THE AUSTRALIAN

Entomologist

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THE ENTOMOLOGICAL SOCIETY OF QUEENSLAND

Volume 34, Part 4, 10 December 2007

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ISSN 1320 6133

THE AUSTRALIAN ENTOMOLOGIST

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Cover: Parobia alipilus Seeman & Nahrung (Podapolipidae) is one of three sexually-

transmitted mites that infest the eucalyptus leaf beetle Paropsis atomaria. The adult

male (pictured) is unlike the female: his legs have spurs, the genital capsule is mid-

dorsal and the fourth pair of legs is modified. Nevertheless, Parobia species are

among the most mite-like of the Podapolipidae, where physogastry and loss of legs is

common and some males have their genitalia on prongs above their heads. Parobia

mites are often more common on female beetles and at least one species significantly

reduces survival of the host during overwintering. Illustration by Owen Seeman.

Australian Entomologist, 2007, 34 (4): 97-106 97

DISTINGUISHING BETWEEN LYNX SPIDERS (ARANEAE:

OXYOPIDAE) RELEVANT TO IPM IN COTTON IN THE NAMOI

VALLEY, NEW SOUTH WALES

M.E.A. WHITEHOUSE! and J.F. GRIMSHAW?

'CSIRO Entomology, Australian Cotton Research Institute, Locked Bag 59, Narrrabri,

NSW 2390 (Email: mary.whitehouse(Qcsiro.au)

?PO Box 1356, Toowong, Old 4066 (Email: grimshawjudy@hotmail.com)

Abstract

Characters are presented to enable identification of three similar species of lynx spiders

(Oxyopes molarius L. Koch, Oxyopes amoenus L. Koch and Oxyopes gracilipes (White))

commonly found in cotton in northern New South Wales. The aim is to enable non-specialists to

readily distinguish between these species so that they can be better incorporated within an

Integrated Pest Management (IPM) framework.

Introduction

The advent of transgenic Bt cotton has had a major impact on the cotton

industry by largely controlling lepidopteran pests, with little effect on the rest

of the insect community (Whitehouse ef al. 2005, Hagerty et al. 2005). This

has made cotton more amenable to Integrated Pest Management (IPM)

strategies (Fitt and Wilson 2000, Deutscher et al. 2005) and has led to a drop

in the amount of insecticide used on cotton (Fitt 2004), which has had major

benefits to the community.

However, one consequence of reduced insecticide use is that minor pests, that

were once controlled by sprays for Helicoverpa Hardwick spp. (Lepidoptera:

Noctuidae), have become major pests in cotton. One such group of pests are

the green and brown mirids Creontiades dilutus (Stål) and Creontiades

pacificus (Stål) (Hemiptera: Miridae) respectively. In Bollgard” II cotton, the

need to control mirids and other sucking pests, such as the green vegetable

bug Nezara viridula (Linnaeus) (Hemiptera: Pentatomidae), tends to

undermine the gains in reduced insecticide use that came with Bt cotton. This

is because the broad-spectrum insecticides used to control these pests could

disrupt the beneficial arthropods and increase the risk of secondary pest

outbreaks such as mites and aphids.

One means of controlling mirids without resorting to insecticide spray, is to

maintain or increase the numbers of predators in cotton that are likely to

attack them. One of the main groups of predators in cotton are spiders, which

make up 50% of the beneficial arthropod community in unsprayed cotton

crops in Australia (Bishop and Blood 1981). Although little is known about

the exact role that spiders play in the management of pests in cotton, it is

known that lynx spiders (Oxyopidae) are mirid predators (Bishop and Blood

1981, Young and Lockley 1986, Nyffeler et al. 1992). They are also the most

abundant family of spiders in Australian cotton (Whitehouse et al. in press).

The dominance of lynx spiders in Australian cotton fields indicates that this

family could be effective against mirids.

98 Australian Entomologist, 2007, 34 (4)

In order to utilize specific oxyopid species within an IPM framework for the

control of pests, there is a need to be able to distinguish between species.

However, as is the case with many Australian spider families, there are no

recent revisions of the Oxyopidae. The last published work was a catalogue

(Roewer 1954) that listed 14 Australian species, 12 of which were in the

genus Oxyopes Latreille. Since then only a few papers have been published.

Grimshaw (1989) reported the first record of another genus, Hamataliwa

Keyserling, in Australia. Vink and Sirvid (1998, 2000) provided reports on O.

gracilipes (White), but identification of specimens from their work was not

possible. Townsend er al. (2001) provided good descriptions of some

Australian Oxyopidae, but from that work it is still not possible to

confidently identify a specimen at hand. To date there is only one revisional

work on Australian Oxyopidae, an unpublished thesis by Grimshaw (1991).

The aim of this paper is to provide simple tools to differentiate between the

three species of Oxyopidae commonly found in cotton in the Namoi Valley

of northern New South Wales. By aiding species identification of these

spiders, we hope to facilitate their use within an IPM framework for the

control of insect pests.

Materials and methods

Oxyopidae were collected using beatsheets (Mansfield et a/. 2006) or visual

searches throughout cotton growing seasons at the Australian Cotton

Research Institute (30°11’S, 149°33’E) at Narrabri, NSW. During the

2006/07 cotton growing season, samples of 10 beatsheets on 10 sampling

dates were taken throughout the season. All lynx spiders in the samples were

sorted to species. Three species were found (Figs 1-8): Oxyopes molarius L.

Koch (‘plain brown lynx’), Oxyopes amoenus L. Koch (‘banded lynx’) and

Oxyopes gracilipes (White) (‘stocking lynx’).

Adult specimens were identified using the key to species in Grimshaw (1991)

and through comparisons with specimens held at the Queensland Museum,

which had been verified by comparison with type material held in Hamburg,

Germany. Voucher specimens of each species are lodged with the

Queensland Museum (female O. molarius: QMS 83346; male O. molarius:

QMS 83278; female O. amoenus: QMS 83277; male O. amoenus: QMS

83276; female O. gracilipes: QMS 83347; male O. gracilipes: QMS 83279).

Results

Female genitalia

The external genitalia of females of the three species are easy to differentiate

(Figs 9-11). O. molarius has a median septum with a broad longitudinal

process, while O. amoenus has a median septum with a narrow and attenuate

longitudinal process. O. gracilipes has a broad median septum with no

processes.

Australian Entomologist, 2007, 34 (4) 99

Figs 1-8. Photographs of female (left) and male (right) Oxyopes spp. commonly found

in cotton in the Lower Namoi Valley, NSW. (1-2): O. molarius (pale morph); (3-4):

O. molarius (dark morph); (5-6): O. amoenus; (7-8): O. gracilipes. Note the bands on

the legs of O. amoenus (Figs 5-6), and the longitudinal stripe on the femur of O.

gracilipes (Figs 7-8). The dark morph in O. molarius can be produced by the loss of

the light hair covering from the pale morph.

100 Australian Entomologist, 2007, 34 (4)

Figs 9-11. Epigyna of O. molarius (9), O. amoenus (10) and O. gracilipes (11),

showing differences in the median septum, which extends anteriorly in O. amoenus

(arrowed).

Male genitalia

Distinguishing between the palps of O. gracilipes and that of the other two

species is relatively easy; O. gracilipes has a small, triangular cymbial

apophysis (Fig. 13) instead of the large, distinctively hooked cymbial

apophysis of O. amoenus and O. molarius (Figs 16 and 19 respectively,

arrow a). However, distinguishing between males of O. molarius and O.

amoenus is difficult. The only differences we found between the palps of

these two species were the shape of the hook (Figs 16 and 19, arrow a; the

hook is longer and more curved in O. molarius) and the shape of a second

cymbial apophysis, which is a small mound in O. molarius and bears a

thickened proximal border resembling a hook in O. amoenus (Figs 16 and 19,

arrow b).

Australian Entomologist, 2007, 34 (4): 101

Figs 12-20. Male palps of O. gracilipes (12: lateral; 13: dorsal; 14: ventral view), O.

amoenus (15: lateral; 16: dorsal; 17: ventral view) and O. molarius (18: lateral; 19:

dorsal; 20: ventral view), showing the hooked cymbial apophysis (arrow a) in O.

molarius and O. amoenus. In O. molarius the hook is longer and more curved, while

in O. amoenus a second cymbial apophysis (arrow b) shows a thickened proximal

boarder resembling a hook. Arrow c = retrolateral tibial apophysis.

102 Australian Entomologist, 2007, 34 (4)

Males of all three species have ventral tibial apophyses (opposite the

thickened border: arrow b in Figs 16 and 19). Townsend et al. (2001)

identified a retrolateral tibial apophysis that was present on O. gracilipes and

O. molarius but missing on O. amoenus. In our observations, the structure is

clear on O. gracilipes, while both O. molarius and O. amoenus have a slight

retrolateral tibial apophysis (Figs 14, 17 and 20, arrow c).

In general appearance, O. gracilipes is sexually dimorphic while O. molarius

and O. amoenus are not (Figs 2, 4, 6 and 8). The cephalothorax of some male

O. gracilipes had an underlying orange hue.

Figs 21-23. Markings on leg I of O. molarius (21), O. amoenus (22) and O. gracilipes

(23). O. gracilipes has is single, sharp black stripe; O. molarius has either no stripe or

one or two broader and more diffuse stripes; O. amoenus has no longitudinal stripes

but has transverse bands around the patellae and tibiae (arrowed).

Australian Entomologist, 2007, 34 (4) 103

Legs

All three species are very variable in their abdominal pattern due to abrasion

of the coloured scales; hence it is very difficult to distinguish between them

in the field, especially between juveniles. In general, O. gracilipes has more

slender legs and is slightly smaller (body length, male: mean = 4.5 mm,

std.dev. = 0.3, n = 25; female: mean = 5.9 mm, std.dev. = 0.8, n = 25) than

either O. molarius (body length, male: mean = 6.5 mm, std.dev. = 0.9, n =

24; female: mean = 8.3 mm, std.dev. = 1.4, n = 31) or O. amoenus (body

length, male: mean = 6.4 mm, std.dev. = 1.2, n = 26; female: mean = 7.8 mm,

std.dev. = 1.3, n = 26) but this difference is often difficult to observe. The

most useful distinguishing characteristic of both adults and juveniles of the

three species is probably the markings on the femur of legs I and II (Figs 21-

23). O. gracilipes has a distinct, thin, dark longitudinal stripe on prolateral

femur 1 (like the seam on old-fashioned silk stockings: Figs 7, 8 and 23). O.

molarius has more variable leg markings; in some there are no markings on

femur 1, while in others it has one or two broad and diffuse longitudinal

stripes (Figs 1, 2, 3, 4 and 21). O. amoenus is generally a much darker spider

than the other two; although it has no longitudinal stripe on its femur 1 it may

have dark patches and it has transverse bands on the patella and tibia (Figs 5,

6 and 22).

Distribution

All three lynx spider species were found throughout the cotton growing

season. In the 2006/07 survey, 766 spiders were collected (including 62

adults) and these were identified to species using the characters described

above. Of these, O. molarius was the most common, followed by O.

amoenus. During this drought year O. gracilipes was relatively rare (Fig. 24).

Discussion

We identified three species of lynx spiders in cotton in the Namoi Valley. All

three species are found in all cotton growing regions southwards from

southern Queensland (Grimshaw 1991, Vink and Sirvid 2000, Whitehouse

and Grimshaw, unpublished data). Oxyopes molarius and O. amoenus have

also been collected from Townsville and Emerald in northern and central

Queensland respectively (Whitehouse and Grimshaw, unpublished data) and

both occur as far north as Cape York Peninsula and in South Australia

(Grimshaw 1991). O. amoenus has been found also in the Northern Territory

(Grimshaw 1991). O. gracilipes has a more southerly distribution, with

specimens found in SE Queensland and coastal regions of NSW, Victoria,

South Australia and SW Western Australia (Grimshaw 1991). Thus, although

this study focuses on the Namoi Valley, the species described are relevant to

most cotton growing regions in Australia.

Although spiders are one of the largest and most common invertebrate groups

in cotton, they are often ignored as agents in pest management. This is largely

because most work to date has focused on insects rather than spiders.

104 Australian Entomologist, 2007, 34 (4)

Consequently, the insect fauna in crops is well documented and much is

known about insect responses to crop conditions. This information has

generated models on insect economic thresholds and the effect of beneficial

insects on these thresholds (e.g. ‘the predator to pest ratio’: Mensah 2002,

Deutscher et al. 2005). Because spiders are quite distinct from insects,

models extrapolated from insect work may be not suitable for spiders. In

addition, spider species in cotton have not been well documented, making it

difficult for crop scouts to identify spiders beyond family level. Improving

spider identification will enable information specific to key spider species to

be incorporated into the management of pest species.

