n the

face of the multiple threats it currently faces.

Euastacus 1s the most diverse and perhaps most interesting Australian crayfish genus,

with 50 described and at least four undescribed species ranging along the east coast

from Cooktown in north Queensland to Wilsons Promontory in the south of Victoria.

Some species are very large and conspicuous: at up to 3 kg E. armatus from the

Murray River and southern tributaries is the largest, and E. hystricosus from the

Maleny/Conondale Ranges region 1s Queensland’s largest, growing to 1.2 kg (Figure

1). Many are also very colourful (Figure 2), with different species sporting fetching

combinations of red, green, brown, black, white and blue, with variously developed

spines and tubercles, often in contrasting hues. Others are small, inconspicuous and

much less likely to be casually encountered. Most species have restricted, to very

limited distributions, and very narrow habitat requirements. These characteristics

make Euastacus a fine choice as a subject for a guide book such as this, which covers

their anatomy, life history, ecology, identification, distribution and threats.

a —o

~~. 4

— ™

Figure 1. This Euastacus hystricosus from a stream in the Conondale Ranges is a member of

Queensland’s largest crayfish species and is threatened by illegal fishing. The spiny

morphology characteristic of many members of the genus is clearly evident.

(photo by Jonathan Marshall).

Book author Robert McCormack has over 20 years’ experience as a commercial

crayfish farmer in Australia and is secretary of the NSW Aquaculture Association, 1s a

research associate with the Carnegie Museum in Pittsburgh, USA, and is prominent

within the Australian Crayfish Project, which conducts biological studies of

Australian waterways, collecting and identifying crayfish. Robert’s passion for

BEDS PE DS PF OE bs PE DS De Pe bd Pe be be Pe bd be be

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 33

freshwater crayfish 1s clearly evident from the enthusiasm of his writing in this book

and his sharp observations of the biology and ecology of the various Euastacus

species discussed.

Figure 2. Euastacus sulcatus from a stream in Springbrook National Park is perhaps the best

known, and most colourful, of the Queensland Euastacus. This is a diurnally active species and

often leaves the water to forage amongst moist leaf litter on the rainforest floor, where it loudly

and aggressively hisses if disturbed. (photo by Jonathan Marshall)

The A5 sized, soft covered book 1s produced to a high standard, as one expects from

this publisher. The many maps, figures and tables are clear and accurate. The

numerous photographs of crayfish and their habitats are of a high standard and are

clearly reproduced with accurate colour and at a size appropriate to their content.

The book aims to present the most up to date information on Evastacus taxonomy,

identification, distribution and biology with the hope that increased interest and

awareness will promote better conservation and management. It commences by

giving a good overview of the necessary background to systematics, morphology and

the general biology of the genus. There is a very nice table listing all 137 described

Australian crayfish species in 10 genera with their distributions, plus a web site to

check for new additions. This latter resource is very useful as new species are still

being described — including several from the Queensland Whitsunday region just this

year.

Euastacus species are then classified into three groups, which (although not described

with this ecological term 1n the book) could be considered as guilds of species sharing

broad morphological and ecological characteristics. This divide into ‘giant spiny

Euastacus ’, intermediate Euastacus’ and ‘dwarf Euastacus’ is made by the author

pragmatically and for the purposes of the book. The rationale and justification for this

erouping are quite well argued, but I was disappointed that no reference 1s made to the

considerable information available on evolutionary relationships between Euastacus

species derived from population genetics studies. It seems to me to be of great

relevance to conservation management to know 1f the morphological and ecological

BEDS REDS PF OE be PE DT De PE bs Pe be pe Pe bd Pe be

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 34

guilds recognised by the author represent distinct lineages in the evolution of the

genus. Despite this, should the reader find a specimen to identify, the guild approach

does without doubt assist greatly with the task. This is because of the fascinating

observation by the author that within a single stream system, there can often be one

species from each guild present, each occupying distinct ecological niches according

to the guild it belongs to and its life-stage, but there are rarely multiple species present

from the same guild. I am particularly interested in these observations because they

infer a strong case for niche partitioning by competitive exclusion and differential

environmental tolerances. These processes are common 1n some environments, such

as rocky sea shores for example, but have rarely been demonstrated in stream

ecosystems. The author’s observations provide clear hypotheses ideal for scientific

evaluation. PhD project, anyone?

