Australian Journal of Entomology (2008) 47, 153–159
Diverse cuticular hydrocarbons from Australian canebeetles (Coleoptera: Scarabaeidae) Mary T Fletcher,1 Peter G Allsopp,2* Matthew J McGrath,1 Sharon Chow,1 Oliver P Gallagher,1 Craig Hull,3 Bronwen W Cribb,3 Christopher J Moore4 and William Kitching1 1
School of Molecular and Microbial Sciences, The University of Queensland, Qld. 4072, Australia. BSES Limited, PO Box 86, Indooroopilly, Qld. 4068, Australia. 3 School of Integrative Biology, The University of Queensland, Qld. 4072, Australia. 4 Department of Primary Industries and Fisheries, Yeerongpilly, Qld. 4109, Australia. 2
Abstract
Cuticular hydrocarbon components in beetles of six Australian melolonthines whose larvae damage sugarcane, Antitrogus parvulus (Britton), A. consanguineus (Blackburn), Lepidiota negatoria (Blackburn), L. picticollis (Lea), L. noxia (Britton) and Dermolepida alborhirtum (Arrow), are identified and compared. These species demonstrate species-specific cuticular hydrocarbon profiles with a number of unprecedented structures. Major components have been identified as polymethylated hydrocarbons, 3-methyl substituted n-alkanes, 9,10-allenes and the corresponding C9 alkenes. The similarity of these compounds shows some correlation with the phylogeny of the beetles, but two polymethylated C22 hydrocarbons are unique to A. parvulus. One C25 allene is shown to have a potential role in mate recognition in A. consanguineus.
Key words
allene, Antitrogus, cuticular hydrocarbons, Dermolepida, Lepidiota, pheromone, scarab.
I NTRODUCT IO N Canegrubs are the major pests affecting the production of sugarcane in Australia (Robertson et al. 1995; McLeod et al. 1999). Nineteen species of scarab canegrubs are endemic to Australia; they belong to four genera within the tribe Melolonthini: Antitrogus Burmeister (4 species), Demolepida Arrow (1 species), Lepidiota Kirby (13 species) and Rhopaea Erichson (1 species). Although commonly lumped together in the category of ‘canegrub’, they exhibit diverse life cycles, distributions and adult behaviours (Allsopp et al. 1993; Allsopp & Lambkin 2006). Adult beetles of some short-lived species (Antitrogus, Rhopaea and some Lepidiota) emerge from the soil primarily to mate and return to oviposit within the space of a few hours, while adults of Dermolepida and several species of Lepidiota live for several weeks and congregate away from canefields in feeding trees (Allsopp et al. 1993). Through the 1990s, inadequate control associated with the withdrawal of organochlorine insecticides and degradation of insecticides, as well as a threat of insecticidal resistance, stimulated interest in alternative approaches to management (Robertson et al. 1995). Such approaches have focused on widening and integrating the control strategies available to the sugarcane industry through efficiencies, substitutions and changes to farming practices. They have been underpinned by a better understanding of the pests’ biology and ecology, and
*
[email protected] © 2008 The Authors Journal compilation © 2008 Australian Entomological Society
being adopted rapidly through targeted extension (Allsopp 2001). Sex pheromones have considerable potential for the monitoring and control of other herbivorous scarab beetles (Leal 1995, 1998), and Allsopp (1993) proposed that pheromones may be useful for population control or monitoring within the Australian canegrub complex. Bioassays and field evidence indicate that males of several of the short-lived species, Rhopaea magnicornis (Blackburn), Antitrogus parvulus (Britton), A. consanguineus (Blackburn) and Lepidiota picticollis (Lea) (Soo Hoo & Roberts 1965; Allsopp 1993), are attracted to conspecific virgin females. Allsopp and Stickley (1991), Fletcher et al. (1999, 2001, 2003), McGrath et al. (2003), Chow et al. (2005), Herber and Breit (2005), Zhou et al. (2007) and Zhu et al. (2008) have invested considerable effort in elucidating the chemical compounds responsible for this attraction. Analysis of collected beetle volatiles and extracts of purported pheromonal glands using gas chromatography-mass spectroscopy (GC-MS) have revealed only minor levels of volatile compounds (Fletcher et al. 1999). Laboratory and field bioassays have been conducted with a number of these volatile components, and, although some promising electro-antennogram results were obtained (C Hull unpubl. data 2001), PG Allsopp (unpubl. data 2001) found no field attractancy. Although lacking in volatile chemical components, these canebeetle extracts did demonstrate a considerable diversity of unusual cuticular hydrocarbons. A number of unprecedented structures have now been identified from these species, including long-chain allenes (Fletcher et al. 2001; McGrath et al. doi:10.1111/j.1440-6055.2008.00643.x
