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Cuticular Chemistry of the Queensland Fruit Fly Bactrocera tryoni (Froggatt)

Applied BioSciences, Macquarie University, North Ryde, NSW 2109, Australia
Australian Research Council Centre for Fruit Fly Biosecurity Innovation, Macquarie University, North Ryde, NSW 2109, Australia
Commonwealth Scientific and Industrial Research Organisation Land and Water, Black Mountain, Acton, ACT 2601, Australia
Author to whom correspondence should be addressed.
Molecules 2020, 25(18), 4185;
Received: 8 August 2020 / Revised: 26 August 2020 / Accepted: 10 September 2020 / Published: 12 September 2020
(This article belongs to the Section Natural Products Chemistry)


The cuticular layer of the insect exoskeleton contains diverse compounds that serve important biological functions, including the maintenance of homeostasis by protecting against water loss, protection from injury, pathogens and insecticides, and communication. Bactrocera tryoni (Froggatt) is the most destructive pest of fruit production in Australia, yet there are no published accounts of this species’ cuticular chemistry. We here provide a comprehensive description of B. tryoni cuticular chemistry. We used gas chromatography-mass spectrometry to identify and characterize compounds in hexane extracts of B. tryoni adults reared from larvae in naturally infested fruits. The compounds found included spiroacetals, aliphatic amides, saturated/unsaturated and methyl branched C12 to C20 chain esters and C29 to C33 normal and methyl-branched alkanes. The spiroacetals and esters were found to be specific to mature females, while the amides were found in both sexes. Normal and methyl-branched alkanes were qualitatively the same in all age and sex groups but some of the alkanes differed in amounts (as estimated from internal standard-normalized peak areas) between mature males and females, as well as between mature and immature flies. This study provides essential foundations for studies investigating the functions of cuticular chemistry in this economically important species.
Keywords: cuticular hydrocarbons; cuticle; chemical communication; GC-MS; methyl branched alkanes; chemical ecology; volatiles cuticular hydrocarbons; cuticle; chemical communication; GC-MS; methyl branched alkanes; chemical ecology; volatiles

1. Introduction

The cuticular layer of the insect exoskeleton contains a range of mostly aliphatic compounds, including normal and branched alkanes, alkenes, saturated and unsaturated esters, alcohols, saturated and unsaturated fatty acids, ketones, and aldehydes [1]. Cuticular hydrocarbons usually contain 20 to 50 carbons, and compounds with other functional groups vary from 12 to 54 carbons [2,3,4]. A primary function of cuticular hydrocarbons is to protect against desiccation [5,6], injury, and infection [7,8,9,10,11,12]. Cuticular compounds, including hydrocarbons, are also commonly important for chemical communication [2,5,6,13], including species recognition [14,15,16,17], mimicry [18], and as pheromones [19] in diverse insect taxa. For example, some cuticular hydrocarbons serve as sex pheromones in house fly [20], the circumboreal fly [21], moths [22], bees [23] and the cowpea weevil [24]. Cuticular hydrocarbons also serve as aggregation pheromones in some insects, including Drosophila [25], termites [26] and cockroaches [27]. Sexual selection has been a driving force for the evolution of sexual dimorphism in animals [28], and many insect taxa exhibit sexual dimorphism in cuticular chemistry [29]. For example, sexually dimorphic cuticular hydrocarbons have been found in Drosophila [30,31,32] and have been implicated in female attractiveness and male mating success [21].
Tephritid fruit flies are amongst the world’s most economically damaging insect pests [33]. Some aspects of tephritid fruit fly semiochemistry have received significant attention, particularly the pheromones [34,35] they use to attract mates and for aggregation and the particular compounds found in fruit, food and certain flowers to which they are attracted [36,37,38,39,40,41,42,43,44,45,46,47]. Some work has also been performed on fruit fly cuticular chemistry because their cuticular chemical profiles tend to be highly species-specific [48,49] and have been used to resolve species, cryptic species and geographic variation in larvae [50,51,52] and adults [53,54,55,56,57,58]. Beyond their use as chemotaxonomic tools, however, relatively little work has been performed on tephritid cuticular compounds. In an important recent exception, allyl-2,6-dimethoxyphenol has been proposed as a short-range male attractant in Bactrocera dorsalis [59]. Most cuticular compounds are aliphatic, so this case is also notable for its involvement of an aromatic compound.
In Australia and in some Pacific Islands, the Queensland fruit fly, Bactrocera tryoni (Froggatt), is an economically important pest of horticultural crops [60,61,62]. This species causes significant economic loss by damaging crops [63] and by limiting market access [64]. While rectal gland and volatile emission chemistry of B. tryoni has been documented [65,66,67,68,69,70], the cuticular chemistry of this species has not. Given what is known for other insects, the composition of its cuticular chemical profile is likely to be relevant to homeostasis, protection from pathogens, injury and insecticides [71], and chemical communication. Understanding elements of cuticular chemistry related to homeostasis may help to understand abiotic factors mediating bioclimatic potential of B. tryoni [72] and effects of domestication, sex and age on desiccation resistance [73,74], and may also be important for understanding environmental and sexual competence of sterile B. tryoni released in sterile insect technique (SIT) programs to control pest populations [75,76,77]. To address this knowledge gap, and to provide foundations for subsequent functional studies, the present study reports a qualitative description of B. tryoni cuticular chemistry and identifies qualitative and quantitative variation (the latter estimated from internal standard-normalized peak areas) related to maturity and sex. Bactrocera tryoni specimens were obtained as larvae in infested fruits and cuticle extracts of emerged adults were analyzed by gas chromatography-mass spectrometry (GC-MS).