120

Number of individuals

Key

j O. amoenus

O. molarius

O. gracilipes

O @ mMJuveniles

N Adults

O 8

Sample date

Fig. 24. Number of lynx spiders caught in beatsheets in cotton during the 2006/07

season. In this drought year, O. molarius was the most common, followed by O.

amoenus; O. gracilipes was rare.

As mirids and other secondary pests become more of a problem in cotton, it

is important that key predators are accurately identified. Particular species of

lynx spiders are known to attack mirids. For example, the striped lynx spider,

Oxyopes salticus Hentz, is responsible for 15-18% of the daily mortality of

the cotton fleahopper Pseudatomoscelis seriatus (Reuter) (Hemiptera:

Miridae) in Texan cotton fields (Nyffeler et al. 1992) and 31% of all striped

lynx spiders in cotton had consumed immature fleahoppers (Breene ef al.

Australian Entomologist, 2007, 34 (4) 105

1989). In Australia, Bishop and Blood (1981) identified O. gracilipes (as

Oxyopes mundulus L. Koch) as an important predator of Helicoverpa spp

larvae in Australian cotton crops. In our own work we found that female O.

molarius were particularly good at attacking adult mirids, while O. gracilipes

was not as effective (Whitehouse and Barnes, unpublished data). To make

use of this information and develop it further, it is important to be able to

easily distinguish between these two very similar species.

Acknowledgements

We thank Judy Nobilo (CSIRO Entomology) for the illustrations and

photographs and for maintaining the spiders in the laboratory, and Rob Raven

(Queensland Museum) for comments on the manuscript and access to

collections. This research was funded by the Cotton Research and

Development Corporation (project number: 3.2.19 AC), and the Cotton

Catchment Communities Cooperative Research Centre (project number:

1.01.01).

References

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and pests in cotton and their importance to cotton production in southeastern Queensland.

General and Applied Entomology 13: 98-104.

BREENE, R.G., STERLING, W.L. and DEAN, D.A. 1989. Predators of the cotton fleahopper on

cotton. Southwestern Entomologist 14: 159-167.

DEUTSCHER, S., WILSON, L.J. and MENSAH, R.K. 2005. Integrated pest management

guidelines for cotton production systems in Australia, Australian Cotton Cooperative Research

Centre, Narrabri; 78 pp.

FITT, G.P. 2004. Implementation and impact of transgenic Bt cottons in Australia. Pp 371-381,

in: Cotton production for the new millennium. Proceedings of the third World cotton research

conference, 9-13 March 2003, Cape Town, South Africa. Agricultural Research Council -

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FITT, G.P.and WILSON, L.J. 2000. Genetic engineering in IPM: Bt cotton. Pp 108-125, in:

Kennedy, G.G. and Sutton, T.B. (eds), Emerging technologies for integrated pest management.

American Phytopathological Society Press, St. Paul; 526 pp.

GRIMSHAW, J.F. 1989. The genus Hamataliwa Keyserling (Araneae: Oxyopidae) in Australia

with description of two new species. Journal of the Australian Entomological Society 28: 181-

186.

GRIMSHAW, J.F. 1991. Revision of the Oxyopidae (Araneae: Arachnida) of Australia.

Unpublished M.Sc thesis. University of Queensland, St Lucia; 184 pp.

HAGERTY, A.M., KILPATRICK, A.L., TURNIPSEED, S.G., SULLIVAN, M.J. and

BRIDGES, W.C. 2005. Predaceous arthropods and lepidopteran pests on conventional, Bollgard,

and Bollgard II cotton under untreated and disrupted conditions. Environmental Entomology 34:

105-114.

MANSFIELD, S., DILLON, M.L. and WHITEHOUSE, M.E.A. 2006. Are arthropod

communities in cotton really disrupted? An assessment of insecticide regimes and evaluation of

the beneficial disruption index. Agriculture, Ecosystems and Environment 113: 326-335.

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MENSAH, R. 2002. Development of an integrated pest management programme for cotton. Part

I: Establishing and utilizing natural enemies. International Journal of Pest Management 48: 87-

94.

NYFFELER, M., STERLING, W.L. and DEAN, D.A. 1992. Impact of the striped lynx spider

(Araneae: Oxyopidae) and other natural enemies on the cotton fleahopper (Hemiptera: Miridae)

in Texas cotton. Environmental Entomology 21: 1178-1188.

ROEWER, C.F. 1954. Katalog der Araneae von 1758 bis 1940. Paul Budy, Bremen; 1751 pp.

TOWNSEND, V.R., FELGENHAUER, B.E. and GRIMSHAW, J.F. 2001. Comparative

morphology of the Australian lynx spiders of the genus Oxyopes (Araneae: Oxyopidae).

Australian Journal of Zoology 49: 561-576.

VINK, C.J. and SIRVID, P.J. 1998. The Oxyopidae (lynx spiders) of New Zealand. New

Zealand Entomologist 21: 1-9.

VINK, C.J. and SIRVID, P.J. 2000. New synonymy between Oxyopes gracilipes (White) and

Oxyopes mundulus L. Koch (Oxyopidae: Araneae). Memoirs of the Queensland Museum 45:

637-640.

WHITEHOUSE, M.E.A., HARDWICK, S., SCHOLZ, B.C.G., ANNELLS, A., WARD, A.,

GRUNDY, P. and HARDEN, S. In press. Evidence of a latitudinal gradient in spider diversity in

Australian cotton. Austral Ecology.

WHITEHOUSE, M.E.A., WILSON, L.J. and FITT, G.P. 2005. A comparison of arthropod

communities in transgenic Bt and conventional cotton in Australia. Environmental Entomology

34: 1224-1241.

YOUNG, O.P. and LOCKLEY, T.C. 1986. Predation of striped lynx spider, Oxyopes salticus

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Australian Entomologist, 2007, 34 (4): 107-114 107

A REVIEW OF CALLISTOMYIA BEZZI AND RELATED GENERA

(DIPTERA: TEPHRITIDAE: TRYPETINAE)

D.L. HANCOCK

PO Box 2464, Cairns, Qld 4870

Abstract

The new tribe Callistomyiini is proposed for the Indo-Australian genera Callistomyia Bezzi and

Alincocallistomyia Hardy, plus the African genus Sosiopsila Bezzi. This tribe might also include

the Neotropical genera Alujamyia Norrbom, Molynocoelia Giglio-Tos and Pseudophorellia

Lima. The African Sosiopsila trisetosa Bezzi, stat. rev., is removed from synonymy with S.

metadacus (Speiser). The identity of the Indian Dacus klugii Wiedemann is discussed, with the

species removed from Callistomyia and placed in the new combination Euphranta klugii

(Wiedemann), close to E. apicalis Hendel. A key to the three genera and eight species known

from the Old World is included.

Introduction

The Indo-Australian genera Callistomyia Bezzi and Alincocallistomyia Hardy

have had a chequered taxonomic history. Currently included in the otherwise

Neotropical tribe Hexachaetini (e.g. Korneyev 1999, Hancock and Drew

2003, Agarwal and Sueyoshi 2005), they were previously placed in the tribes

Acanthonevrini (Hardy 1986, 1988) or Trypetini (Hancock and Drew 1994,

Permkam and Hancock 1995, Wang 1998). However, a recent study by

Norrbom (2006) suggests that they represent a distinct clade closely related to

the Neotropical genera Alujamyia Norrbom, Molynocoelia Giglio-Tos and

Pseudophorellia Lima, with Hexachaeta Loew forming a separate clade with

Anastrepha Schiner and Toxotrypana Gerstaecker (the latter three genera all

placed in tribe Toxotrypanini). Norrbom (2006) suggested that they might

also be related to tribe Adramini; however, all lack long, fine hairs on the

anatergite, a key defining character for that tribe and (as indicated by

Norrbom 2006) a sister-group relationship with the Toxotrypanini seems

more certain.

Tribal placement of the African genus Sosiopsila Bezzi has been equally

uncertain. Hancock (1986) transferred it from the Adramini to tribe

Phytalmiini but the yellow dorsal stripe on the anepisternum and the lack of

preapical setae on the aculeus suggest that this was incorrect. It was returned

tentatively to [or near] the Adramini by Korneyev (1999) and Hancock

(2003). However, it also lacks long, fine hairs on the anatergite and its

inclusion within the Adramini remained doubtful. The shape and structure of

the spermathecae, the male fifth sternite and the aedeagus closely resemble

those of Callistomyia (see Munro 1984, Hardy 1973, 1974, Permkam and

Hancock 1995), the unusually shaped, apically trilobed aculeus is similar to

that seen in Pseudophorellia (see Hancock 1986, Norrbom 2006), while the

shape of the epandrium and proctiger resemble that of Molynocoelia (see

Munro 1984, Norrbom 2006). Sosiopsila shares with Alincocallistomyia and

Callistomyia the pubescent arista, ocellar setae vestigial or absent, vein R4+5

108 Australian Entomologist, 2007, 34 (4)

extensively setose and an apically pointed aculeus without preapical setae.

Sosiopsila has only one pair of scutellar setae (2-3 pairs in the other genera)

but this appears to be the medial pair, with the basal and apical pairs lost.

Sosiopsila is here tentatively included with the latter two genera within the

new tribe Callistomyiini, which might also include the Molynocoelia group of

genera (see Norrbom 2006).

Subfamily TRYPETINAE

Tribe CALLISTOMYIINI nov.

Type genus Callistomyia Bezzi.

This tribe is proposed to accommodate a group of apparently related genera

that lack defining characters of other tribes currently included in the

subfamily Trypetinae (see Korneyev 1999, Norrbom 2006). Two Indo-

Australian genera (Alincocallistomyia Hardy, Callistomyia Bezzi) and one

African genus (Sosiopsila Bezzi) are included. Three Neotropical genera

(Alujamyia Norrbom, Molynocoelia Giglio-Tos and Pseudophorellia Lima)

might also belong here.

Diagnosis. Head generally with 1-2 pairs of orbital setae and 2-3 pairs of

frontal setae; ocellars weak or absent; all setae dark and acuminate; arista

usually pubescent; occiput swollen ventrally. Scutum generally fulvous to

reddish, with or without dark spots or vittae; anepisternum usually with a

yellow dorsal band from postpronotal lobe to wing base; anatergite without

long, fine hairs; postpronotal, presutural, dorsocentral and prescutellar

acrostichal setae present or absent; dorsocentrals, when present, posterior in

position, close to line of postalars; intrapostalars lacking; 1-3 pairs of

scutellar setae. Legs often with a second, smaller spine at apex of mid tibia

and with or without two rows of ventral spinules on mid and hind femora.

Wing usually banded but pattern sometimes modified or reduced; veins R;

and R4+s extensively setose; no distinct costal seta at base of pterostigma;

pterostigma usually narrow and apically acute; vein M not distinctly curved

upwards at apex; cell bcu weakly or strongly acuminate but apical extension

not basally constricted. Abdominal tergites not fused, often with dark spots or

bands but without shiny black bullae; aculeus apically acute, sometimes

trilobed with the preapical lobes also acute, and without preapical setae;

eversible membrane with ventral spicules much more extensive than dorsal

spicules (Norrbom 2006: unchecked for Sosiopsila); 2-3 spermathecae,

usually round or mushroom-shaped.

The identity of ‘Callistomyia’ klugii

Dacus klugii Wiedemann was described from ‘India orient[alis]’ by

Wiedemann (1824) [‘orientalis’ signifying India proper rather than the West

Indies]. Bezzi (1913) tentatively suggested it might belong in Callistomyia

and that was followed, without further comment, by Hardy (1951) and all

subsequent authors. No additional specimens have been referred to it since its

Australian Entomologist, 2007, 34 (4) 109

original description and neither Senior-White (1924) nor Kapoor (1970)

mentioned it. However, the name has been recognised as valid in several

recent catalogues (Hardy 1977, Kapoor 1993, Norrbom et al. 1999, Agarwal

and Sueyoshi 2005) and in at least three systematic keys (Hardy 1951, 1974,

Kapoor 1993).

The type (in the Zoological Museum, University of Copenhagen) is possibly

from the Calcutta district of West Bengal, where Dagobert Daldorf (the likely

collector) was based between 1798 and his death in 1802 (Courtice 2006).

Unfortunately, I have not been able to obtain any additional information on

the type but Wiedemann’s (1830) description suggests it belongs in genus

Euphranta Loew. Note that the basal, transverse wing band enclosing

crossvein BM-Cu is [dark] brown, not yellowish as in Callistomyia, and that

the hyaline distal areas are ‘whitish’. It appears closest to, and is possibly

synonymous with, E. apicalis Hendel, seemingly differing only in the

apparent absence of dark brown vittae on the scutum and a smaller hyaline

apical spot on the wing (confined to cell r4,5).