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2003) and polymethylated hydrocarbons (Fletcher et al. 2003; Chow et al. 2005). We report here for the first time the full array of hydrocarbons present in these canebeetle species, and discuss the chemotaxonomic implications. Insect cuticular hydrocarbons are generally complex mixtures of normal and branched saturated and unsaturated hydrocarbons (Howard et al. 1978) that have a primary role in the protection against desiccation, micro-organisms and abrasion (Lange et al. 1989; Singer 1998). The usefulness of cuticular hydrocarbons in arthropod taxonomy was established by Lockey (1988, 1991), and there is considerable evidence that these characters are species-specific and that the variation in these compounds accompanies speciation. They have been used to differentiate closely related species in a wide range of groups, including beetles (Lockey & Metcalfe 1988; Golden et al. 1992; Page et al. 1997; Ramamurthy et al. 1998), ants (Steiner et al. 2002), flies (Sutton & Carlson 1997), wasps (Nakabou & Ohno 2001), mole crickets (Broza et al. 2000) and termites (Page et al. 2002). Our hypothesis was that the more closely related that canebeetle species are taxonomically, the more similar should be their hydrocarbon profiles. Allsopp and Lambkin (2006) developed a phylogeny of canebeetle species. Cuticular hydrocarbons also function in the recognition systems of both social and solitary insects, and facilitate species, sex and colony recognition (Singer 1998). In social insects, cuticular hydrocarbons are known to act as sex pheromones and allow conspecific mate recognition (Lange et al. 1989). The canebeetle cuticular hydrocarbons reported here represent complex mixtures, and we have undertaken preliminary investigations into their role in mate recognition.
M ETHODS AND MAT E RIALS Insect material We collected larvae of Antitrogus parvulus, A. consanguineus, Lepidiota negatoria (Blackburn), L. noxia (Britton) and L. picticollis from canefields in the Bundaberg-Childers district of southern Queensland (25°S, 152.20°E) and larvae of Dermolepida albohirtum (Arrow) from fields in the Burdekin area of northern Queensland (19.40°S, 147.25°E). We chose two species of Antitrogus that show evidence of sex-attraction in these species (Allsopp 1993), two major lineages of Lepidiota (Allsopp & Lambkin 2006), one of which shows sex-attraction (Allsopp 1993), and the most-important canegrub species D. albohirtum. We reared them to adults in their native soil in individual containers at 25°C using a diet of germinated grass seeds. All adults were killed by freezing – none could have mated and could have come in contact only with soil material. Adults were identified using the keys of Miller and Allsopp (2000).
Analytical procedures We used GC-MS technology to identify the cuticular hydrocarbons. In this technique, the gas chromatograph utilises a © 2008 The Authors Journal compilation © 2008 Australian Entomological Society
capillary column and, depending on the column’s dimensions (length, diameter, film thickness) as well as the phase properties, the difference in the chemical properties among different molecules in a mixture will separate the molecules as the sample travels the length of the column. The molecules take different times (retention time) to come out of the gas chromatograph, and this allows the mass spectrometer downstream to capture, ionise and detect the molecules separately. The mass spectrometer does this by breaking each molecule into ionised fragments and detecting these fragments using their mass-to-charge ratio. We used a Shimadzu QP5050 instrument with a DB5-MS capillary column. Hydrocarbon analyses were conducted using a temperature program from 190°C (0 min) increasing to 300°C at a rate of 5°C/min.