2. Results

Cuticular Chemistry and Statistical Analysis

The identified compounds are all aliphatic; no trace of aromatic compounds was found. Typical chromatograms of both immature and mature female and male B. tryoni are shown in Figure 1. The chromatogram sections of shorter (A) and longer retention time compounds (B) of B. tryoni adults are shown in Figure 2, where a typical chromatogram of a female is presented because the chromatogram includes all the compounds that are also found in immature and mature females and males. A range of non-alkanes, including spiroacetals, amides and esters and an assortment of C29 to C33 methyl-branched alkanes represent the cuticular chemistry of wild B. tryoni adults.
The identities of 22 of the 32 non-alkane compounds found were confirmed with authentic standards and the other ten were tentatively identified by comparison to fragment patterns in NIST libraries (Table 1). Two of the compounds were 6,6-membered ring spiroacetals (compounds A2 and A5 in Table 1 and Figure 2), four were aliphatic amides (compounds A1, A3, A4 and A6 in Table 1 and Figure 2), and the remaining 26 were all esters of saturated/unsaturated and methyl branched saturated fatty acids (compounds A7A32 in Table 1 and Figure 2). The spiroacetals and saturated/unsaturated and branched saturated esters were found to be specific to mature females, while the amides were found in mature flies of both sexes (Table 1). In mature females, ethyl esters of saturated or unsaturated C12, C14, C16 and C18 are the most abundant, while methyl and propyl esters, and branched saturated fatty acid esters are minor and trace, respectively. Methyl positions in the branched fatty acid esters are ambiguous, because trace amounts of the compounds made it difficult for further analyses. The amount of N-(3-methylbutyl)isobutyramide (A6) was about 2.5 times larger in mature male than in mature females (p < 0.05, t-test), but the amount of N-(3-methylbutyl)propanamide (A4) was about 3.2 times larger in mature females than in mature males (p < 0.05, t-test). The differences in the amounts of the other amides, N-(3-methylbutyl)acetamide (A1) and N-(2-methylbutyl)propanamide (A3) were not significant (p > 0.05 for all, t-test). The total amount of the amides was 2.6 times larger in mature females than in mature males (p < 0.05, t-test). The results are illustrated in Figure 3.
The 34 tentatively identified hydrocarbons, all methyl-branched alkanes with C29 to C33 carbon backbones, are summarized in Table 2. Unsaturated hydrocarbons were not detected. Most are mono- or dimethylalkanes, with only a few trimethylalkanes found. Monomethyl branches appeared exclusively at odd carbon positions in odd carbon alkanes and at even carbon positions in even carbon alkanes. Comparisons in the amounts of individual hydrocarbons between sexes are illustrated in Figure 4. The most abundant alkanes were mono- and dimethylhentriacontane (C31) isomers (compounds B18 to B24 in Figure 2B and Figure 4), with the 11-, 13- and 15-methylhentriacontanes (B18) appearing at the highest intensity in chromatograms. Although dimethyl branches were separated by 1, 3, 5, 7, 9, 11, or 13 methylene groups, a majority of dimethyl branches were separated by 3 and 5 methylenes. Trimethyl branches were separated by [3,3], [3,5] or [3,11] methylenes.
There were significant effects of sexual maturity, sex and the interaction between these two variables on the amounts of the normal and methyl branched alkanes (sexual maturity F1,2276 = 81.90, p < 0.0001; F1,2276 = 10.56, p < 0.005; sexual maturity * sex F1,2276 = 4.10, p < 0.05, respectively). The amounts of alkanes in mature flies were 2.3 (±1.3) times larger than those in immature flies on average. The amounts of compounds B1, B3, B4, B5, B6, B7, B9, B10, B11, B12, B13, B18, B19, B20, B21, B22, B22, and B24 were higher in mature females than in mature males (p < 0.05, t-test). The amounts of these compounds were 1.7 (±0.5) times greater in mature females than in mature males on average. There was no significant difference in the amount of hydrocarbons between immature females and immature males (p > 0.05, t-test).