A relationship with Euphranta apicalis, a species widespread in southeast

Asia but not yet recorded from India, is suggested by size and overall body

colour, apparent similarities in wing pattern and the presence of a pair of

blackish facial spots (Wiedemann 1830, Hendel 1915). However, the nearest

recorded locality for E. apicalis is the Moulmein district of southern Burma

(Hering 1938, Wang 1998) and, in the absence of additional Indian material

and without further information on the type, formal synonymy would be

premature. Therefore, Euphranta klugii (Wiedemann, 1824), comb. n. and

Euphranta apicalis Hendel, 1915 are regarded here as separate species placed

in the apicalis group of Hancock and Drew (2004).

It should be noted that, prior to Daldorf’s return to India in 1798, he also

collected in Sumatra (Courtice 2006), where E. apicalis is known to occur

(Hancock and Drew 2004); however, given the original type locality of ‘India

[of the East]? (Wiedemann 1824), the Calcutta district is a more likely

provenance. Wiedemann (1830) subsequently misstated the type locality as

‘Ostindien’, leading some authors (e.g. Hardy 1951) to incorrectly interpret

the type locality as ‘East Indies’ [Indonesia].

Key to Old World genera and species of tribe Callistomyiini

I One pair of scutellar setae; postpronotal and dorsocentral setae absent;

mid and hind femora without rows of black ventral spinules; wing without

a transverse band across R-M crossvein; cell bcu weakly acute apically

(Africa JE SE EN Sosiopsila Bezzi ... 2

— Two or three pairs of scutellar setae; postpronotal and dorsocentral setae

present; mid and hind femora with rows of black ventral spinules; wing

with a transverse band across R-M crossvein; cell bcu strongly acute

aplcallyg(IndO-AUStralid) POE er str. 4

Australian Entomologist, 2007, 34 (4)

Postnotum fulvous, at most with a slight medial darkening; apical wing

spot extending narrowly (for about 1/3 length) below vein R4+5 (South

ATTICA) Pts kr ota, AR) S. rotunda Munro

Postnotum with a distinct medial black or brown patch; apical wing spot

extending broadly (for 1/2-2/3 length) below vein R45 ...........0e0eeeee 3

Costal band in cell r; narrow but distinct, wider than costal vein (Nigeria

to Ethiopia and ?Kenya) ia a E A E S. metadacus (Speiser)

Costal band in cell r, absent or vestigial, no wider than costal vein

(Mala Wiito South Africa) PE S. trisetosa Bezzi

Three pairs of scutellar setae; prescutellar setae present; no distinct

propleural seta (Borneo) .................. Alincocallistomyia imitator Hardy

Two pairs of scutellar setae; prescutellar setae absent; a distinct

propleuralisetalpresen AP Callistomyia Bezzi ... 5

Wing with large apical spot broadly connected in posterior half of cell dm

with transverse band across R-M crossvein; abdomen without black

transverse bands on terga II-V [present or absent on tergite II] (northern

Australia, southern Papua New Guinea) ..............0.000 C. horni Hendel

Wing with large apical spot broadly isolated or at most narrowly

connected along vein Cu; with transverse band across R-M crossvein;

abdomen usually with black transverse bands on terga H-V ............... 6

Wing with large apical spot narrowly connected with transverse band

along vein Cu, (Philippines) a a C. icarus (Osten Sacken)

Wing with large apical spot broadly separated from transverse band ..... 7

Wing with large apical spot diffusely connected with costal band along

apical margin of cell r}, (India and China to western Indonesia)

RS ORARAA AIE PA E OAAR avenue C. pavonina Bezzi

Wing with large apical spot not connected with costal band along apical

margin of cell r}, (eastern Indonesia, northern Papua New Guinea)

A AARI L TA N E nis a a ot bebe C. flavilabris Hering

Systematics

Genus ALINCOCALLISTOMYIA Hardy

Alincocallistomyia Hardy, 1986: 28. Type species A. imitator Hardy.

One species, from the island of Borneo. Larval hosts unknown.

Alincocallistomyia imitator Hardy

Alincocallistomyia imitator Hardy, 1986: 29. (near Tawau, Sabah, Malaysia).

Distribution. Known only from Sabah, east Malaysia.

Australian Entomologist, 2007, 34 (4) 111

Genus CALLISTOMYIA Bezzi

Callistomyia Bezzi, 1913: 124. Type species C. pavonina Bezzi.

Four allopatric Indo-Australian species. Larval hosts Rutaceae (subfamily

Aurantiodeae). For a habitus illustration of the type species, see Hancock and

Drew (1994). The Australian C. horni has a distinctive wing pattern but the

other three species (C. flavilabris, C. icarus and C. pavonina) are only

weakly separable.

Callistomyia flavilabris Hering

Callistomyia flavilabris Hering, 1953: 513. (Misool, Indonesia).

Distribution. Eastern Indonesia (Maluku Province: Misool) and northern

Papua New Guinea (Madang Province).

Host plant. Berries of Wenzelia dolichophylla (Rutaceae) (Hancock and

Drew 2003).

Callistomyia horni Hendel

Callistomyia horni Hendel, 1928: 361. (Palmerston [Darwin], Northern Territory).

Distribution. Southern Papua New Guinea (Central Province) and Australia

(northern areas of Western Australia, Northern Territory and Queensland).

Host plants. Berries of Clausena, Glycosmis and Micromelum species

(Rutaceae) (Permkam and Hancock 1995).

Callistomyia icarus (Osten Sacken)

Dacus icarus Osten Sacken, 1882: 224. (Philippines).

Callistomyia icarus: Hardy, 1974: 160. (Luzon).

Distribution. Philippines (Luzon).

Host plants. None recorded.

Callistomyia pavonina Bezzi

Callistomyia pavonina Bezzi, 1913: 125. (NE India).

Callistomyia flavilabris: Hardy, 1973: 177-178. (Malaysia). Misidentification.

In specimens previously referred to C. pavonina, the dark facial spot is

variable in intensity and is often absent. Malaysian specimens of ‘C.

flavilabris” recorded by Hardy (1973) were regarded as examples of C.

pavonina with the facial spot absent by Hancock and Drew (1994).

Distribution. India, Sri Lanka, China, Taiwan, Thailand, Laos, Vietnam, west

Malaysia and western Indonesia (Sumatra, Java).

Host plants. Berries of Clausena and Glycosmis species (Rutaceae) (Hancock

and Drew 1994).

112 Australian Entomologist, 2007, 34 (4)

Genus SOSIOPSILA Bezzi

Sosiopsila Bezzi, 1920: 214. Type species S. trisetosa Bezzi.

Three poorly differentiated (and apparently allopatric) African species that

are probably little more than subspecies. Larval hosts unknown. Munro

(1984) suggested that larvae were probably stem borers, but that is unlikely to

be the case. For a habitus illustration of the type species, see Hancock (1986,

as metadacus).

Sosiopsila metadacus (Speiser)

Polystodes metadacus Speiser, 1915: 99. (Zela, Mandarra Mts, Cameroon).

Sosiopsila metadacus: Hancock, 1986: 304. (Cameroon).

Distribution. Known from northern Nigeria, Cameroon and Ethiopia

(material in The Natural History Museum, London) and possibly western

Kenya (Copeland et al. 2005, as Sosiopsila sp. cf. metadacus).

Sosiopsila rotunda Munro

Sosiopsila rotunda Munro, 1933: 26. (Rosslyn, South Africa).

Distribution. Known from northern and eastern South Africa (North-West

Province: Rosslyn and Rustenburg; KwaZulu-Natal: Durban).

Sosiopsila trisetosa Bezzi; stat. rev.

Sosiopsila trisetosa Bezzi, 1920: 215. (East of Mt Mlanje, Mozambique).

Sosiopsila metadacus: Hancock, 1986: 303-304. (Zimbabwe). Misidentification.

This species was placed as a synonym of S. metadacus by Hancock (1986).

However, examination of further material (in The Natural History Museum,

London) suggests that the variation observed (particularly the presence or

absence of a distinct costal band) is at least partly geographical.

Distribution. Known from southern Malawi, Mozambique, Zimbabwe and

northeastern South Africa (Mpumalanga Province: Nelspruit).

Acknowledgement

I thank Nigel Wyatt (The Natural History Museum, London) for access to

material in his care.

References

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Tephritidae). Oriental Insects 39: 371-433.

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Memoirs of the Indian Museum 3: 53-175, pls 8-10.

BEZZI, M. 1920. Notes on the Ethiopian fruit flies of the family Trypaneidae, other than Dacus

— II. Bulletin of Entomological Research 10: 211-272.

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COPELAND, R.S., OKEDA, W., FREIDBERG, A., MERZ, B., WHITE, I.M., DE MEYER, M.

and LUKE, Q. 2005. Fruit flies (Diptera, Tephritidae) of Kakamega Forest, Kenya. Journal of

East African Natural History 94(2): 247-278. i

COURTICE, A.C. 2006. Of peaches and maggots: the story of Queensland fruit fly. Hillside

Books, Marengo, NSW; 336 pp.

HANCOCK, D.L. 1986. Classification of the Trypetinae (Diptera: Tephritidae), with a

discussion of the Afrotropical fauna. Journal of the Entomological Society of Southern Africa

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HANCOCK, D.L. 2003. An annotated checklist of the Tephritidae of Zimbabwe (Diptera:

Schizophora), with a key to subgenera of Dacus Fabricius. Cimbebasia 19: 111-148.

HANCOCK, D.L. and DREW, R.A.I. 1994. New species and records of Asian Trypetinae

(Diptera: Tephritidae). Raffles Bulletin of Zoology 42(3): 555-591.

HANCOCK, D.L. and DREW, R.A.I. 2003. New species and records of Trypetinae (Diptera:

Tephritidae) from Australia and the South Pacific. Australian Entomologist 30(3): 93-106.

HANCOCK, D.L. and DREW, R.A.I. 2004. Notes on the genus Euphranta Loew (Diptera:

Tephritidae), with description of four new species. Australian Entomologist 31(4): 151-168.

HARDY, D.E. 1951. The Krauss collection of Australian fruit flies. Pacific Science 5(2): 115-

189.

HARDY, D.E. 1973. The fruit flies (Tephritidae-Diptera) of Thailand and bordering countries.

Pacific Insects Monograph 31: 1-353.

HARDY, D.E. 1974. The fruit flies of the Philippines (Diptera: Tephritidae). Pacific Insects

Monograph 32: 1-266.

HARDY, D.E. 1977. Family Tephritidae (Trypetidae, Trupaneidae). Pp 44-134, in: Delfinado,

M.D. and Hardy, D.E. (eds), A catalogue of Diptera of the Oriental Region. Volume III,

Suborder Cyclorrhapha (excluding Division Aschiza). University of Hawaii Press, Honolulu; x +

854 pp.

HARDY, D.E. 1986. Fruit flies of the subtribe Acanthonevrina of Indonesia, New Guinea, and

the Bismarck and Solomon Islands (Diptera: Tephritidae: Trypetinae: Acanthonevrini). Pacific

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HARDY, D.E. 1988. Fruit flies of the subtribe Gastrozonina of Indonesia, New Guinea, and the

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114 Australian Entomologist, 2007, 34 (4)

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73-113, in: Aluja, M. and Norrbom, A.L. (eds), Fruit flies (Tephritidae): phylogeny and

evolution of behavior. CRC Press, Boca Raton; xvi + 944 pp.

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No. 61: ix + 313 pp.

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A. (ed.), Biotaxonomy of Tephritoidea. Israel Journal of Entomology 35-36: xii + 599 pp.

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1999. Systematic database of names. Pp 65-251, in: Thompson, F.C. (ed.), Fruit fly expert

identification system and systematic information database. Myia 9. Backhuys Publishers, Leiden;

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Semper [concl.]. Berliner Entomologische Zeitschrift (Berlin) 26: 187-252.

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Government of India, Calcutta; iii + 33 pp.

SPEISER, P.G.E. 1915. Beitrage zur Dipterenfauna von Kamerun. III. Deutsche Entomologische

Zeitschrift (Berlin) 1915: 91-106.

WANG, X.-J. 1998. The fruit flies (Diptera: Tephritidae) of the East Asia Region. Acta

Zootaxonomixa Sinica 21(Supplement 1996): i-viii, 1-419.

WIEDEMANN, C.R.W. 1824. Munus rectoris in Academia Christiana Albertina aditurus

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illustrat. Kiliae [Kiel]; 60 pp.