Retention indices We calculated retention indices or Kovát’s indices (KIs) from interpolation between the retention times of a standard solution containing n-alkanes from C23 to C30. The standard solution was analysed before acquisition of data on each beetle extract. Co-injections of beetle extract and standard n-alkanes were also performed. The KI expresses the number of carbon atoms (¥100) of a hypothetical normal alkane that would have an adjusted retention volume (time) identical to that of the peak of interest when analysed under identical conditions.
Extracts Abdominal tips from female beetles were excised by scalpel and extracted into dichloromethane. Number of beetles extracted was dependent on availability (typical sample size was 20 beetles).
Statistical analysis We used ordinal multidimensional scaling (MDS) to visualise data on the presence and proportions in each species of each of the 21 named compounds and the two unidentified polymethyl-hydrocarbons in Table 1. MDS (Young & Hamer 1994) places each species at a particular locus in an n-dimensional space, so that the interpoint distances correspond to the dissimilarity of those points. Thus, two species that have very similar hydrocarbon compositions will be placed close together, whereas dissimilar ones will be widely spaced. Theoretically, there is no limit to the number of dimensions that can be used to define the locations of points. However, a solution in two dimensions is easiest to portray, but the placement may be distorted. The amount of distortion imposed can be assessed using Kruskal’s Stress Formula One (Spence 1978). We calculated Spearman rank correlations of the percentage composition of the hydrocarbons of each species compared with those of every other species. This matrix (after scaling so that identical profiles (rank correlation of 1) equalled 0) was used as the input for MDS. We used Unistat 5.0 (Unistat 2000),
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Table 1 Proportion (%) of cuticular hydrocarbons from abdominal tips of female canebeetles (compounds are referred to in the text and in Figure 1 by the bolded numbers that reflect increasing retention indices) Compound Tricosane (4) 3-methyltricosane (5) 9-tricosene (1) 9,10-tricosadiene (3) Pentacosane (10) 3-methylpentacosane (11) 9-pentacosene (8) 9,10-pentacosadiene (9) Heptacosane (14) 3-methylheptacosane (16) 9-heptacosene (12) 9,10-heptacosadiene (13) 13-methylheptacosane (15) Nonacosane (18) 3-methylnonacosane (21) 9-nonacosene (17) 13-, 15-methylnonacosane (19) and (20) 4,6,8,10,16-penta-methyldocosane (6) 4,6,8,10,16,18-hexa-methyldocosane (7) Unidentified polymethyl-hydrocarbon Unidentified polymethyl-hydrocarbon 1-cosanol (2) Other hydrocarbons
Retention index (KI) 2300 2371 2275 2297 2500 2571 2472 2494 2700 2774 2672 2693 2732 2900 2972 2876 2929 2396 2424 2880 2952 2288
Antitrogus parvulus
Antitrogus consanguineus
13 12 14 50 Trace
Lepidiota negatoria
Lepidiota picticollis
10 19
9 27 16 30
6
Lepidiota noxia
Dermolepida albohirtum
11
12 11 23 3 4
27 16
4 6
2
4 3 2 Trace 14 10 10 6 5 8 6 5
45 38 10 8 17
1
14 8
18
16†
4 23†
†L. noxia and D. albohirtum extracts contained a number of unidentified hydrocarbons with retention indices between 2700 and 3000.
and made comparisons of the pattern with the phylogeny developed by Allsopp and Lambkin (2006).