3. Discussion

The present study finds that the n-hexane-extracted cuticular chemistry of B. tryoni includes a complex mixture of at least 66 compounds, including two spiroacetals, four aliphatic amides, 26 saturated/unsaturated C12 to C20 methyl, ethyl and propyl esters and 34 methyl branched saturated alkanes with a range of C29 to C33 carbon backbones. A previous study reported 14 cuticular compounds in B. tryoni, including five fatty acid esters, two siloxanes and seven methyl branched alkanes [98]. We did not detect the siloxanes or the methyl branched alkanes, which are all shorter than the alkanes found in the present study. We suspect that occurrence of the siloxanes in the previous work may reflect impurities, and incorrect assignments may have been given for the shorter methyl branched alkanes. The differences might have been also caused by technical differences; for example the previous work extracted cuticular compounds in methanol for 20 min [98], while the present study extracted in n-hexane for 3 min. The solvent choice of n-hexane was based on anticipated polarities of insect cuticular compounds that are generally less or non-polar [99]. The method used in the present study is more similar to methods widely used in studies of the cuticular hydrocarbons of other tephritids [53,54,55,56,57,58,59,100].
We found sexual dimorphism in several aspects of the cuticular chemistry of B. tryoni. In particular, our data suggest that spiroacetals and esters are specific to mature females. Sex-related differences in cuticular chemistry have also been reported in B. dorsalis, in which mature males have 7-monoenes that are absent from mature females and immatures of both sexes [100]. We also found quantitative differences between the sexes in their cuticular amides, which overall were more abundant in mature females than mature males, even though they are known to be particularly abundant in the rectal glands of mature males. Although the amides from male rectal glands have been suggested to function as male sex pheromones [65,66,68], the functions of these compounds on the cuticle, and in rectal glands of mature females, are not known. Our results indicate that more work is now needed on their functions in both sexes, both in rectal glands and on the cuticle.
Most of the non-alkane components which we find in cuticular extracts of B. tryoni have also been reported previously in rectal gland extracts of this species, with some of the same sex differences also evident [65,67,68,69]. In particular, the spiroacetals, amides and saturated/unsaturated esters we found in female cuticles had also been reported in female rectal glands. The presence of minute amounts of the amides and absence of the spiroacetals found on male cuticles also matches the earlier findings for male rectal glands, although the previous work also reported shorter chain esters (C5 or less) in male rectal glands that were not found on cuticles. Notwithstanding the difference in the esters, the similarities between the two extract types otherwise suggest that at least some of the non-alkane components on the cuticle could originate in the rectal glands and be distributed over the body when grooming, as has been described in some social insects [101,102]. The differences in the esters and the quantitative differences in the amides noted above might in part reflect differences in volatility of the various compounds.
The C12 to C20 esters we found in cuticle of mature female B. tryoni have also been reported in the cuticle of females in five B. dorsalis complex species, but at different relative abundances between the species, suggesting that they may have a role in species recognition [58,59]. On the other hand, the methyl branched fatty acid esters found in the present study have not been previously reported in either the emission profile or rectal gland extracts of B. tryoni [67], or other fruit flies. Methyl branched fatty acid methyl, propyl or longer alkyl esters occur in other arthropods, such as spiders [103,104], but, to the best of our knowledge, specific roles have not yet been identified for them in any species. Branched fatty acid esters are biosynthetically feasible [105,106] and fatty acids are also the precursors of other cuticular compounds and hydrocarbons [2]. Hence, the branched esters found in this study may be intermediate products in the biosynthesis of other cuticular compounds.
While most of the C29 to C33 methyl branched alkanes detected in the present study have also been reported in other organisms (see references in Table 2), several of them have not previously been described in tephritids. Branched alkanes and/or unsaturated hydrocarbons have lower melting points than the corresponding normal alkanes, allowing the waxy layer of insect cuticles to be flexible over a wide range of ambient temperatures [107,108]. Long chain hydrocarbons, and in particular branched and unsaturated alkanes, have also been linked to desiccation resistance in a variety of insects [109,110,111]. However, to what extent that applies in B. tryoni adults remains unclear. While qualitatively similar across the sexes and age groups studied, we found higher amounts of alkanes (as estimated from internal-standard-normalised peak areas) in mature than immature flies but, at least in domesticated flies from colonies maintained on artificial diets, desiccation resistance of B. tryoni decreases with age [73].
Several normal and methyl branched C28 to C40 alkanes have also been reported previously in two closely related taxa in the Bactrocera dorsalis complex, but as different isomers and in different amounts between the two [57]. Such variation in cuticular hydrocarbon chemistry has also been reported among species of another tephritid genus, Anastrepha [53,55,56,58,100], and even among populations within Ceratitis rosa [55]. As with the esters above, these taxon-specific cuticular hydrocarbon signatures may play a role in taxon recognition in nature, but they may also make excellent tools for taxonomists [53,55,56,57,58,100]. Their use in taxonomy is directly relevant to the B. tryoni complex which, despite its importance as a pest, is not well understood taxonomically. The B. tryoni complex includes three other taxa; B. neohumeralis, B. aquilonis and B. melas [60]. Bactrocera tryoni is clearly differentiated from B. neohumeralis in the timing of mating behavior [112], but the species status of B. aquilonis and B. melas is still debated [113] and the four taxa differ in their pest status and quarantine restrictions in various jurisdictions [113]. However, before cuticular chemistry can be used to resolve such taxonomic issues it will be important to investigate the extent to which it can be influenced by diet, maturity and the physical environment [114,115,116]. Given the population differences found in Ceratitis rosa [55], it will also be important to determine whether cuticular chemistry varies between geographic regions and during the course of domestication of species in the B. tryoni complex.
In summary, the present study provides the first detailed description of the cuticular chemistry of B. tryoni, and finds some clear qualitative and quantitative differences as flies mature and between the sexes. Our findings provide a foundation for studies addressing the roles of cuticular chemistry in functions such as desiccation resistance, protection from pathogens and injury, and chemical communication, as well as its potential application in resolving the taxonomy of the B. tryoni complex.

4. Materials and Methods

4.1. Chemicals

Ethyl dodecanoate (ethyl laurate), ethyl tridecanoate, propyl dodecanoate, methyl tetradecanoate, ethyl (E)-9-tetradecenoate (ethyl myristelaidate), ethyl (Z)-9-tetradecenoate (ethyl myristoleate), ethyl tetradecanoate (ethyl myristate), ethyl pentadecanoate, methyl (Z)-9-hexadecenoate, methyl hexadecanoate, ethyl (E)-9-hexadecenoate (ethyl palmitolelaidate) (Z)-9-hexadecenoate (ethyl palmitoleate), ethyl hexadecanoate (ethyl palmitate), ethyl (Z,Z)-octadeca-9,12-dienoate (ethyl linoleate), ethyl (Z)-9-octadecenoate (ethyl oleate), ethyl (E)-9-octadecenoate (ethyl elaidate), ethyl octadecanoate, hexadecane and straight-chain C8–C40 alkane standards were purchased from Sigma-Aldrich (St. Louis, MO, USA), Nu-Chek-Prep and INC (Minneapolis, MN, USA) or Alfa-Aesar. N-(3-methylbutyl)acetamide, N-(2-methylbutyl)propanamide, N-(3-methylbutyl)propanamide, N-(3-methylbutyl)isobutyramide were synthesized by reactions of an appropriate amine and an acid anhydride in water. Synthetic details are presented in Supplementary Materials. 2,8-Dimethyl-1,7-dioxaspiro[5.5]undecane was kindly provided by Ms. Sally Noushini.