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Australian Entomologist, 2007, 34 (4): 115-118 115

A NEW SPECIES OF THE SUBGENUS POLYRHACHIS

(CYRTOMYRMA) FOREL (HYMENOPTERA: FORMICIDAE:

FORMICINAE) FROM BORNEO

RUDOLF J. KOHOUT

Biodiversity Program, Queensland Museum, PO Box 3300, South Brisbane, Qld 4101

(Email: rudolfkohout@qm.qld.gov.au)

Abstract

Polyrhachis acuminata, a new species of the subgenus Cyrtomyrma Forel, is described and

illustrated from Sabah, Malaysia.

Introduction

During a recent, one day visit to Poring Hot Springs in Kinabalu Park, Sabah,

Malaysia, I collected a few Polyrhachis Fr. Smith specimens, including two

that I regarded at the time to belong to a species recently described as

Polyrhachis (Cyrtomyrma) danum Kohout (Kohout 2006). These specimens

featured several characteristics of that species, including distinctly reddish-

brown appendages and a similar mesosomal outline. However, following my

return to Brisbane, closer examination of the specimens revealed them to be

an undescribed species. This discovery was made too late for the species to

be included in my recent revision of the Bornean fauna of subgenus

Cyrtomyrma Forel (Kohout 2006) and consequently it is described below.

Abbreviations of institutions (with names of curators) are: ITBC = Institute

for Tropical Biology and Conservation, Universiti Malaysia Sabah, Kota

Kinabalu, Sabah, East Malaysia (Dr Maryati Mohamed); QMBA =

Queensland Museum, Brisbane, Australia (Dr Chris J. Burwell).

Methods

Photographs of the holotype were taken by Geoff Thompson (QMBA) with a

Leica DFC500 Camera and Leica MZ16A stereomicroscope, using Leica

Application Suite Software. The images were then processed using Auto-

Montage (Syncroscopy, Division of Synoptics Ltd, USA) and Adobe CS2

(Adobe Systems Inc, USA) software.

Standard measurements and indices are as follows: TL = Total length (the

necessarily composite measurement of the outstretched length of the entire

ant measured in profile); HL = Head length (the maximum measurable length

of the head in perfect full face view, measured from the anterior-most point

of the clypeal border or teeth, to the posterior-most point of the occipital

margin); HW = Head width (width of the head in perfect full face view,

measured immediately in front of the eyes); CI = Cephalic index (HW x

100/HL); SL = Scape length (length of the antennal scape, excluding the

condyle); SI = Scape index (SL x 100/HW); PW = Pronotal width (greatest

width of the pronotal dorsum); MTL = Metathoracic tibial length (maximum

measurable length of the tibia of the hind leg). All measurements were taken

116 Australian Entomologist, 2007, 34 (4)

using a Zeiss SR stereomicroscope with an eyepiece graticule calibrated

against a stage micrometer and are expressed in millimetres (mm).

Polyrhachis acuminata sp. n.

(Figs 1-2)

Types. Holotype worker, EAST MALAYSIA (SABAH): Kinabalu Park, Poring Hot

Springs, 06°02°N, 116°43’E, 27.vi.2006, R.J. & E. Kohout acc. 06.5. Paratype: |

worker, same data as holotype. Holotype in ITBC; paratype in QMBA.

Description. Worker. Dimensions (holotype cited first): TL c. 6.55, 6.75; HL

1.59, 1.62; HW 1.50, 1.53; CI 94, 94; SL 1.96, 2.03; SI 131, 133; PW 1.18,

1.22; MTL 2.34, 2.37 (2 measured). Mandibles with 5 teeth, apical tooth

longest, other teeth subequal in length. Anterior clypeal margin obtusely

truncate with shallow median notch. Clypeus in profile convex with shallow

depression behind anterior margin and moderately impressed basal margin.

Frontal triangle indistinct. Frontal carinae sinuate with weakly raised

margins; central area shallowly concave with shallowly impressed frontal

furrow. Sides of head in front of eyes converging towards mandibular bases

in weakly convex line; behind eyes sides rounding into convex occipital

margin. Eyes moderately convex, in full face view only marginally breaking

lateral cephalic outline. Ocelli lacking. Pronotum in dorsal view with humeri

widely rounded, with greatest pronotal width just before mid-length of

segment. Mesosoma in profile moderately convex, with promesonotal suture

distinctly impressed; metanotal groove lacking dorsally, weakly indicated

laterally. Propodeal dorsum with indication of rudimentary propodeal spines;

rounding into relatively short, oblique declivity. Petiole with dorsal spines

reduced to short, blunt, wide-based teeth; lateral spines up to four times as

long as their basal width, slender and acute. Subpetiolar process relatively

long, angular anteriorly, narrowly rounded posteriorly. Anterior face of first

gastral segment in lateral view lower than full height of petiole, very weakly

concave at base, narrowly rounding onto dorsum of segment.

Mandibles finely longitudinally striate with numerous piliferous pits;

sculpture becoming rather smooth and polished towards bases. Head,

mesosoma, petiole and gaster very finely shagreened, rather polished with

numerous shallow punctures; sculpture becoming somewhat finely wrinkled

on meso- and metapleurae. Petiole very finely transversely wrinkled, with

sculpture more intensely reticulate-rugose at base.

Mandibles with numerous semierect hairs at masticatory borders. Anterior

clypeal margin with several moderately long, anteriorly directed setae and a

few short setae lining margin laterally. Two pairs of erect hairs arising near

anterior margin and one pair along frontal carinae. Rather long, erect hairs on

anterior face of fore coxae; distinctly shorter hairs on posterior face and on

ventral surfaces of trochanters. Gaster with medium length, erect hairs lining

posterior margins of apical segments, hairs on gastral venter more abundant.

Australian Entomologist, 2007, 34 (4) 117

Black, with mandibular masticatory borders, condylae and extreme tips of

apical funicular segments reddish-brown. Legs, including trochanters,

distinctly red or reddish-brown with proximal ends of tibiae, coxae and tarsi

virtually black.

Sexuals and immature stages unknown.

Etymology. From the Latin acuminatus, meaning pointed, in reference to the

long, sharply pointed lateral petiolar spines.

Figs 1-2. Polyrhachis acuminata sp. n., holotype worker: (1) dorsal view; (2) lateral

view.

Remarks. Polyrhachis acuminata is very similar to P. danum and P. lepida

Kohout, both also known from Sabah. All three species are distinctly

bicoloured with the head, mesosoma, petiole and gaster black and most of the

legs bright red or reddish-brown. They have a rather similar lateral

mesosomal outline, except that the propodeal declivity is virtually vertical in

P. danum and P. lepida, while it is oblique in P. acuminata. Polyrhachis

acuminata is also distinguished by its rudimentary propodeal spines that are

completely absent in P. danum or indicated only as barely visible tubercles in

some P. lepida specimens. The mesosomal sculpturation is uniformly finely

shagreened in P. danum, while the sides of the mesosoma are wrinkled in P.

acuminata and distinctly reticulate-rugose in P. lepida. However, the main

character that distinguishes the three species is the configuration of petiolar

spines. In P. danum, all the petiolar spines are reduced to minute denticles,

118 Australian Entomologist, 2007, 34 (4)

while in P. acuminata and P. lepida the dorsal pair are wide-based and tooth-

like and the lateral spines are long and slender. In P. acuminata the lateral

spines are up to four times as long as their basal widths, while they are only

twice as long or shorter in P. lepida. The species also differ in their relative

sizes, with P. lepida the smallest and P. danum the largest (HL 1.40-1.50 in

P. lepida, 1.59-1.62 in P. acuminata and 1.65-1.87 in P. danum).

Key to Bornean P. (Cyrtomyrma) species

Polyrhachis acuminata is the latest addition to the list of Bornean

Cyrtomyrma species and can be identified using the following modification

to the key to Bornean species in Kohout (2006). Figure numbers refer to

illustrations in the original article. Polyrhachis rastellata (Latreille),

erroneously recorded from Borneo in the past (see Kohout 2006 for details),

is included in the key for completeness.

6 Larger species (HL >1.65); petiole with sides more-or-less parallel,

spines reduced to minute denticles (Fig. 6F) ..............+ danum Kohout

— Smaller species (HL <1.65); petiole with sides diverging, spines acute

(HIGH BAD) PE SEE ects. EE eis teeters LSE 7

7 Pronotal shoulders broadly rounded; lateral petiolar spines up to four

times as long as their basal width .....................seee ee acuminata sp. n.

— Pronotal shoulders narrowly rounded or subangular (Fig. 7A); lateral

petiolar spines at most twice as long as their basal width ................... 8

8 Propodeal declivity very steep, virtually vertical (Fig. 7A); lateral

petiolar spines distinctly longer than dorsal pair (Fig. 7B); legs dark

Teddish=brownp Fer eet tee aae tame nes Pentre? Ort lepida Kohout

— Propodeal declivity oblique (Fig. 7C); petiolar spines subequal or lateral

pair shorter than dorsal pair (Fig.7D); legs mostly orange or light

reddish-brown ............ecssessssseseseseseeseseseseeseteeee es Pastellata (Latreille)

Acknowledgements

I am very grateful to Prof. Datin Dr Maryati Mohamed (Director of the

Institute for Tropical Biology and Conservation, Universiti Malaysia Sabah)

for continued logistic and financial support of my work on the taxonomy of

Bornean Polyrhachis. My gratitude goes also to Geoff Thompson

(Queensland Museum) for producing the digital images and Dr Chris Burwell

(Queensland Museum) for reading and commenting on a draft of the

manuscript.

Reference

KOHOUT, R.J. 2006. Review of Polyrhachis (Cyrtomyrma) Forel (Hymenoptera: Formicidae:

Formicinae) of Australia, Borneo, New Guinea and the Solomon Islands with descriptions of

new species. Memoirs of the Queensland Museum 52(1): 87-146.

Australian Entomologist, 2007, 34 (4): 119-120 119

THE IDENTITY OF TERELLIA IMMACULATA MACQUART

(DIPTERA: TEPHRITIDAE: TEPHRITINAE)

D.L. HANCOCK

PO Box 2464, Cairns, Old 4870

Abstract

Terellia immaculata Macquart, 1855 is placed as a new synonym of the Palaearctic Terellia

longicauda (Meigen, 1838). Its stated type locality of Marquesas Islands, French Polynesia, is

regarded as an error, possibly for Marquise in NW France.

Introduction

The fruit fly species Terellia immaculata Macquart has remained

unrecognised since its original description (Macquart 1855). The type female

is in the Oxford University Museum of Natural History (OUMNH) and is

presumed to be of French Polynesian origin. Bezzi (1913) listed it from the

Marquise Islands, while Hardy and Foote (1989) and Norrbom er al. (1999)

both regarded it as an unplaced species of Tephritidae from French Polynesia

(Marquesas Islands). However, the tephritine tribe Terelliini, to which

Terellia Robineau-Desvoidy belongs, has not otherwise been reported from

either the Australasian or Oceanian Regions.

Through the kindness of Adrian Pont and James Hogan (OUMNH), I have

been able to examine photographs of the type female. Apart from a missing

head, the type is in reasonably good condition and diagnostic characters are

clearly visible. It is undoubtedly the same taxon as Terellia longicauda

(Meigen), a widespread Palaearctic species.

Terellia longicauda (Meigen)

Trypeta longicauda Meigen, 1838: 356. (Bavaria, Germany).

Trypeta acuticornis Loew, 1846: 520. (? Wurttemburg, Germany).

Terellia immaculata Macquart, 1855: 145. (Iles Marquises [? Marquesas Is, French

Polynesia] — error?). Syn. n.

Terellia (Terellia) longicauda (Meigen): Norrbom et al., 1999: 222.

Type data. The type of T. immaculata carries the following labels: (1) [in handwriting

of P.J.M. Macquart] — ‘Terellia R.D. / immaculata / 9, Macq. n. sp.’; (2) [in

handwriting of J.M.F. Bigot] — ‘Trypeta immaculata. 9. / Terellia. id. Macq. / Ins.

Marquis. Macq.’ [‘rypeta’ subsequently inserted after ‘T’ by J.E. Collin];

(3) [handwritten & printed] — ‘T. immaculata / EX COLL. BIGOT’, (4) [circular

printed label with red border] — ‘Holo-/ type’.

Comments. The characters of T. immaculata (particularly the scutal pattern,

yellow scutellum, hyaline wing with a pale yellow stigma, sectional lengths

of the medial vein, white-setose abdomen, long oviscape and aculeus shape)

are consistent with those of T. longicauda as discussed and illustrated by

White (1988) and Merz (1994). The stated type locality of ‘iles Marquises’

[Marquesas Is] is evidently erroneous and is possibly a misrepresentation of

Marquise, a town near Boulogne in northwestern France.