Behavioural bioassays To determine if at least one of these hydrocarbons could be involved in sex attraction, we coated small polypropylene microcentrifuge tubes (1.5 mL) with a solution of synthetic (R)-9,10-pentacosadiene (C25 (R)-allene) (9) (McGrath et al. 2003) in hexane (ca. 200 mL of a 3.5 mg/mL solution) and coated duplicate tubes with similar quantities of hexane (as a control). Hexane was allowed to evaporate from each tube before it was presented to A. consanguineus beetles during the normal active time period (19:00–22:00 h). Nine virgin laboratory-reared adult male A. consanguineus were placed individually in contact with both a control tube and then a C25 (R)-allene coated tube. Non-responsive beetles were retested later.
RE SULTS AND D ISCU SSIO N Identification of cuticular components The GC-MS analysis of abdominal tip extracts from females of the canebeetles Antitrogus parvulus, A. consanguineus, Lepidiota negatoria, L. noxia, L. picticollis and Dermolepida albohirtum revealed a complex of components (Fig. 1) but with substantial differences in presence and amount among the six
species (Table 1). Compounds are identified by the numbers (bolded and in parentheses in the following text) that reflect increasing retention indices given in Table 1 and Figure 1. The polymethylated hydrocarbons 4,6,8,10,16-pentamethyldocosane (6) and 4,6,8,10,16,18-hexa-methyldocosane (7), and the allenes 9,10-tricosadiene (3), 9,10-pentacosadiene (9) and 9,10-heptacosadiene (13) were identified by comparison with synthetic samples (Fletcher et al. 2001, 2003; McGrath et al. 2003). The stereochemistry of the 4,6,8,10,16penta- and 4,6,8,10,16,18-hexamethyldocosanes (6) and (7) was established by comparisons of mass spectra, highresolution nuclear magnetic resonance spectroscopy and chromatographic properties of various synthetic samples and natural hydrocarbons – both have an anti-anti-anti stereochemistry of the methyl tetrad moiety, and a syn methyl diad at the other end of the hexamethyldocosane (Chow et al. 2005; Herber & Breit 2005; Zhou et al. 2007; Zhu et al. 2008). These five compounds represent unprecedented structures in nature. The extract from L. noxia contained two unidentified components (KI 2880 and 2952). These appear, from mass spectral analysis, to be novel polymethyl hydrocarbons with the same alternate methyl substitution pattern as present in compounds (6) and (7) (Fletcher et al. 2003), albeit with a longer carbon chain. Z-9-Tricosene (1), 1-cosanol (2) and the n-alkanes (4), (10), (14) and (18) were identified by comparison of mass spectra and retention indices with commercially available samples. Methoxymercuration was used to confirm the position of the double bond in 9-pentacosene (8) and 9-heptacosene (12) © 2008 The Authors Journal compilation © 2008 Australian Entomological Society
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C8H17
nC H 8 17
R
(1) R = nC13H27 (8) R = nC15H31 (12) R = nC17H35 (17) R = nC19H39
R Me (5) R = nC19H39 (11) R = nC21H43 (16) R = nC23H47 (21) R = nC25H51
R
(3) R = nC12H25 (9) R = nC14H29 (13) R = nC16H33
nC H OH 20 41
nC H 23 48
(2)
nC H 25 52
(4)
Me
Me
Me
nC H 27 56
(10)
Me
(14)
Me
nC H 29 60
(18)
R
(6) R = H (7) R = Me
Me
R1
(15)
R2
(19) R1 = Me, R2 = H (20) R1 = H, R2 = Me
(Fletcher et al. 2001; McGrath et al. 2003), together with published retention indices for 9-pentacosene (8) (Bagnères et al. 1990; Hedlund et al. 1996). The mass spectra of 9-heptacosene (12) and 9-nonacosene (17) were similar to their homologues and retention indices agreed with published values (Subchev & Jurenka 2001). Literature retention indices for 3-methyltricosane (5), 3-methylpentacosane (11), 3-methylheptacosane (16) and 3-methylnonacosane (21) (Nikolova et al. 1999), together with mass spectral comparisons with reported mass spectra for 3-methylpentacosane (11) (Howard et al. 1978) and mass spectral data for (11), (16) and (21) (Marukawa et al. 2001) confirmed the identity of these compounds. Mass spectra for these compounds all showed prominent ions at M–C2H5 indicative of methyl branching at C-3 (Howard et al. 1978). Similar mass spectral interpretation enabled structures to be deduced for 13-methylheptacosane (15) and 13- and 15-methylnonacosane (19) and (20) (coeluting as one peak), which agreed with literature retention indices for these compounds (Bernier et al. 1997; Nikolova et al. 1999).