4.2. Origin of Flies

Bactrocera tryoni larvae were collected from infested loquat fruits from a tree located in Marsfield NSW Australia (33.766080 S, 151.100722 E). A 500 mL plastic container with approximately 300 g of infested loquat fruits was placed on approximately a 1 cm deep layer of vermiculite in a 12.5 L plastic box for larvae to complete development and exit the fruit to pupate. Pupae were sieved and moved to a fine mesh cage (47.5 × 47.5 × 47.5 cm) (Megaview Bugdorm 4S4545, Taiwan) where the flies emerged. Identity of B. tryoni was confirmed by examining emerged flies under a stereomicroscope using the key to tropical tephritid fruit flies [117] and Jane Royer kindly provided additional confirmation for the identity of B. tryoni. Adults were separated by sex within three days of emergence, when still sexually immature [118], and thereafter kept as single-sex cohorts of 80 flies in mesh cages (30.5 × 30.5 × 30.5 cm) (Megaview Bugdorm 4S4545, Taiwan). Adult flies were provided with food by coating a small area of the top of mesh cage with a paste prepared by mixing 15 g sugar, 5 g hydrolysate yeast (MP Biomedicals LLC) and 4 mL tap water. Water was provided by placing an inverted vial (6 cm height, 4 cm diameter) full of water on a sponge on the top of the mesh cage. All cages were maintained in a controlled environment room at 25 ± 0.5 °C, 65 ± 5% RH and photoperiod of 11.5:0.5:11.5:0.5 light: dusk: dark: dawn.

4.3. Extraction of Cuticular Compounds

Fifteen two-day old sexually immature flies of each sex and fifteen 20-day old sexually mature virgin flies of each sex were used for the extraction of cuticular compounds. The flies were killed by placing them in a 5 mL plastic vial on dry ice. Frozen flies were allowed to defrost at room temperature for three minutes immediately before the following extraction procedure. n-Hexane was chosen as the extraction solvent because many cuticular compounds found in other tephritids are less, or non-polar [53,54,55,56,57,58,59,100]. A single fly was immersed in 400 µL of n-hexane that contained 1.5 μg/mL n-hexadecane (Sigma-Aldrich, St. Louis, MO, USA) in a 1.5 mL clear glass vial. n-Hexadecane was used as an internal standard to normalize peak areas for comparisons between groups. The vial containing a fly was allowed to stand for three minutes at room temperature, then the fly was removed from the n-hexane extract. If n-hexane extracts contained aqueous droplets, the droplets were removed by adding sodium sulfate (Na2SO4) (Sigma-Aldrich, St. Louis, MO, USA) and gravity filtration. If samples contained solid organic matter, the solid was removed by gravity filtration. Gravity filtration was achieved by filtering the sample through a glass wool plug on the neck of a Pasteur pipette. n-Hexane extracts were concentrated to 120 µL under a gentle stream of nitrogen gas and transferred to a glass insert (150 µL) in a clear 1.5 mL GC vial. The samples were stored at 4 °C until analyzed.

4.4. Gas Chromatography Mass Spectrometry (GC-MS) Analysis

GC-MS analysis was carried out on a Shimadzu GCMS TQ8040 spectrometer equipped with a split/splitless injector and SH Rtx-5MS (30 m × 0.25 mm, 0.25 µm film) fused silica capillary column. The carrier gas was helium (99.999%) (BOC, North Ryde, NSW, Australia) at a flow rate of 1.0 mL/min. An aliquot of 1 µL sample was injected at splitless mode where the injector temperature was 270 °C. The temperature program was set initially at 50 °C (1 min), increased to 280 °C at a rate of 10 °C/min, then increased to 302 °C at a rate of 2 °C/min. The ion source and transfer line temperatures were 200 and 290 °C, respectively. The ionization method was electron impact at a voltage of 70 eV. The spectra were obtained over a mass range of m/z 47–650. The data were analyzed by Shimadzu GCMS Postrun program (Shimadzu, Kyoto, Japan) and compared with the mass fragmentation patterns in NIST libraries (NIST17-1, NIST17-2), and authentic standards. Retention indices (KI) were calculated with the Kovats retention index equation [119] and compared with KI values published in the literature. The structures of methyl-branched alkanes were assigned by using methods described in previous studies [120,121,122]. Briefly, the chain length and number of methyl groups of a methyl-branched alkane were established by examining an equivalent chain length and molar mass. The position of a methyl group was then assigned by examining fragment ions.

4.5. Comparison between Groups

The data were normalized by dividing the GC peak areas of individual components by the peak areas of the internal standard. The data were not normally distributed and hence were log-transformed for statistical analyses. However, the raw data were used to generate the graphs. The data for normal and branched alkanes were analyzed by ANOVA to test for effects of sex and maturity on the amounts of compounds. The individual amides and alkanes were compared between sexes by t-test.

Supplementary Materials

The following are available online. Synthetic details of the amides and Figure S1. Mass spectra of 2-ethyl-8-methyl-1,7-dioxaspiro[5.5]undecane and methyl branched fatty acid esters.