120 Australian Entomologist, 2007, 34 (4)

Host plant. Larvae of T. longicauda feed in the flower heads of the thistle

Cirsium eriophorum (L.) Scop. (Asteraceae: Cardueae) (White 1988, Merz

1994).

Distribution. Great Britain, central Europe and western Siberia to Spain, the

Balkans and Iran (Norrbom er al. 1999).

Discussion

With the removal of Terellia immaculata from the faunal list for the

Marquesas Islands, only two other species of Tephritidae remain, the

widespread Dioxyna sororcula (Wiedemann) and the endemic Trupanea

simplex Malloch (Hardy and Foote 1989). Both belong in tribe Tephritini in

the flower-infesting subfamily Tephritinae. No fruit-infesting species are

known from these remote Pacific islands (Purea et al. 1996).

Acknowledgements

I thank Adrian Pont and James Hogan (UOMNH) for photographs of the type

of Terellia immaculata and for help in interpreting the handwritten label data.

References

BEZZI, M. 1913. Indian trypaneids (fruit flies) in the collection of the Indian Museum, Calcutta.

Memoirs of the Indian Museum 3: 53-175, pis 8-10.

HARDY, D.E. and FOOTE, R.H. 1989. Family Tephritidae. Pp 502-531, in: Evenhuis, N.L.

(ed.), Catalog of the Diptera of the Australian and Oceanian Regions. Bishop Museum Special

Publication 86. Bishop Museum Press, Honolulu and E.J. Brill, Leiden; 1155 pp.

LOEW, H. 1846. Fragmente zur Kenntniss der europåischen Arten einiger Dipterengattungen.

Linnaea Entomologica, Berlin 1: 319-530, pl. III.

MACQUART, P.J.M. 1855. Diptéres exotiques nouveau ou peu connus. 5e supplément.

Mémoires de la Société des Sciences, de l’Agriculture et des Arts de Lille (1854) (2) 1: 25-156, 7

pls.

MEIGEN, J.W. 1838. Systematische Beschreibung der bekannten europdischen zweifliigeligen

Insekten. Siebenter Theil oder Supplementband. Schultz, Hamm; xii + 434 + [1] pp.

MERZ, B. 1994. Diptera Tephritidae. Insecta Helvetica Fauna 10. Schweitzerische

Entomologische Gesellschaft, Geneva; [vii] + 198 pp.

NORRBOM, A.L., CARROLL, L.E., THOMPSON, F.C., WHITE, I.M. and FREIDBERG, A.

1999. Systematic database of names. Pp 65-251, in: Thompson, F.C. (ed.), Fruit fly expert

identification system and systematic information database. Myia 9. Backhuys Publishers, Leiden;

ix + 524 pp.

PUREA, M., PUTOA, R. and MUNRO, E. 1996. Fauna of fruit flies in the Cook Islands and

French Polynesia. Pp 54-56, in: Allwood, A.J. and Drew, R.A.I. (eds), Management of fruit flies

in the Pacific. Aciar Proceedings No. 76. Australian Centre for International Agricultural

Research, Canberra; 267 pp.

WHITE, I.M. 1988. Tephritid flies (Diptera: Tephritidae). [Royal Entomological Society of

London] Handbooks for the identification of British insects. Vol. 10. Part 5a. British Museum

(Natural History), London; 134 pp.

Australian Entomologist, 2007, 34 (4): 121-122 121

NOTES ON THE DISTRIBUTION AND BIOLOGY OF TRAPEZITES

GENEVIEVEAE ATKINS, SIGNETA TYMBOPHORA (MEYRICK &

LOWER) AND HESPERILLA SARNIA ATKINS

(LEPIDOPTERA: HESPERIIDAE)

ANDREW ATKINS

PO Box 42, Eudlo, Old 4554

Abstract

Biological notes and distribution records are presented for three uncommon species of trapezitine

skippers: Trapezites genevieveae Atkins, Signeta tymbophora (Meyrick & Lower) and

Hesperilla sarnia Atkins.

Introduction

Between 2000 and 2007, a number of observations were made of three

endemic trapezitine skippers in SE Queensland. Although the larval food

plants of each species are local but fairly widespread within temperate and

subtropical biomes in eastern Australia, the skippers are uncommon to rare

with short flight seasons and are found only within limited areas. It appears

that further, individually specific requirements, possibly climatic, altitude and

microhabitat associations, are necessary to each species and thus are probable

subjects of conservation value that emphasise the importance of the particular

biomes that support them.

Trapezites genevieveae Atkins is restricted to dense old-growth montane

rainforest from the Barrington Ranges in New South Wales to the Blackall

Ranges in SE Queensland; Signeta tymbophora (Meyrick & Lower) is found

very locally in coastal rainforest from near Narooma in New South Wales to

upland temperate rainforest at Bunya Mts in SE Queensland; Hesperilla

sarnia Atkins occurs from SE Queensland to the Cairns area of northern

Queensland (Braby 2000). All three species have been found in mixed wet

sclerophyll/upland rainforest at Mapleton in the Blackall Ranges (personal

observations). Further searches for larvae and adults of these species beyond

the northern extensions of the Sunshine Coast (southern Queensland) and

also at Eungella Range (hinterland of Mackay, northern Queensland) so far

have been unsuccessful.

Observations

Trapezites genevieveae

Adults are rarely seen and fly mostly near the tree canopy but larvae or

‘larval eats’ [distinctive oblique leaf cuts] are more easily found on the food

plant, Lomandra spicata (Xanthorrhoeaceae), which grows on dark, damp

slopes and near river banks deep within the forest (Atkins 1999). It is

recorded from disjunct localities in SE Queensland but seems to be not

uncommon on the border ranges at Springbrook, Green Mountain and Binna

Burra. The food plant extends much further north, to the Atherton Tableland

in northern Queensland. In recent surveys I have found L. spicata in both

122 Australian Entomologist, 2007, 34 (4)

upland and lowland rainforests in the hinterlands of the Sunshine Coast, as

far north as 15 km NE of Kin Kin, and also in a small area (containing about

50 plants) on the slopes of Mt Dalrymple, Eungella Range west of Mackay

(possibly a new locality record for the plant). There were no larval eats at this

latter locality. Recent observations (2006-07) of larvae confirm the skipper’s

presence at Mt Mee, Conondale Ranges, Maleny, Montville and Mapleton,

with possible larval eats at Peachester and in the forests north of Kin Kin. A

further search of areas west of Miriam Vale might prove fruitful.

Signeta tymbophora

In February 2007, I observed and collected a male flying in bright sunshine

around and settling on 3 m high shrubs growing in a rainforest ravine in the

Mapleton State Forest. Its food plants are various species of forest wire-

grasses (Poaceae), Gahnia and Carex (Cyperaceae). A wire-grass was found

nearby. Lomandra spicata and Scleria sphacelata also grow in this area. This

is a new northern record for this small, dark skipper (probably about 20 km

north of the inland locality at Bunya Mts).

Hesperilla sarnia

As with T. genevieveae, observations of adults of this fast flying, dark brown

skipper are rare; however, larval leaf-tube shelters and zigzag cuts to the leaf

are more indicative of its presence. More than 15 localities, both lowland and

upland, are known from the Sunshine Coast and hinterland (Atkins 2004). In

2006, three eggs were found on Scleria sphacelata (Cyperaceae), two on one

plant at the same Mapletom forest locality as S. tymbophora and one on a

plant at Forest Glen. The eggs were all located on the upper side of leaves

close to the base of the plants. The eggs were 1 mm in diameter and had

approximately 20 vertical ribs; all three appeared to be infertile. In February

2007, a slightly worn female was collected resting on grass growing within a

new housing estate at Forest Glen. Compared with males, it was somewhat

reluctant to fly and, when disturbed, flew rather sluggishly. This is probably

the fifth record of a female of the southeastern form of this skipper.

References

ATKINS, A.F. 1999. The skippers, Trapezites (Hesperiidae). Chapter 5, pp 75-104, in: Kitching,

R.L., Scheermeyer, E., Jones, R.E. and Pierce, N.E. (eds), Biology of Australian butterflies.

Monographs on Australian Lepidoptera, Vol. 6. CSIRO Publishing, Collingwood; xvi + 395 pp.

ATKINS, A.F. 2004. New larval food plant records and notes on the biology of Trapezites

symmomus Hiibner, T. praxedes (Plötz), T. maheta (Hewitson) and Hesperilla sarnia Atkins

(Lepidoptera: Hesperiidae: Trapezitinae) from southeast Queensland. Australian Entomologist

31(4): 137-140.

BRABY, M.F. 2000. Butterflies of Australia: their identification, biology and distribution. 2

vols, CSIRO Publishing, Collingwood; xxvii + 976 pp.

Australian Entomologist, 2007, 34 (4): 123-125 123

LOPHODIPLOSIS TRIFIDA GAGNÉ (DIPTERA: CECIDOMYIIDAE),

A STEM-GALLING MIDGE WITH POTENTIAL AS A BIOLOGICAL

CONTROL AGENT OF MELALEUCA QUINQUENERVIA

(MYRTACEAE)

M. PURCELL’, S. WINEWRITER? and B. BROWN!

'CSIRO Entomology, United States Department of Agriculture, Agricultural Research Service,

Australian Biological Control Laboratory, 120 Meiers Road, Indooroopilly, Old 4068

(Email: matthew.purcell@csiro.au)

”United States Department of Agriculture, Agricultural Research Service, FDACS Florida

Biocontrol Laboratory, PO Box 147100, Gainesville, Florida 32614-7100, USA

Abstract

The gall midge Lophodiplosis trifida Gagné was originally described as an inquiline of galls

formed by three other Lophodiplosis Gagné species on Melaleuca dealbata and M.

quinquenervia. However, field observations conducted throughout the native range of M.

quinquenervia, coupled with replicated laboratory studies, have shown that L. trifida forms

unique stem galls on Melaleuca species within the M. leucadendron complex. Melaleuca

quinquenervia is an invasive weed in Florida, USA and L. trifida is a candidate biological

control agent of that species.

Introduction

Melaleuca quinquenervia S. T. Blake is a federally and state listed invasive

tree in southern Florida, USA (Turner et al. 1998). Since 1996, explorations

for host-specific natural enemies have been conducted by the United States

Department of Agriculture, Agriculture Research Service, Australian

Biological Control Laboratory (USDA-ARS ABCL) throughout the range of

M. quinquenervia in Australia.

A gall-forming cecidomyiid fly, then undescribed, was first collected in 1995

from M. quinquenervia growing in Queensland, by ABCL scientist J. K.

Balciunas. It was described subsequently as Lophodiplosis trifida Gagné by

Gagné et al. (1997), who reported it as an inquiline of three other

Lophodiplosis Gagné galls — in leaf blister galls with L. indentata Gagné and

L. denticulata Gagné on M. quinquenervia, and in bud rosette galls with L.

bidentata Gagné on M. dealbata S. T. Blake (Gagné et al. 1997).

Since then, ABCL scientists have collected L. trifida from unique stem galls

throughout the native range of M. quinquenervia. Specimens were sent to R.

J. Gagné, who confirmed their identity. Voucher specimens are held at the

Australian National Insect Collection, Canberra (ANIC) and the U.S.

National Museum of Natural History, Smithsonian Institution, Washington,

D.C.

Observations

Lophodiplosis trifida galls young shoots, predominantly during the autumn-

winter period, when a flush of young foliage is produced by M.

quinquenervia trees following flowering. The galls are variable in size and

shape and can persist on the plant for long periods, resulting in deformed

124 Australian Entomologist, 2007, 34 (4)

branches. Close or overlapping utilisation of host tissue by L. trifida, L.

indentata, L. denticulata and L. bidentata accounts for L. trifida’s original

designation as an inquiline in other Lophodiplosis galls. In the absence of

congeners, colonies of L. trifida have been established and sustained on

young seedlings and plants of M. quinquenervia for many generations at

ABCL. Lophodiplosis trifida readily galls the stems, curtailing shoot growth,

that can sometimes lead to death of the plants.

Discussion

The discovery that L. trifida is a stem galler of M. quinquenervia, rather than

an inquiline, is significant, given that this particular gall possesses some of

the traits considered desirable for biological control of weeds (Dennill 1988,

Harris and Shorthouse 1996). The larvae of L. trifida live within the gall, the

galled shoots occur at high densities, gall development spans the entire

growth phase of the plant and gall development severs vascular tissue.