Cuticular composition and taxonomic implications The hydrocarbon compositions (Table 1) are unusual in that in four species (A. parvulus, A. consanguineus, L. picticollis and L. negatoria), a small number of components (2–4) account for 80–90% of the identified cuticular hydrocarbons. This is in © 2008 The Authors Journal compilation © 2008 Australian Entomological Society
Fig. 1. Cuticular hydrocarbons identified from Australian melolonthine canebeetles. Numbers refer to compounds identified in Table 1 and the text and reflect increasing retention indices.
contrast to the complex mixtures of hydrocarbons generally found in insect cuticular hydrocarbons (e.g. Howard et al. 1978; Lockey & Metcalfe 1988; Steiner et al. 2002), of which the multitude of components found in D. albohirtum and to a lesser extent L. noxia (Table 1) is more typical. However, the differences in type and proportion of the compounds easily allow differentiation of the five species (Fig. 2) and have the potential to be used for identification and/or in developing phylogenies. However, comparisons with other species of canebeetles would be required to improve our knowledge of the occurrence of the compounds. The predominant major compounds in the cuticles of the six canebeetles are n-alkanes, 3-methyl substituted n-alkanes, 9,10-allenes and the corresponding C9 alkenes. They range from C23-chain compounds in L. picticollis, through C23-chain and C25-chain compounds in L. negatoria, to mainly C25-chain but with some C23-chain and C27-chain compounds in L. noxia, to C25-chain and C27-chain compounds in A. consanguineus. C29-chain compounds occur in D. albohirtum, along with C27chain and some C25-chain and C23-chain compounds. The occurrence of the two novel polymethylated C22-chain alkanes in A. parvulus is unique and contributes to that species’ isolated placement on the MDS plot. The similarity of the hydrocarbon profiles shows good correlation with our understanding of the phylogeny (Allsopp & Lambkin 2006). The three genera are well separated in the MDS plot. Of the three Lepidiota spp., negatoria and noxia are more closely related to each other than either is to picticollis.
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0.5 0.4
pic
0.3
Dimension 2
0.2
nox
neg
0.1 0.0
–0.6
–0.4
–0.2
0.0
0.2
0.4
0.6
–0.1 par con
alb
–0.2 –0.3 –0.4
Stress < 0.0001
Dimension 1
Fig. 2. Multidimensional scaling plot based on the presence and proportion of compounds (1)–(21) and the two unidentified polymethyl-hydrocarbons in Table 1. Species are coded: alb, Dermolepida albohirtum; con, Antitrogus consanguineus; neg, Lepidiota negatoria; nox, Lepidiota noxia; par, Antitrogus parvulus; pic, Lepidiota picticollis.
The dissimilarity of A. consanguineus and A. parvulus, in both chain length and the extent of methylation in the two parvulus hydrocarbons, is surprising given their reasonably close taxonomic relationship.