Author Contributions

Conceptualization, V.M., P.W.T. and S.J.P.; methodology, V.M., P.W.T. and S.J.P.; formal analysis, G.P. and S.J.P.; investigation, C.C.-V., G.P., V.M. and S.J.P.; resources, P.W.T. and J.G.O.; writing—original draft preparation, S.J.P.; writing—review and editing, C.C.-V., G.P., J.G.O., P.W.T., V.M. and S.J.P.; supervision, J.G.O., P.W.T. and V.M.; project administration, P.W.T.; funding acquisition, P.W.T. All authors have read and agreed to the published version of the manuscript.


This research was conducted as part of the SITplus collaborative fruit fly program. Project Raising Q-fly Sterile Insect Technique to World Standard (HG14033) is funded by the Hort Frontiers Fruit Fly Fund, part of the Hort Frontiers strategic partnership initiative developed by Hort Innovation, with co-investment from Macquarie University and contributions from the Australian Government. This work received additional support from the Australian Research Council Industrial Transformation Training Centre (ITTC) for Fruit Fly Biosecurity Innovation (Project IC50100026), funded by the Australian Government, including a fellowship to SJP.


We thank Sally Noushini for providing 2,8-dimethyl-1,7-dioxaspiro[5,5]undecane, and Jane Royer for expert identification of B. tryoni.

Conflicts of Interest

The authors declare no conflict of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript, or in the decision to publish the results.