As part of a management plan for control of M. quinquenervia, three

biological control agents have been introduced to Florida (USA) by scientists

at the USDA-ARS Invasive Plant Research Laboratory (IPRL) since 1997,

including a bud gall fly Fergusonina turneri Taylor (Goolsby et al. 2000,

Davies et al. 2001). However, additional natural enemies are required that

attack other plant stages. Elsewhere, gall formers have been used extensively

in weed biological control programs (Julien and Griffiths 1998).

Initial screening of non-Melaleuca myrtaceous species by ABCL indicated

that the host range of L. trifida is limited to Melaleuca species in the M.

leucadendron complex. As a result, researchers of the IPRL and ABCL

selected L. trifida to be imported and subjected to additional host range

testing at the Florida Department of Agriculture and Consumer Services,

Division of Plant Industry (FDACS DPI) quarantine facility in Gainesville,

Florida, USA, in 2003.

Acknowledgements

We wish to thank ABCL staff members Jeff Makinson, Ryan Zonneveld and

Dalio Mira for their assistance with field collections and observations, and Dr

R. J. Gagné (USDA-ARS) for identification of specimens.

References

DAVIES, K.A., MAKINSON, J. and PURCELL, M.F. 2001. Observations on the development

and parasitoids of Fergusonina/Fergusobia galls on Melaleuca quinquenervia (Myrtaceae) in

Australia. Transactions of the Royal Society of South Australia 125: 45-50.

DENNILL, G.B. 1988. Why a gall former can be a good biocontrol agent: the gall wasp

Trichilogaster acaciaelongifoliae and the weed Acacia longifolia. Ecological Entomology 13: 1-

9.

GAGNE, R.J., BALCIUNAS, J.K. and BURROWS, D.W. 1997. Six new species of gall midges

(Diptera: Cecidomyiidae) from Melaleuca (Myrtaceae) in Australia. Proceedings of the

Entomological Society of Washington 99: 312-334.

Australian Entomologist, 2007, 34 (4) 125

GOOLSBY, J.A., MAKINSON, J.R. and PURCELL, M.F. 2000. Seasonal phenology of the

gall-making fly Fergusonina sp. (Diptera: Fergusoninidae) and its implications for biological

control of Melaleuca quinquenervia. Australian Journal of Entomology 39: 336-343.

HARRIS, P. and SHORTHOUSE, J.D. 1996. Effectiveness of gall inducers in weed biological

control. Canadian Entomologist 128: 1021-1055.

JULIEN, M.H. and GRIFFITHS, M.W. (eds.). 1998. Biological control of weeds: a World

catalogue of agents and their target weeds. 4th edition. CAB International, Wallingford, United

Kingdom; 223 pp.

TURNER, C.E., CENTER, T.D., BURROWS, D.W. and BUCKINGHAM, G.R. 1998. Ecology

and management of Melaleuca quinquenervia, an invader of wetlands in Florida, U.S.A.

Wetlands Ecology and Management 5: 165-178.

126 Australian Entomologist, 2007, 34 (4)

NOTES ON THE GENUS-GROUP PLACEMENT OF

PENEPAROXYNA HARDY & DREW AND SORAIDA HERING

(DIPTERA: TEPHRITIDAE: TEPHRITINAE)

D.L. HANCOCK

PO Box 2464, Cairns, Qld 4870

Abstract

The Australian genus Peneparoxyna Hardy & Drew is transferred from the Tephritis group to the

Campiglossa group of genera in tribe Tephritini, while the Indonesian genus Soraida Hering is

newly placed in the Campiglossa group.

Peneparoxyna Hardy & Drew

Peneparoxyna minuta Hardy & Drew, known from New South Wales and the

Northern Territory (Hardy and Drew 1996), was placed in the Tephritis group

of genera by Hancock (2001) and Hancock and Drew (2003), largely on the

basis of its superficial resemblance to Actinoptera Rondani. However, the

narrow, geniculate mouthparts, weakly reticulate wing pattern and internal

structure of the male distiphallus are more consistent with the Campiglossa

group of genera (especially Dioxyna Frey, Desmella Munro and Tanaica

Munro: see Merz and Dawah 2005), to which it is transferred.

Soraida Hering

Soraida tenebricosa Hering, known from Lombok and Sunda Is in Indonesia

(Hering 1941), was included in the tribe Tephritini by Hardy (1988) but its

precise relationships remained unresolved. It differs from all other Indo-

Australian members of the subfamily Tephritinae in its wing pattern, being

smoky-grey to pale brownish with a pale brown pterostigma but without

hyaline markings. Its generic characters (see Hardy 1988), particularly the

geniculate mouthparts, are typical of the Campiglossa group of genera to

which it is referred. Soraida and Peneparoxyna are the only known genera of

the group restricted to the Indo-Australian Region.

References

HANCOCK, D.L. 2001. Systematic notes on the genera of Australian and some non-Australian

Tephritinae (Diptera: Tephritidae). Australian Entomologist 28(4): 111-116.

HANCOCK, D.L. and DREW, R.A.I. 2003. A new genus and new species, combinations and

records of Tephritinae (Diptera: Tephritidae) from Australia, New Zealand and the South Pacific.

Australian Entomologist 30(4): 141-158.

HARDY, D.E. 1988. The Tephritinae of Indonesia, New Guinea, the Bismarck and Solomon

Islands (Diptera: Tephritidae). Bishop Museum Bulletin in Entomology 1: i-vii, 1-92.

HARDY, D.E. and DREW, R.A.I. 1996. Revision of the Australian Tephritini (Diptera:

Tephritidae). Invertebrate Taxonomy 10: 213-405.

HERING, E.M. 1941. Dipteren von den Kleinen Sunda-Inseln. Aus der Ausbeute der Sunda-

Expedition Rensch. II. Trypetidae. Arbeiten über Morphologische und Taxonomische

Entomologie aus Berlin-Dahlem 8: 24-45.

MERZ, B. and DAWAH, H.A. 2005. Fruit flies (Diptera, Tephritidae) from Saudi Arabia, with

descriptions of a new genus and six new species. Revue suisse de Zoologie 112(4): 983-1028.

Australian Entomologist, 2007, 34 (4): 127-139 127

SONGS AND CALLING BEHAVIOUR OF FROGGATTOIDES

TYPICUS DISTANT (HEMIPTERA: CICADOIDEA: CICADIDAE),

A NOCTURNALLY SINGING CICADA

A. EWART! and L.W. POPPLE?

‘Queensland Museum, PO Box 3300, South Brisbane, Old 4101

”School of Integrative Biology, The University of Queensland, St Lucia, Qld 4072

Abstract

The nocturnal male singing behaviour and songs of Froggattoides typicus Distant are

documented, based on observations and sound recordings made at the Southwood National Park

in southern Queensland, during early December 2005, in an open net after dusk. Two distinct

song components are recognized: (i) a continuous, soft clicking song (calling song) with some

accompanying clicks and click phrases, emitted during the earlier part of the evening and

believed to be predominantly timbal produced; (ii) sets of multiple ticks produced during the

later part of the evening, commonly accompanying wing flicking behaviour and together with

sporadically emitted, short, sharp buzz phrases which sound similar to the sudden expulsion of

air from a restricted nozzle.

Introduction

Known by its popular name ‘eastern bent-winged cicada’ (Moulds 1990),

Froggattoides typicus Distant is a very distinctive, predominantly pale green,

endemic Australian cicada. Moulds (1990) noted that it had never been heard

singing, at least during the day. It is, however, frequently captured at light,

usually arriving well after sunset, when it commonly appears in significant

numbers and often emits a marked clicking noise. As noted by Moulds

(1990), hand-held specimens of both sexes produce an audible clicking noise,

apparently resulting from a rapid beating together of the distal half of the

wings while closed. The production of click sounds in this species may,

however, also involve an alternative mechanism as the wings are flicked

open, generated when the forewing leaves the wing grooves on the margins

of the mesonotum (Ewing 1989; see also Gogala and Trilar 2003).

F. typicus occurs widely throughout southern, southwestern and central

Queensland, being noticeably common in forest communities associated with

brigalow (Acacia harpophylla) and gidyea (A. cambagei) woodlands. It is

very rarely seen during the day, sitting camouflaged in tree foliage. We were

initially alerted to the nocturnal behaviour of F. typicus by observations made

of insects calling after 2000 h on the highway between Charleville and

Cunnamulla, southern Queensland (S. Peck, pers. comm.). Our observations

indicate that it is indeed nocturnally active, with song production only after

dusk. Such behaviour is unusual within known Australian Cicadidae, which

predominantly sing during the day and/or at dusk (see Moulds 1990).

Materials and methods

Observations and recordings were collected in brigalow forest in the

Southwood National Park (~27°50'S 150°06'E), southern Queensland,

128 Australian Entomologist, 2007, 34 (4)

between 5-9 December 2005. Recordings were made with a Sony cassette

recorder WM-D6C, with Sennheiser microphone K6/ME66. Song analyses

utilised Avisoft SASLab Pro4 software. After capture at light, the insects

were placed in a cylindrical net cage (36 cm x 30 cm diameter), containing

small fragments of local vegetation, and placed in a canvas tent (3 x 2 x 2 m)

with no artificial lighting but with one window uncovered to allow entry of

filtered weak moonlight. Some of the insects were kept alive in the net cage

during the following day under strongly shaded conditions. Song and timbal

terminology follows that used by Ewart (2005). Amplitude spectra were

produced using a 556-point Fast Fourier Transform with Hamming window.

Singing behaviour

During and especially after dusk the cicadas became active, constantly

moving around the vegetation with frequent wing ‘flicking’. This activity

continued until or slightly beyond midnight. The nature of ‘song’ production,

however, changed between the earlier and later parts of the evening. During

the observation period, sunset and end of civil twilight occurred at

approximately 1848 h and 1915 h (Eastern Standard Time) respectively.

From approximately 2030-2115 h the cicadas, while still actively walking

around the vegetation in the cage, produced a continuous, soft clicking song

(Fig. 1), sporadically interspersed with sets of regularly emitted and regularly

spaced clicks as well as short, sharp and relatively loud individual clicks

(Figs 2A-C), in some cases visibly associated with wing flicking. After

approximately 2115 h, the soft clicking song became progressively more

subdued and ceased. Instead, the cicadas continued to be active, constantly

(but erratically) flicking their wings, which produced very short sets of

distinct multiple clicks (Fig. 3A), not exactly the same in structure as those

emitted earlier, together with additional and very sporadic short sharp ‘buzz’

phrases (Fig. 3D); this activity continued to slightly beyond midnight. All

song types recorded during this study were produced by male insects only.

Fig. 1. Waveform plots (amplitude versus time) of the early evening calling song of

Froggatoides typicus. (A): the continuous echeme sequences, each echeme sounding

as a soft click; the low amplitude phrases occurring between some of the echemes are

due to the songs of other F. typicus in the background; recording filtered (IIR) to 5.5

kHz. (B): expanded time plot showing the multiple macrosyllables comprising each

echeme, the numbers associated with each echeme showing the numbers of

macrosyllables present; the definitions of echeme lengths and repetition rates are

shown [as also in A]. (C): further time expanded plot of a single echeme showing

individual macrosyllables, each comprising multiple syllables. (D): details of

individual syllables within the initial macrosyllable shown in Fig. C; also note the

long final syllable, possibly indicating syllable coalescence. Records B to D filtered

(IIR) to 1 kHz.

Australian Entomologist, 2007, 34 (4) 129

Continuous echeme sequences (soft clicks)

hI

i

|

| i

Single macrosyllable with at least

20 distinct syllables

130 Australian Entomologist, 2007, 34 (4)

20 40 60 80 100 120 140 160 180 200 220 240 260

400— Ẹ Single discrete clicks

DA a

200

100

0

-100

-200

-300-

-400

40 80 120 160 200 240 280 320 360 400ms

Fig. 2. Waveform plots (A, C, E) and their respective amplitude spectra (B, D, F;

amplitude vs frequency) of early evening clicking sounds of Froggatoides typicus.

(A-B): three groups of single clicks, each group with changing repetition rates,

emitted between calling song echeme; individual clicks have broadband frequency

spectra with dominant frequencies between 8.4 and 11.0 kHz; amplitude spectrum

shown is based on all three groups. (C-F): temporally patterned clicks emitted at

beginning and end of a sequence of calling song echemes; individual clicks include

both single and double pulses; additional higher amplitude isolated single clicks are

shown, which may not have been emitted by the same insect; these clicks are single

ps with narrowband frequency spectra and dominant frequencies between 6.9 and

Z.