Biological role The novelties of the allenic and polymethyl structures and their distribution among the species suggest an important biological role. Five of these canebeetle species occur in the same geographical area, so require different mate-recognition systems to prevent incorrect matings. These hydrocarbons may play such a role in conspecific mate recognition. Male Antitrogus consanguineus beetles appear to virtually ‘pat’ their mate with fully extended and splayed antennal clubs during mating (PG Allsopp unpubl. data 1991). Thus, as a preliminary bioassay, we placed virgin A. consanguineus males in contact with microcentrifuge tubes coated with synthetic C25 (R)-allene (9) (the major cuticular component of female A. consanguineus) in hexane. Of the nine male beetles tested, six beetles gripped the allene-coated tube with splayed antennae and probed with their aedeagus (Fig. 3) in an apparent attempt to mate with the tube. Beetles did not respond to similar tubes treated with hexane alone. We conclude that the C25 allene (9) must be a significant stimulus in mate recognition and acts as some form of contact sexual pheromone in
Fig. 3. Antitrogus consanguineus male beetle with antennae splayed and aedeagus extended attempting to mate with a microcentrifuge tube coated with synthetic C25 R-allene (9).
this species. Major cuticular components identified in the other canebeetle species (Table 1) may well perform a similar contact pheromonal function. However, allene (9) was tested in the field and does not appear to draw in A. consanguineus beetles over any distance (PG Allsopp unpubl. data 2001). The long chain length of the allene means that it is of very low volatility. Mate attraction © 2008 The Authors Journal compilation © 2008 Australian Entomological Society
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and recognition can often involve a series of chemical cues, for example, males of the closely related Melolontha hippocastani (Fabricius) are attracted by feeding-induced plant volatiles and a female-driven sex pheromone (Ruther et al. 2000). Similar situations, but with the addition of a contact pheromone, may exist with the species that we studied. It would explain why we sometimes see male–male copulation among at least A. consanguineus (Allsopp & Morgan 1991).
ACKNOWLE D G E ME NT This research was supported by an Australian Research Council grant (SPIRT Award 1999–2001).
R EFE RE NCE S Allsopp PG. 1993. Evidence for sex attraction in three species of Australian canegrub beetles (Coleoptera: Scarabaeidae: Melolonthinae). Coleopterists’ Bulletin 47, 51–52. Allsopp PG. 2001. The research process – canegrub management in the Australian industry. Proceedings of the Australian Society of Sugar Cane Technologists 23, 9–16. Allsopp PG & Lambkin CL. 2006. Canegrubs and cladistics: what story do adult, larval and ecological characters tell? Australian Journal of Entomology 45, 55–66. Allsopp PG & Morgan TA. 1991. Male–male copulation in Antitrogus consanguineus (Blackburn) (Coleoptera: Scarabaeidae). Australian Entomological Magazine 18, 147–148. Allsopp PG & Stickley BDA. 1991. Assessment of the Potential of Sex Pheromones as Strategic Lures for the Control of Canegrubs. Final Report SRDC Project BS17S. BSES, Brisbane, Australia. Allsopp PG, Chandler KJ, Samson PR & Story PG. 1993. Pests of Australian Sugarcane. Bureau of Sugar Experiment Stations, Brisbane, Australia. Bagnères AG, Clément JL, Blum MS, Severson RF, Joulie C & Lange C. 1990. Cuticular hydrocarbons and defensive compounds of Reticulitermes flavipes (Kollar) and R. santonensis (Feytaud): polymorphism and chemotaxonomy. Journal of