The high species diversity of Evastacus 1s to a large extent a consequence of two

phenomena: firstly, the physiological requirements of the genus for cool, moist

environmental conditions, and secondly, the process of gradual aridification of the

Australian continent over the past 40 million years. Suitable habitat for Euvastacus has

steadily contracted over this time and the genus 1s in fact a relic of the cool, wet

Gondwanan forests that once covered our continent. For approximately the past five

million years Evastacus populations derived from several widespread ancestors

(perhaps representing the author’s three guilds?) have been isolated in relic patches of

cool wet conditions, usually on mountain tops along the east coast, and this has

resulted in vicariant speciation to produce the many species we see today. Once again

to my disappointment, this fascinating aspect of Euastacus biology 1s little mentioned

in the book, and the several published studies which have presented this story are not

cited (see further reading at the end of this review).

The main section of the book presents at least one nice colour photograph and a

comprehensive description of each species, arranged in alphabetical order and for

each including their habitat/activities, diagnosis, distribution, colour, breeding,

conservation status and comments. An additional table and regional maps outline

known species distributions. There is no taxonomic key presented, so species

identification using the book relies on knowing where a specimen was collected and

which of the three guilds it belongs to. Considering that the information necessary to

generate a key is included in the species diagnoses, it 1s a missed opportunity that

none was created for the book.

The penultimate chapter deals with threats to the viability of Euastacus populations.

All Queensland species are protected from ‘fishing’ by the Fisheries Management Act

but illegal harvesting is a major threat, especially to larger individuals of the giant

species, pushing some close to extinction. According to the author, even some

National Park Rangers are unaware that Euastacus in their park are protected by law!

Let’s hope they look at a copy of this book to put them straight. Climate change 1s an

insidious threat to all species with limited capacity for dispersal and requirements for

cool, moist conditions, with Euastacus particularly susceptible. Populations restricted

BEDS REDS PF RE be PE DS De Pe bd Pe be pe Pe bd Pe bg

Magazine of the Butterfly and Other Invertebrates Club #68 — Page 35

to mountain tops have nowhere to go 1f temperatures rise, but the author offers some

hope for burrowing species as they may be able to retreat to their burrows as refuges

from extreme temperatures. Habitat alteration is another broad threat, but fortunately

many Queensland species are sheltered from this within National Parks. Anecdotal

evidence suggests feral pigs actively seek Euastacus for food and indeed I have seen

evidence of pig rooting in the vicinity of crayfish burrows within National Parks.

Finally there is a section on diseases and ectocommensals, including the cute little

flatworms called temnocephala, which live on the exoskeleton of the crayfish without

doing much harm and look like miniature inflated rubber gloves. Different Evastacus

host unique temnocephala species, and they too are being studied to help understand

the evolution of the Australian aquatic biota.

Overall, I very much like and recommend this book to anyone interested in Australian

natural history, despite the few criticisms I have made. It provides a very good, if not

comprehensive, overview of what is known of the genus and presents much that is

new based on the keen observations of the author and his colleagues. It should be a

valuable resource to students and researchers and I would be delighted to see

ecological studies to formally evaluate some of the important anecdotal observations

made by the author. Will it fulfill its ambition to improve Euastacus crayfish

conservation and management? It 1s difficult to say, but as it 1s well presented, looks

pleasant and includes ample information to encourage such an outcome, it does seem

possible. Let us hope the relevant managers and policy makers take note, recognise

the need and act to protect our beautiful and scientifically valuable Euvastacus spiny

crayfish.

Finally, I would like to thank CSIRO Publishing for kindly providing a copy of this

book for review.

Ed.: A preliminary key by Robert McCormack is now available and was presented on the

6/02/2013 at the taxonomic workshop on aquatic invertebrates held at La Trobe University,

Albury-Wodonga Campus.