  1. Gibbs, A.; Crockett, E.L. The Biology of Lipids: Integrative and Comparative Perspectives. Am. Zool. 1998, 38, 265–267. [Google Scholar] [CrossRef]
  2. Howard, R.W.; Blomquist, G.J. Chemical Ecology and Biochemistry of Insect Hydrocarbons. Annu. Rev. Èntomol. 1982, 27, 149–172. [Google Scholar] [CrossRef]
  3. Lockey, K.H. Lipids of the insect cuticle: Origin, composition and function. Comp. Biochem. Physiol. Part B Comp. Biochem. 1988, 89, 595–645. [Google Scholar] [CrossRef]
  4. Blomquist, G.J.; Jackson, L.L. Chemistry and biochemistry of insect waxes. Prog. Lipid Res. 1979, 17, 319–345. [Google Scholar] [CrossRef]
  5. Blomquist, G.J.; Nelson, D.R.; De Renobales, M. Chemistry, biochemistry, and physiology of insect cuticular lipids. Arch. Insect Biochem. Physiol. 1987, 6, 227–265. [Google Scholar] [CrossRef]
  6. Ferveur, J.-F. Cuticular Hydrocarbons: Their Evolution and Roles in Drosophila Pheromonal Communication. Behav. Genet. 2005, 35, 279–295. [Google Scholar] [CrossRef]
  7. Gołębiowski, M.; Boguś, M.I.; Paszkiewicz, M.; Stepnowski, P. Cuticular lipids of insects as potential biofungicides: Methods of lipid composition analysis. Anal. Bioanal. Chem. 2010, 399, 3177–3191. [Google Scholar] [CrossRef]
  8. Gołębiowski, M.; Urbanek, A.; Pietrzak, A.; Naczk, A.M.; Bojke, A.; Tkaczuk, C.; Stepnowski, P. Effects of the entomopathogenic fungus Metarhizium flavoviride on the fat body lipid composition of Zophobas morio larvae (Coleoptera: Tenebrionidae). Naturwissenschaften 2020, 107, 1–11. [Google Scholar] [CrossRef]
  9. Gołębiowski, M.; Cerkowniak, M.; Urbanek, A.; Dawgul, M.; Kamysz, W.; Boguś, M.I.; Stepnowski, P. Identification and antifungal activity of novel organic compounds found in cuticular and internal lipids of medically important flies. Microbiol. Res. 2015, 170, 213–222. [Google Scholar] [CrossRef]
  10. Boguś, M.I.; Czygier, M.; Gołębiowski, M.; Kędra, E.; Kucińska, J.; Mazgajska, J.; Samborski, J.; Wieloch, W.; Włóka, E. Effects of insect cuticular fatty acids on in vitro growth and pathogenicity of the entomopathogenic fungus Conidiobolus coronatus. Exp. Parasitol. 2010, 125, 400–408. [Google Scholar] [CrossRef]
  11. Urbanek, A.; Szadziewski, R.; Stepnowski, P.; Boros-Majewska, J.; Gabriel, I.; Dawgul, M.; Kamysz, W.; Sosnowska, D.; Gołębiowski, M. Composition and antimicrobial activity of fatty acids detected in the hygroscopic secretion collected from the secretory setae of larvae of the biting midge Forcipomyia nigra (Diptera: Ceratopogonidae). J. Insect Physiol. 2012, 58, 1265–1276. [Google Scholar] [CrossRef] [PubMed]
  12. Gołębiowski, M.; Cerkowniak, M.; Dawgul, M.; Kamysz, W.; Boguś, M.I.; Stepnowski, P. The antifungal activity of the cuticular and internal fatty acid methyl esters and alcohols in Calliphora vomitoria. Parasitology 2013, 140, 972–985. [Google Scholar] [CrossRef]
  13. Howard, R.W.; Blomquist, G.J. Ecological, behavioral, and biochemical aspects of insect hydrocarbons. Annu. Rev. Èntomol. 2005, 50, 371–393. [Google Scholar] [CrossRef] [PubMed]
  14. Carlson, D.A.; Nelson, D.R.; Langley, P.A.; Coates, T.W.; Davis, T.L.; Der Linden, M.E.L.-V. Contact sex pheromone in the tsetse flyGlossina pallidipes (Austen) Identification and Synthesis. J. Chem. Ecol. 1984, 10, 429–450. [Google Scholar] [CrossRef]
  15. Ginzel, M.D.; Blomquist, G.J.; Millar, J.G.; Hanks, L.M. Role of contact pheromones in mate recognition in Xylotrechus colonus. J. Chem. Ecol. 2003, 29, 533–545. [Google Scholar] [CrossRef]
  16. Wilson, E.O. Chemical Communication within Animal Species; Academic Press, Inc.: New York, NY, USA, 1970; pp. 133–155. [Google Scholar]
  17. Xue, H.-J.; Segraves, K.A.; Wei, J.; Zhang, B.; Nie, R.-E.; Li, W.-Z.; Yang, X.-K. Chemically mediated sexual signals restrict hybrid speciation in a flea beetle. Behav. Ecol. 2018, 29, 1462–1471. [Google Scholar] [CrossRef]
  18. Dettner, K.; Liepert, C. Chemical Mimicry and Camouflage. Annu. Rev. Entomol. 1994, 39, 129–154. [Google Scholar] [CrossRef]
  19. Heifetz, Y.; Voet, H.; Applebaum, S.W. Factors affecting behavioral phase transition in the desert locust, Schistocerca gregaria (Forskål) (Orthoptera: Acrididae). J. Chem. Ecol. 1996, 22, 1717–1734. [Google Scholar] [CrossRef]
  20. Carlson, D.A.; Mayer, M.S.; Silhacek, D.L.; James, J.D.; Beroza, M.; Bierl, B.A. Sex Attractant Pheromone of the House Fly: Isolation, Identification and Synthesis. Science 1971, 174, 76–78. [Google Scholar] [CrossRef]
  21. Jennings, J.H.; Etges, W.J.; Schmitt, T.; Hoikkala, A. Cuticular hydrocarbons of Drosophila montana: Geographic variation, sexual dimorphism and potential roles as pheromones. J. Insect Physiol. 2014, 61, 16–24. [Google Scholar] [CrossRef]
  22. Roelofs, W.L.; Liu, W.; Hao, G.; Jiao, H.; Rooney, A.P.; Linn, J.C.E. Evolution of moth sex pheromones via ancestral genes. Proc. Natl. Acad. Sci. USA 2002, 99, 13621–13626. [Google Scholar] [CrossRef] [PubMed]
  23. Mant, J.; Brändli, C.; Vereecken, N.J.; Schulz, C.M.; Francke, W.; Schiestl, F.P. Cuticular Hydrocarbons as Sex Pheromone of the Bee Colletes cunicularius and the Key to its Mimicry by the Sexually Deceptive Orchid, Ophrys exaltata. J. Chem. Ecol. 2005, 31, 1765–1787. [Google Scholar] [CrossRef] [PubMed]