Chemical Ecology 16, 3213–3244. Bernier UR, Carlson DA & Geden CJ. 1997. Gas chromatography/mass spectrometry analysis of the cuticular hydrocarbons from parasitic wasps of the genus Muscidifurax. Journal of the American Society for Mass Spectrometry 9, 320–332. Broza M, Nation JL, Milne K & Harrison J. 2000. Cuticular hydrocarbons as a tool supporting recognition of Gryllotalpa tali and G. marismortui (Orthoptera: Gryllotalpidae) as distinct species in Israel. Annals of the Entomological Society of America 93, 1022–1030. Chow S, Fletcher MT, Lambert LK et al. 2005. Novel cuticular hydrocarbons from the cane beetle Antitrogus parvulus – 4,6,8,10,16-pentaand 4,6,8,10,16,18-hexamethyldocosanes – unprecedented anti-antianti-stereochemistry in the 4,6,8,10-methyltetrad. Journal of Organic Chemistry 70, 1808–1827. Fletcher MT, Hayes P, Kitching W, Cribb BW, Allsopp PG & Moore CJ. 1999. Australian Sugarcane Beetle Pheromones. First Asia-Pacific Conference on Chemical Ecology, Shanghai, China. Fletcher MT, McGrath MJ, Kitching W et al. 2001. A novel group of allenic hydrocarbons from five Australian (Melolonthine) beetles. Chemical Communications 2001, 885–886. Fletcher MT, Chow S, Lambert LK et al. 2003. 4,6,8,10,16-penta- and 4,6,8,10,16,18-hexamethyl-docosanes from the cane beetle, Antitrogus parvulus – cuticular hydrocarbons with unprecedented structure and stereochemistry. Organic Letters 5, 5083–5086. Golden KL, Meinke LJ & Stanley-Samuelson DW. 1992. Cuticular hydrocarbon discrimination of Diabrotica (Coleoptera: Chrysomelidae) sibling species. Annals of the Entomological Society of America 85, 561–570. © 2008 The Authors Journal compilation © 2008 Australian Entomological Society
Hedlund K, Bartelt RJ, Dicke MC & Vet LEM. 1996. Aggregation pheromones of Drosophila immigrans, D. phalerata, and D. subobscura. Journal of Chemical Ecology 22, 1835–1844. Herber C & Breit B. 2005. Enantioselective total synthesis and determination of the absolute configuration of the 4,6,8,10,16,18hexamethyldocosane from Antitrogus parvulus. Angewandte Chemie International Edition 44, 5267–5269. Howard RW, McDaniel CA & Blomquist GJ. 1978. Cuticular hydrocarbons of the eastern subterranean termite, Reticulitermes flavipes (Kollar) (Isoptera: Rhinotermitidae). Journal of Chemical Ecology 4, 233–245. Lange C, Basselier JJ, Bagneres AG et al. 1989. Strategy for the analysis of cuticular hydrocarbon waxes from insects using gas chromatography/mass spectrometry with electron impact and chemical ionization. Biomedical and Environmental Mass Spectrometry 18, 787–800. Leal WS. 1995. Sex pheromones of plant-feeding scarab beetles. Korean Journal of Entomology 34, 9–14. Leal WS. 1998. Chemical ecology of phytophagous beetles. Annual Review of Entomology 43, 39–61. Lockey KH. 1988. Lipids of the insect cuticle: origin, composition and function. Comparative Biochemistry and Physiology, B Comparative Biochemistry 89, 595–645. Lockey KH. 1991. Insect hydrocarbon classes: implications for chemotaxonomy. Insect Biochemistry 121, 91–97. Lockey KH & Metcalfe NB. 1988. Cuticular hydrocarbons of adult Himatismus species and a comparison with 21 other species of adult tenebrionid beetle using multivariate analysis. Comparative Biochemistry and Physiology, B Comparative Biochemistry 91, 371–382. McGrath MJ, Fletcher MT, König WA et al. 2003. A suite of novel allenes from Australian melolonthine scarab beetles. Structure, synthesis, and stereochemistry. Journal of Organic Chemistry 68, 3739–3748. McLeod RS, McMahon GG & Allsopp PG. 1999. Costs of major pests and diseases to the Australian sugar industry. Plant Protection Quarterly 14, 42–46. Marukawa