Australian Entomologist, 2007, 34 (4) 131

6001 A Later evening sounds showing examples of sets of See Fig. D below

my multiple clicks and sharp buzz phrase

See Fig. C below See Fig. B below

E j

-200 -

24 ES

Sets of multiple Sets of multiple

clicks clicks

=T T

0 1 2 3 4 5 6 7 s

Expanded time plot of a single set of

multiple clicks

0 10 20 30 40 50 60 70 80 ms

Expanded time plot of a single set of

multiple clicks

0 10 20 30 40 50 60 70 80 90 ms

Expanded time plot

of a single sharp

buzz phrase

200 240 280 320 360 ms 400

Fig. 3. Waveform plots of later evening clicks and ‘buzz’ sounds of F. typicus. (A):

sets of multiple sets of clicks and the short sharp ‘buzz’ phrase. (B-C): time expanded

plots of two sets of clicks shown in Fig. A. (D): expanded time plot of the single sharp

‘buzz’ phrase shown in Fig. A. Recordings A to D filtered (IIR) to I kHz.

132 Australian Entomologist, 2007, 34 (4)

Song analyses

(a) Early evening continuous soft clicking song (Fig. 1)

This consisted of continuous echeme sequences, each echeme producing a

single audible click. The mean repetition rate was 1.35 s” (range 1.2-1.5) and

mean echeme length was 325 ms (range 267-378). Each echeme, as seen in

expanded time plots (Figs 1B-C), comprised a sequence of macrosyllables, 8-

12 in number, with a mean length of 21.7 ms (range 16.4-30.3), the variation

reflecting the number of component macrosyllables. Macrosyllable repetition

rates varied from 24-37 s' (27.9-41.5 ms). The initial macrosyllable in each

echeme was the longest. Each macrosyllable was further resolved, at further

time expansion (Fig. 1D), into 16-24 discrete syllables. Syllable repetition

rates varied from 295-2040 s (mean 910), equivalent to syllable lengths of

0.49-3.4 ms (mean 1.13); these equated closely to the syllable amplitude

modulation of 885 s”. The final two high amplitude syllables within each

macrosyllable apparently have coalesced, as illustrated by the macrosyllable

shown in Fig. 1D.

The amplitude spectrum (Fig. 4A) of the continuous clicking song component

shows the emitted frequency maxima to lie between approximately 7 and 9

kHz, with significant frequency peaks, with reduced magnitudes, extending

to 16 kHz. The spectrum is notable for the extensive array of apparent

sidebands (listed in Fig. 4A), their complexity attributed to the complexity of

the fine scale variability within the macrosyllable and syllable structures.

Very detailed time plots and amplitude spectra of the syllables (not shown)

indicated that the final high amplitude syllables (as shown in Fig. 1D) are

characterised by a rather narrow band of frequency emission centred at 7.2

kHz, the frequencies slightly decreasing during syllable emission and rising

markedly either side of the syllable. In contrast, the initial 5 syllables shown

in Fig. 1D had frequencies between 9.4 and 10.1 kHz, compared with the low

amplitude inter-syllable regions which exhibited higher frequencies between

10.6 and 11 kHz. These demonstrate rapid temporal changes in frequencies

during syllable emission, on time scales of <1 ms.

(b) Early evening click phrases and single clicks

Short phrases, less than 300 ms in length and comprised of temporally

structured clicks, were observed to be sporadically interspersed within, or

between, the echemes of the continuous soft clicking song (Fig. 2). The

regularly emitted click phrases (Figs 2C-D) had click repetition rates of 145-

146 s` and comprised both single and doublet pulses. When emitted as short

groups of clicks (each single pulses: Fig. 2A), the click repetition rates

decreased during emission. The frequency structures of these structures click

phrases are variable (Figs 2B, D, F), ranging from relatively sharply tuned

frequency maxima near 8 kHz (Fig. 2F), to more broadband frequency

maxima between about 8 and 11.5 kHz (Fig. 2A), to very broadband

frequency distributions between 2 and 11 kHz (Fig. 2D).

Australian Entomologist, 2007, 34 (4) 133

Continuous soft , H Apparent sidebands: 429,

clicking song 332, 276, 221, 147, 128,

111, 92, 64, 51, 38, 32,

(Early evening) Å 23. 15.8,6,4.3 Hz

Background

cricket

(4.2-5.2 kHz)

rt

Single set E E Single set

multiple clicks 8 multiple clicks

(initial segment; |3. (second segment;

see Fig. 3B) 97 see Fig. 3B)

See Fig. 3D

Single short

Fig. 4. Amplitude spectra of Froggatoides typicus songs. (A): multiple echemes

(40 seconds) of the continuous soft calling song shown in Fig. 1A; note the weak

background song of an unidentified cricket between 4.2 and 5.1 kHz. (B-C): initial

and later segments, respectively, of the set of clicks shown in the expanded time plot

in Fig. 3B. (D): set of multiple clicks shown in the expanded time plot in Fig. 3C. (E):

The short ‘buzz’ phrase shown on Fig. 3D. Spectra shown in Figs C and D filtered

(IIR) to 1 kHz.

134 Australian Entomologist, 2007, 34 (4)

The relatively higher amplitude sharp single clicks (Figs 2C, E) are emitted

apparently randomly during the continuous soft clicking song. Expanded time

plots (not shown) showed these to be narrowly tuned single pulses with

relatively narrow dominant frequency spectra between 6.9 and 7.9 kHz.

Those shown in Figs 2C-D may not all emanate from a single insect in the

cages.

(c) Later evening sets of multiple clicks (Fig. 3A)

These are here linked to wing ‘flicking’ behaviour accompanying the

constant walking activity. The timing between the sets of clicks was very

variable although they were emitted frequently. Individual sets of clicks

varied in detailed structure, as seen in expanded time plots (Figs 3B-C).

Within a given set, individual clicks ranged from 2 to 12 in number. Their

structures also varied, some comprising high amplitude pulses with

logarithmically decaying tail, others that were relatively closely spaced and

partially coalescing, with extended complex decaying tail (Fig. 3B). The

initiation of the high amplitude pulse trains of the clicks were mostly abrupt

and were commonly preceded (~5-10 ms interval) by a sharp, low amplitude

pulse or pulses. The two sets of amplitude spectra illustrated (Figs 4B-D)

exhibit strongly contrasting frequency patterns, particularly in the click

sequence shown in Fig. 3B. The initial segment of this sequence had a

narrowband frequency structure between 6.5 and 8.5 kHz (Fig. 4B). The

following segment of the click sequence (Fig. 4C) exhibited broadband

frequency emissions between <l and 10 kHz, extending with reduced

amplitude to 15 kHz. The most significant difference between these two

spectra was the presence of a strong lower frequency component below 5

kHz in the later segment of the ticking sequence, which we attribute to wing

flicking (see below). The amplitude spectrum of the click sequence shown in

Fig. 3C exhibits an extremely broad frequency emission extending between

<1 and 16 kHz (Fig. 4D).

(d) Later evening short sharp ‘buzz’ phrases (Fig. 3D)

These clearly differ from the sets of clicks previously described. Those

measured ranged between 0.3 and 0.35 s in length. They are abrupt, relatively

loud and emitted only sporadically and irregularly. To the human ear, they

have a distinct resemblance to the sudden expulsion of air from a restricted

nozzle. As shown by the waveform plots (Fig. 3D), they initiate abruptly,

continue briefly at constant amplitude and decay nearly exponentially. A

small precursor double pulse is present. The amplitude spectra showed a

broad frequency emission range between approximately 2 and 10 kHz, with

maxima near 7-7.5 kHz (Fig. 4E). There appeared to be an absence of clearly

defined temporal patterning and thus the overall structure was similar to

white noise. The small precursor pulse phases exhibited a very narrowly

ned spectrum between 6.7 and 8.1 kHz, quite distinct from that of the main

phrase.

Australian Entomologist, 2007, 34 (4) 135

Discussion

The early evening continuous soft clicking song is interpreted as a timbal-

generated calling song with a dominant frequency range between ~6.5 and 9

kHz. Although a cursory examination of the timbals suggest that they are

poorly developed, this is only because they are effectively ‘sandwiched’

between the bulbous tergites 1 and 2 (Figs 5B, 6B) and require the removal of

much of tergite 2 to be fully visible (Fig. 5A). They are similar in overall

form to those of diurnally singing cicadas. The six timbal ribs (long ribs) are

very pale green in colour and weakly sclerotised. The four posterior ribs (1 to

4) are fused dorsally to the basal spur and ventrally to each other. Ribs 1 to 3

are continuous across the timbal, whereas rib 4 is discontinuous medially.

The two most anterior ribs (5, 6) are short, unfused and appear to represent

remnant long ribs. No inter-rib sclerites between ribs 1 to 4 were observed.

The dorsal and ventral fusion of ribs 1 to 4 suggest that these may act as a

single rib during timbal contractions and relaxation, as suggested in certain

diurnal ticking cicadas (Ewart 2005). The male opercula are well developed

and also similar in overall structure to those of diurnal cicadas (Fig. 6A).

Those in F. typicus are notable for the absence of spikes on the meracantha

and the marked curvature and undulations of the surfaces of the opercula, the

latter best seen in lateral profile (Fig. 6B).

The various clicking sounds shown above exhibit complex frequency spectra

varying between narrowband and very broadband. As described, wing

flicking occurs frequently during early and later evening songs. Some clicks

were observed to correlate with wing flicks, suggesting that both timbals and

wing flicks are used, either singly or in combination, as part of song

production. An additional factor is the differential role of sound radiation

structures (timbals, tympana and abdomen) in modulating the frequency

signatures of the emitted sounds (Fonseca and Popov 1994).

We suggest that the earlier evening songs are predominantly timbal produced,

including the patterned and separate click components, which we further

correlate with dominant spectral frequencies between approximately 6-11

kHz (e.g. Figs 2B, 2F; 4A, 4B). Nevertheless, many clicks have dominant

frequencies which include a significant component at <6 kHz and in some

clearly extend to >11 kHz (e.g. Figs 4C-D). We interpret these clicks and

their spectral frequencies to have originated predominantly through wing

flicking. The later evening clicks are mainly, but not entirely, of these types.

Nevertheless, amplitude spectra of some clicks, including early and later

evening types, showed frequencies which suggest the presence of both timbal

and wing clicking components (e.g. Figs 2D, 4B-C), pointing to combined

timbal and wing flicking in sound production. The short sharp ‘buzz’ phrases

are enigmatic in their origin. Both their structure, as seen in the waveform

plots, and their frequency spectra distinguish them from the other sounds

produced by F. typicus, implying a different production mechanism. As noted

136 Australian Entomologist, 2007, 34 (4)

above, the sound resembles a sharp expulsion of air. The frequency spectrum

of the precursor pulses, however, suggests that these may initiate via a timbal

origin.

Dorsal

Anterio-

lateral

margin

Timbal

Fig. 5. Froggattoides typicus. (A): view of right timbal. (B): view of dorsal surface

between tergites I and 2 showing the position of the timbals ‘sandwiched’ between

the tergites. Scale bars = 1 mm.

Australian Entomologist, 2007, 34 (4) 137

Fig. 6. Froggattoides typicus. (A): ventral surface of left operculum. (B): lateral view

of left operculum (Op) and proximal structures. Tymp = tympanum (graded shading);

Aud Cap = auditory capsule; FW and HW = bases of fore and hind wings; Tg 1-3 =

tergites 1, 2 and 3; Timb = timbal (‘sandwiched’ between tergites 1 and 2). Two

external openings are shown as the shaded areas. Scale bars = 1 mm.

138 Australian Entomologist, 2007, 34 (4)

Froggattoides typicus seemingly exhibits certain behavioral and song

adaptations to nocturnal activity. First is the constant movement at night

within tree foliage, observed as walking in the cages, but is also inferred to

include at least short flights in the natural environment (noting their ready

attraction to light). This activity starts during dusk, again based on

observations on caged specimens. A second aspect is the soft continuing

clicking song, here identified as the calling song, emitted during the earlier

part of the evening. This song has a dominant frequency range of 6.5-9 kHz,

with a weaker extension to about 14 kHz. Additional patterned and single

clicks are emitted during this song component, whose frequencies overlap,

even slightly extending the range of the continuous calling song. These early

evening songs are inferred to be predominantly timbal produced, although

some clicks seem to have a wing-flicking component.

The third aspect is the change towards the increasing importance of clicking

songs later in the evening, tending to differ in their structures and frequency

properties from those emitted earlier. These clicks appear to be produced

predominantly through wing flicking, although some have a timbal

component. Sound production is again accompanied by movements of the

cicadas. These clicks, as described, have very wide frequency ranges, from

<2 to ~16 kHz, believed to facilitate sound transmission and localisation.

Wing flicking may also be associated with pheromone dispersal, although

this remains to be demonstrated. A fourth aspect is the production of the

sporadic short, sharp ‘buzz’ song, which occurs both in the early and

especially later evening. The dominant frequency of this component lies

between approximately 2 and 10 kHz, thereby effectively complementing the

frequency ranges of the other emitted sounds.