  24. Nojima, S.; Shimomura, K.; Honda, H.; Yamamoto, I.; Ohsawa, K. Contact Sex Pheromone Components of the Cowpea Weevil, Callosobruchus maculatus. J. Chem. Ecol. 2007, 33, 923–933. [Google Scholar] [CrossRef]
  25. Keesey, I.W.; Koerte, S.; Retzke, T.; Haverkamp, A.; Hansson, B.S.; Knaden, M. Adult Frass Provides a Pheromone Signature for Drosophila Feeding and Aggregation. J. Chem. Ecol. 2016, 42, 739–747. [Google Scholar] [CrossRef]
  26. Mitaka, Y.; Matsuyama, S.; Mizumoto, N.; Matsuura, K.; Akino, T. Chemical identification of an aggregation pheromone in the termite Reticulitermes speratus. Sci. Rep. 2020, 10, 7424. [Google Scholar] [CrossRef]
  27. Hamilton, J.A.; Wada-Katsumata, A.; Schal, C. Role of Cuticular Hydrocarbons in German Cockroach (Blattodea: Ectobiidae) Aggregation Behavior. Environ. Èntomol. 2019, 48, 546–553. [Google Scholar] [CrossRef] [PubMed]
  28. Cooper, W.E.; Andersson, M. Sexual Selection. Copeia 1995, 756. [Google Scholar] [CrossRef]
  29. Blomquist, G.J.; Bagnères, A.-G. Introduction: History and overview of insect hydrocarbons. In Insect Hydrocarbons: Biology, Biochemistry, and Chemical Ecologyi; Bagnères, A.-G., Blomquist, G.J., Eds.; Cambridge University Press: Cambridge, UK, 2010; pp. 3–18. [Google Scholar]
  30. Luo, Y.; Zhang, Y.; Farine, J.; Ferveur, J.; Ramírez, S.; Kopp, A. Evolution of sexually dimorphic pheromone profiles coincides with increased number of male-specific chemosensory organs in Drosophila prolongata. Ecol. Evol. 2019, 9, 13608–13618. [Google Scholar] [CrossRef]
  31. Ngumbi, E.N.; Hanks, L.M.; Suarez, A.V.; Millar, J.G.; Berenbaum, M.R. Author Correction: Factors Associated with Variation in Cuticular Hydrocarbon Profiles in the Navel Orangeworm, Amyelois transitella (Lepidoptera: Pyralidae). J. Chem. Ecol. 2020, 46, 232. [Google Scholar] [CrossRef]
  32. Farine, J.-P.; Ferveur, J.-F.; Everaerts, C. Volatile Drosophila Cuticular Pheromones Are Affected by Social but Not Sexual Experience. PLoS ONE 2012, 7, e40396. [Google Scholar] [CrossRef]
  33. Dias, N.P.; Zotti, M.J.; Montoya, P.; Carvalho, I.R.; Nava, D.E. Fruit fly management research: A systematic review of monitoring and control tactics in the world. Crop. Prot. 2018, 112, 187–200. [Google Scholar] [CrossRef]
  34. Robledo, N.; Arzuffi, R.; Robledo-Quintos, N. Influence of Host Fruit and Conspecifics on the Release of the Sex Pheromone By Toxotrypana curvicauda Males (Diptera: Tephritidae). Environ. Èntomol. 2012, 41, 387–391. [Google Scholar] [CrossRef] [PubMed]
  35. Tan, H.K.; Nishida, R. Incorporation of Raspberry Ketone in the Rectal Glands of Males of the Queensland Fruit Fly, Bactrocera tryoni FROGGATT (Diptera: Tephritidae). Appl. Èntomol. Zool. 1995, 30, 494–497. [Google Scholar] [CrossRef]
  36. Jang, E.B.; Carvalho, L.A.F.N.; Chen, C.-C.; Siderhurst, M.S. Cucumber Lure Trapping of Zeugodacus cucurbitae (Diptera:Tephritidae) in Hawaii and Taiwan: Longevity and Nontargets Captures. J. Econ. Èntomol. 2016, 110, 201–207. [Google Scholar] [CrossRef]
  37. Royer, J.E.; De Faveri, S.G.; Lowe, G.E.; Wright, C.L. Cucumber volatile blend, a promising female-biased lure for B actrocera cucumis (French 1907) (Diptera: Tephritidae: Dacinae), a pest fruit fly that does not respond to male attractants. Austral Èntomol. 2014, 53, 347–352. [Google Scholar] [CrossRef]
  38. Siderhurst, M.S.; Jang, E.B. Female-Biased Attraction of Oriental Fruit Fly, Bactrocera dorsalis (Hendel), to a Blend of Host Fruit Volatiles From Terminalia catappa L. J. Chem. Ecol. 2006, 32, 2513–2524. [Google Scholar] [CrossRef]
  39. Siderhurst, M.S.; Jang, E.B. Attraction of female Oriental fruit fly, Bactrocera dorsalis, to Terminalia catappa fruit in wind tunnel and olfactometer tests. Formos. Entomol. 2006, 26, 45–55. [Google Scholar]
  40. Siderhurst, M.S.; Jang, E.B. Cucumber Volatile Blend Attractive to Female Melon Fly, Bactrocera cucurbitae (Coquillett). J. Chem. Ecol. 2010, 36, 699–708. [Google Scholar] [CrossRef]
  41. Tan, K.H. Fruit fly pests as pollinators of wild orchids. In Proceedings of the 7th International Symposium on Fruit Flies of Economic Importance, Salvador, Brazil, 10–15 September 2006; pp. 195–206. [Google Scholar]
  42. Tan, K.H.; Nishida, R. Zingerone in the floral synomone of Bulbophyllum baileyi (Orchidaceae) attracts Bactrocera fruit flies during pollination. Biochem. Syst. Ecol. 2007, 35, 334–341. [Google Scholar] [CrossRef]
  43. Tan, K.H.; Tan, L.T.; Nishida, R. Floral Phenylpropanoid Cocktail and Architecture of Bulbophyllum vinaceum Orchid in Attracting Fruit Flies for Pollination. J. Chem. Ecol. 2006, 32, 2429–2441. [Google Scholar] [CrossRef]
  44. Tan, K.-H.; Nishida, R. Synomone or kairomone?—Bulbophyllum apertum flower releases raspberry ketone to attract Bactrocera fruit flies. J. Chem. Ecol. 2005, 31, 497–507. [Google Scholar]
  45. Tan, K.-H.; Nishida, R.; Toong, Y.-C. Floral synomone of a wild orchid, Bulbophyllum cheiri, lures Bactrocera fruit flies for pollination. J. Chem. Ecol. 2002, 28, 1161–1172. [Google Scholar] [CrossRef] [PubMed]
  46. Jang, E.B.; Carvalho, L.A.; Stark, J.D. Attraction of Female Oriental Fruit Fly, Bactrocera dorsalis, to Volatile Semiochemicals from Leaves and Extracts of a Nonhost Plant, Panax (Polyscias guilfoylei) in Laboratory and Olfactometer Assays. J. Chem. Ecol. 1997, 23, 1389–1401. [Google Scholar] [CrossRef]