K, Takikawa H & Mori K. 2001. Synthesis of the enantiomers of some methyl-branched cuticular hydrocarbons of the ant, Diacamma sp. Bioscience, Biotechnology and Biochemistry 65, 305–314. Miller LJ & Allsopp PG. 2000. Identification of Australian canegrubs (Coleoptera: Scarabaeidae: Melolonthini). Invertebrate Taxonomy 14, 377–409. Nakabou M & Ohno T. 2001. Identification of Orius spp. by cuticular hydrocarbons. Japanese Journal of Applied Entomology and Zoology 45, 18–21. Nikolova N, Rezanka T, Nikolova-Damyanova B & Kalushkov P. 1999. Hydrocarbons in adult Chrysomela vigintipunctata (Scopoli) (Coleoptera: Chyrsomelidae). Comparative Biochemistry and Physiology, B Comparative Biochemistry 123, 67–77. Page M, Nelson LJ, Blomquist GJ & Seybold SJ. 1997. Cuticular hydrocarbons as chemotaxonomic characters of pine engraver beetles (Ips spp.) in the grandicollis subgeneric group. Journal of Chemical Ecology 23, 1053–1099. Page M, Nelson LJ, Forschler BT & Haverty MI. 2002. Cuticular hydrocarbons suggest three lineages in Reticulitermes (Isoptera: Rhinotermitidae) from North America. Comparative Biochemistry and Physiology, B Comparative Biochemistry and Molecular Biology 131, 305–324. Ramamurthy VV, Poorani J, Devakumar C & Dureja P. 1998. Cuticular hydrocarbons and biosystematics of emtimine weevil genera Lepropus Schoenherr and Brachyaspistes Fahraeus (Curculionidae: Coleoptera). Entomon 23, 251–257. Robertson LN, Allsopp PG, Chandler KJ & Mullins RT. 1995. Integrated management of canegrubs in Australia: current situation and future research directions. Australian Journal of Agricultural Research 46, 1–16. Ruther J, Reinecke A, Thiemann K, Tolasch T, Francke W & Hilker M. 2000. Mate finding in the forest cockchafer, Melolontha hippocastani, mediated by volatiles from plants and females. Physiological Entomology 25, 172–179. Singer TL. 1998. Roles of hydrocarbons in the recognition systems of insects. American Zoologist 38, 394–405.
Diverse cuticular hydrocarbons Soo Hoo CF & Roberts RJ. 1965. Sex attraction in Rhopaea. Nature 205, 724–725. Spence I. 1978. Multi-dimensional scaling. In: Quantitative Ethology (ed. PW Colgan), pp. 175–217. Wiley, New York, USA. Steiner FM, Schlick-Steiner BC, Nikiforov A, Kalb R & Mistrik R. 2002. Cuticular hydrocarbons of Tetramorium ants from central Europe: analysis of GC-MS data with self-organising maps (SOM) and implications for systematics. Journal of Chemical Ecology 28, 2569–2584. Subchev M & Jurenka RA. 2001. Sex pheromone levels in pheromone glands and identification of the pheromone and hydrocarbons in the hemolymph of the moth Scoliopteryz libatrix L. (Lepidoptera: Noctuidae). Archives of Insect Biochemistry and Physiology 47, 35–43. Sutton BD & Carlson DA. 1997. Cuticular hydrocarbon variation in the Tabanidae (Diptera): Tabanus nigrovittatus complex of the North
159
American Atlantic coast. Annals of the Entomological Society of America 90, 542–549. Unistat Limited. 2000. Unistat 5.0 Statistical Package for Windows™. User’s Guide. Unistat, London, UK. Young FW & Hamer RM. 1994. Theory and Applications of Multidimensional Scaling. Eribaum Associates, Hillsdale, New Jersey, USA. Zhou J, Zhu Y & Burgess K. 2007. Synthesis of (S,R,R,S,R,S)4,6,8,10,16,18-hexamethyldocosane from Antitrogus parvulus via diastereoselective hydrogenations. Organic Letters 9, 1391–1393. Zhu G, Liang B & Negishi E. 2008. Efficient and selective synthesis of (S,R,R,S,R,S)-4,6,8,10,16,18-hexamethyldocosane via Zr-catalyzed asymmetric carboalumination of alkenes (ZACA reaction). Organic Letters 10, 1099–1101. Accepted for publication 14 October 2007.
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