A fifth aspect concerns sound interference during nocturnal singing. During

the period of the present observations, the only interference encountered was

a continuous song of an unidentified cricket. The frequency of this song lies

within the narrow range of 4.2-5.1 kHz (Fig. 4A). This is below the dominant

frequency of the F. typicus calling song and only minimally overlaps with

that of the other two song types.

One significant feature of the variety of sounds emitted by F. typicus is their

resulting broad frequency range. Such frequency ranges are expected to

facilitate more efficient survival of song structures which are degraded

through absorption, scattering, reflection and refraction, which lead to

frequency filtering of sound propagating through foliage (Michelsen 1992,

Richards and Wiley 1980, Michelsen and Larsen 1983, Römer and Lewald

1992). The higher frequency components should also aid in sound

localisation (Gerhardt and Huber 2002). It is suggested that the early evening

calling songs and clicking sounds function to alert and attract females to the

presence of males within a given area. The later evening clicking and ‘buzz’

sounds may facilitate the final stages of localisation of both males and

Australian Entomologist, 2007, 34 (4) 139

females. It is likely that females also use wing flicking to respond to male

calls (given that female F. typicus also possess strongly angulated forewings),

as has been observed in a number of diurnal cicada species (Sueur and Aubin

2004).

Acknowledgements

Thanks are due to the Rangers and Officers of the Southern Region of the

Queensland Parks and Wildlife Service, especially Stephen Peck for alerting

us to the night songs of F. typicus and Rod Hobson for organising a fauna

survey of the Southwood National Park, during which the above observations

were made. Dr M.S. Moulds, two anonymous referees and Kathy Hill and D.

Marshall (University of Connecticut, U.S.A.) are thanked for their

constructive comments and observations on the manuscript.

References

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Cicadoidea: Cicadidae) from Queensland, with descriptions of their songs. Memoirs of the

Queensland Museum 51(2): 439-500.

EWING, A.W. 1989. Arthropod bioacoustics: neurobiology and behaviour. Edinburgh

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FONSECA, P.J. and POPOV, A.V. 1994. Sound radiation in a cicada: the role of different

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GERHARDT, H.C. and HUBER, F. 2002. Acoustic communication in insects and anurans:

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GOGALA, M. and TRILAR, T. 2003. Video analysis of wing clicking in cicadas of the genera

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Slovenica 11(1): 5-15.

MICHELSEN, A. 1992. Hearing and sound communication in small animals: evolutionary

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MICHELSEN, A. and LARSEN, O.N. 1983. Strategies for acoustic communication in complex

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RECENT ENTOMOLOGICAL LITERATURE

BRABY, M.F.

2007 Collecting biological specimens in the Northern Territory with particular reference to

terrestrial invertebrates: guidelines to current legislation and permits. Northern Territory

Naturalist 19: 35-45.

BRABY, M.F., PIERCE, N.E. and VILA, R.

2007 Phylogeny and historical biogeography of the subtribe Aporiina (Lepidoptera: Pieridae):

implications for the origin of Australian butterflies. Biological Journal of the Linnean

Society 90: 413-440.

GRIMSHAW, J.F. and DONALDSON, J.F.

2007 Records of two sugarcane pests Eumetopina flavipes Muir (Hemiptera: Delphacidae) and

Chilo terrenellus Pagenstecher (Lepidoptera: Pyralidae) from Torres Strait and far north

Queensland. Australian Journal of Entomology 46(1): 35-39.

KOLESIK, P., WOODS, B., CROWHURST, M. and WIRTHENSOHN, M.G.

2007 Dasineura banksiae: a new species of gall midge (Diptera: Cecidomyiidae) feeding on

Banksia coccinea (Proteaceae) in Australia. Australian Journal of Entomology 46(1): 40-

44,

LOCKER, B., FLETCHER, M.J. and GURR, G.M.

2007 Revision of the genus Innobindus Jacobi (Hemiptera: Fulgoromorpha: Cixiidae) with the

description of six new species and comments on other Australian Brixiini genera.

Australian Journal of Entomology 46(1): 45-55.

LOCKER, B., FLETCHER, M.J., GURR, G.M., HOLZINGER, W.E. and LOCKER, H.

2006 Taxonomic and phylogenetic revision of the Gelastocephalini (Hemiptera: Cixiidae).

Invertebrate Systematics 20: 59-160.

PRINSLOO, G.L. and NESER, O.C.

2007 Revision of the pteromalid wasp genus Trichilogaster Mayr (Hymenoptera:

Chalcidoidea): gall-inducers on Australian acacias. African Entomology 15(1): 161-184.

SCRIBER, J.M., LARSEN, M.L. and ZALUCKI, M.F.

2007 Papilio aegeus Donovan (Lepidoptera: Papilionidae) host plant range evaluated

experimentally on ancient angiosperms. Australian Journal of Entomology 46(1): 65-74.

[incorrectly records subsp. P. a. ormenus from Victoria and South Australia]

SOULIER-PERKINS, A.

2005 Phylogenetic evidence for multiple invasions and speciation in caves: the Australian

planthopper genus Solonaima (Hemiptera: Fulgoromorpha: Cixiidae). Systematic

Entomology 30: 281-288.

THEISCHINGER, G. and HAWKING, J.

2006 The complete field guide to dragonflies of Australia. CSIRO Publishing, Melbourne.

WATTS, C.H.S. and HUMPHREYS, W.F.

2004 Thirteen new Dytiscidae (Coleoptera) of the genera Boongurrus Larson, Tjirtudessus

Watts & Humphreys and Nirripirti Watts & Humphreys, from underground waters in

Australia. Transactions of the Royal Society of South Australia 128: 99-120.

2006 Twenty-six new Dytiscidae (Coleoptera) of the genera Limbodessus Guignot and

Nirripirti Watts & Humphreys, from underground waters in Australia. Transactions of the

Royal Society of South Australia 130: 123-185.

WATTS, C.H.S., HANCOCK, P.J. and LEYS, R.

2007 A stygobitic Carabhydrus Watts (Dytiscidae, Coleoptera) from the Hunter Valley in New

South Wales, Australia. Australian Journal of Entomology 46(1): 56-59.

THE —

AUSTRALIAN

ENTOMOLOGIST

VOLUME 34

2007

Published by:

THE ENTOMOLOGICAL SOCIETY OF QUEENSLAND

THE AUSTRALIAN ENTOMOLOGIST

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THE AUSTRALIAN ENTOMOLOGIST

Contents

Volume 34, 2007

ATKINS, A.

Notes on the distribution and biology of Trapezites genevieveae

Atkins, Signeta tymbophora (Meyrick & Lower) and Hesperilla

sarnia Atkins (Lepidoptera: Hesperiidae)

EWART, A. and POPPLE, L.W.

Songs and calling behaviour of Froggattoides typicus Distant

(Hemiptera: Cicadoidea: Cicadidae), a nocturnally singing cicada

FERGUSON, D.J.

Field observations of Perissomma mcalpinei Colless (Diptera:

Perissommatidae)

GINN, S.G., BRITTON, D.R. and BULBERT, M.W.

New records of butterflies (Lepidoptera) in the Pilbara region of

Western Australia, with comments on the use of malaise traps

for monitoring

HANCOCK, D.L.

Phylogeny of the troidine butterflies (Lepidoptera: Papilionidae)

revisited: are the red-bodied swallowtails monophyletic?

The identity of ‘Trypeta’ nigricans Wiedemann (Diptera:

Tephritidae: Tephritinae)

A new synonym and a new combination in the fruit fly tribe

Pliomelaenini (Diptera: Tephritidae: Tephritinae)

A note on the genus Hemiristina Permkam & Hancock (Diptera:

Tephritidae: Trypetinae)

A review of the fruit fly tribe Eutretini (Diptera: Tephritidae:

Tephritinae) in the Indo-Australian region

A review of Callistomyia Bezzi and related genera (Diptera:

Tephritidae: Trypetinae)

The identity of Terellia immaculata Macquart (Diptera:

Tephritidae: Tephritinae)

Notes on the genus-group placement of Peneparoxyna Hardy &

Drew and Soraida Hering (Diptera: Tephritidae: Tephritinae)

HARDY, N.B. and GULLAN, P.J.

A new species of Phacelococcus Miller (Hemiptera: Coccoidea:

Eriococcidae) on Bursaria (Pittosporaceae)

HOPKINSON, M. and HOPKINSON, A.

A range extension for Graphium aristeus parmatum (Gray)

(Lepidoptera: Papilionidae) in northern Queensland

JOHNSON, S.J. and VALENTINE, P.S.

A new subspecies of Jalmenus inous Hewitson (Lepidoptera:

Lycaenidae) from Shark Bay, Western Australia

iii

121

127

93

65

33

49

57

84

92

107

119

126

85

76

77

KOHOUT, R.J.

A new species of the subgenus Polyrhachis (Cyrtomyrma) Forel

(Hymenoptera: Formicidae: Formicinae) from Borneo

LAMBKIN, T.L. !

The immature stages of Cephrenes moseleyi (Butler) (Lepidoptera:

Hesperiidae) from Torres Strait, Queensland

LAMBKIN, T.L. and KNIGHT, A.I.

Confirmation of Euploea leucostictos (Gmelin) and E. netscheri

erana (Fruhstorfer) (Lepidoptera: Nymphalidae) in Torres Strait,

Queensland, and the first record of E. tulliolus dudgeonis

(Grose-Smith) in Australia

MAGINNIS, T.L. and MAGINNIS, L.P.

Leg autotomy and regeneration in a population of Didymuria

violescens (Leach) (Phasmatodea: Phasmatidae) in New South

Wales, Australia

MOUND, L.A. and TREE, D.J.

Oriental and Pacific Thripidae (Thysanoptera) new to Australia,

with a new species of Pseudodendrothrips Schmutz

OLIVE, J.C.

Å new species of Gudanga Distant (Hemiptera: Cicadidae) from

northern Queensland

PURCELL, M., WINEWRITER, S. and BROWN, B.

Lophodiplosis trifida Gagné (Diptera: Cecidomyiidae), a stem-

galling midge with potential as a biological control agent of

Melaleuca quinquenervia (Myrtaceae)

SMITH, A.B.T.

Review of the genus Dungoorus Carne (Coleoptera: Scarabaeidae:

Rutelinae: Anoplognathini)

WHITEHOUSE, M.E.A. and GRIMSHAW, J.F.

Distinguishing between lynx spiders (Araneae: Oxyopidae) relevant

to IPM in cotton in the Namoi Valley, New South Wales

115

27

123

43

97

BOOK REVIEWS 56, 58, 61

RECENT LITERATURE

Publication dates: Part 1 (pp 1-32) 1 March 2007

Part 2 (pp 33-64) 25 May 2007

Part 3 (pp 65-96) 10 September 2007

Part 4 (pp 97-140) 10 December 2007

63, 140

ENTOMOLOGICAL NOTICES

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THE AUSTRALIAN

len

å "Volume 34, Part 4, 10 December 2007

CONTENTS

ATKINS, A.

Notes on the distribution and hoe of Trapezites genevieveae Atkins,

Signeta tymbophora (Meyrick & Lower) and Hesperilla sarnia

Atkins (Lepidoptera: Hesperiidae).

EWART, A. AND POPPLE, L.W.

Songs and calling behaviour of Froggattoides typicus Distant

(Hemiptera: Cicadoidea: Cicadidae), a nocturnally singing cicada.

HANCOCK, D.L.

A review of Callistomyia Bezzi and related genera (Diptera: Tephritidae: Trypetinae).

HANCOCK, D.L.

The identity of Terellia immaculata Macquart (Diptera: Tephritidae: Tephritinae).

HANCOCK, D.L.

Notes on the genus-group placement of Peneparoxyna Hardy & Drew and

Soraida Hering (Diptera: Tephritidae: Tephritinae).

KOHOUT, RJ.

A new species of the subgenus Polyrhachis (Cyrtomyrma) Forel

(Hymenoptera: Formicidae: Formicinae) from Borneo.

PURCELL, M., WINEWRITER, S. AND BROWN, B.

Lophodiplosis trifida Gagne (Diptera: Cecidomyiidae), a stem-galling midge with

potential as a biological control agent of Melaleuca quinquenervia (Myrtaceae).

WHITEHOUSE, M.E.A, AND GRIMSHAW, J.F.

Distinguishing between lynx spiders (Araneae: Oxyopidae) relevant

to IPM in cotton in the Namoi Valley, New South Wales.

RECENT ENTOMOLOGICAL LITERATURE

ISSN 1320 6133 GYT O