  47. Siderhurst, M.S.; Park, S.J.; Buller, C.N.; Jamie, I.M.; Manoukis, N.C.; Jang, E.B.; Taylor, P.W. Raspberry Ketone Trifluoroacetate, a New Attractant for the Queensland Fruit Fly, Bactrocera Tryoni (Froggatt). J. Chem. Ecol. 2016, 42, 156–162. [Google Scholar] [CrossRef]
  48. Kather, R.; Martin, S.J. Cuticular hydrocarbon profiles as a taxonomic tool: Advantages, limitations and technical aspects. Physiol. Èntomol. 2012, 37, 25–32. [Google Scholar] [CrossRef]
  49. Vanickova, L.; Hernández-Ortiz, V.; Bravo, I.S.J.; Dias, V.S.; Roriz, A.K.P.; Laumann, R.A.; Mendonça, A.D.L.; Paranhos, B.A.J.; Nascimento, R.R.D. Current knowledge of the species complex Anastrepha fraterculus (Diptera, Tephritidae) in Brazil. ZooKeys 2015, 540, 211–237. [Google Scholar] [CrossRef]
  50. Carlson, D.A.; Yocom, S.R. Cuticular hydrocarbons from six species of tephritid fruit flies. Arch. Insect Biochem. Physiol. 1986, 3, 397–412. [Google Scholar] [CrossRef]
  51. Sutton, B.D.; Carlson, D.A. Interspecific variation in tephritid fruit fly larvae surface hydrocarbons. Arch. Insect Biochem. Physiol. 1993, 23, 53–65. [Google Scholar] [CrossRef]
  52. Sutton, B.D.; Steck, G.J. Discrimination of Caribbean and Mediterranean Fruit Fly Larvae (Diptera: Tephritidae) by Cuticular Hydrocarbon Analysis. Fla. Èntomol. 1994, 77, 231. [Google Scholar] [CrossRef]
  53. Vaníčková, L.; Virgilio, M.; Tomčala, A.; Břízová, R.; Ekesi, S.; Hoskovec, M.; Kalinová, B.; Do Nascimento, R.R.; De Meyer, M. Resolution of three cryptic agricultural pests (Ceratitis fasciventris, C. anonae, C. rosa, Diptera: Tephritidae) using cuticular hydrocarbon profiling. Bull. Entomol. Res. 2014, 104, 631–638. [Google Scholar]
  54. Vanickova, L.; Břízová, R.; Mendonça, A.L.; Pompeiano, A.; Nascimento, R.R.D. Intraspecific variation of cuticular hydrocarbon profiles in theAnastrepha fraterculus(Diptera:Tephritidae) species complex. J. Appl. Èntomol. 2015, 139, 679–689. [Google Scholar] [CrossRef]
  55. Vanickova, L.; Břízová, R.; Pompeiano, A.; Ekesi, S.; De Meyer, M. Cuticular hydrocarbons corroborate the distinction between lowland and highland Natal fruit fly (Tephritidae, Ceratitis rosa) populations. ZooKeys 2015, 540, 507–524. [Google Scholar] [CrossRef] [PubMed]
  56. Vanickova, L.; Břízová, R.; Pompeiano, A.; Ferreira, L.L.; De Aquino, N.C.; Tavares, R.D.F.; Rodriguez, L.D.; Mendonça, A.D.L.; Canal, N.A.; Nascimento, R.R.D. Characterisation of the chemical profiles of Brazilian and Andean morphotypes belonging to the Anastrepha fraterculus complex (Diptera, Tephritidae). ZooKeys 2015, 540, 193–209. [Google Scholar] [CrossRef] [PubMed]
  57. Goh, S.; Ooi, K.; Chuah, C.; Yong, H.; Khoo, S.; Ong, S. Cuticular hydrocarbons from two species of Malaysian Bactrocera fruit flies. Biochem. Syst. Ecol. 1993, 21, 215–226. [Google Scholar] [CrossRef]
  58. Vanickova, L.; Nagy, R.; Pompeiano, A.; Kalinová, B. Epicuticular chemistry reinforces the new taxonomic classification of the Bactrocera dorsalis species complex (Diptera: Tephritidae, Dacinae). PLoS ONE 2017, 12, e0184102. [Google Scholar] [CrossRef]
  59. Shen, J.; Hu, L.; Zhou, X.; Dai, J.; Chen, B.; Li, S. Allyl-2,6-dimethoxyphenol, a female-biased compound, is robustly attractive to conspecific males of Bactrocera dorsalis at close range. Èntomol. Exp. Appl. 2019, 167, 811–819. [Google Scholar] [CrossRef]
  60. Clarke, A.R.; Powell, K.S.; Weldon, C.W.; Taylor, P.W. The ecology of Bactrocera tryoni (Diptera: Tephritidae): What do we know to assist pest management? Ann. Appl. Biol. 2011, 158, 26–54. [Google Scholar] [CrossRef]
  61. Gilchrist, A.S.; Dominiak, B.; Gillespie, P.S.; Sved, J.A. Variation in population structure across the ecological range of the Queensland fruit fly. Bactrocera tryoni. Aust. J. Zool. 2006, 54, 87. [Google Scholar] [CrossRef]
  62. Sutherst, R.W.; Collyer, B.S.; Yonow, T. The vulnerability of Australian horticulture to the Queensland fruit fly, Bactrocera (Dacus) tryoni, under climate change. Aust. J. Agric. Res. 2000, 51, 467. [Google Scholar] [CrossRef]
  63. Dominiak, B.; Ekman, J.H.; Broughton, S. Mass trapping and other management options for Mediterranean fruit fly and Queensland fruit fly in Australia. Gen. App. Entomol. 2016, 44, 1–8. [Google Scholar]
  64. Florec, V.; Sadler, R.J.; White, B.; Dominiak, B.C.; White, B. Choosing the battles: The economics of area wide pest management for Queensland fruit fly. Food Policy 2013, 38, 203–213. [Google Scholar] [CrossRef]
  65. Bellas, T.E.; Fletcher, B.S. Identification of the major components in the secretion from the rectal pheromone glands of the queensland fruit fliesDacus tryoni andDacus neohumeralis (Diptera: Tephritidae). J. Chem. Ecol. 1979, 5, 795–803. [Google Scholar] [CrossRef]
  66. Fletcher, B.S. Storage and Release of a Sex Pheromone by the Queensland Fruit Fly, Dacus tryoni (Diptera: Trypetidae). Nature 1968, 219, 631–632. [Google Scholar] [CrossRef] [PubMed]
  67. El-Sayed, A.M.; Venkatesham, U.; Unelius, C.R.; Sporle, A.; Pérez, J.; Taylor, P.W.; Suckling, D.M. Chemical Composition of the Rectal Gland and Volatiles Released by Female Queensland Fruit Fly, Bactrocera tryoni (Diptera: Tephritidae). Environ. Èntomol. 2019, 48, 807–814. [Google Scholar] [CrossRef]
  68. Pérez, J.; Park, S.J.; Taylor, P.W. Domestication modifies the volatile emissions produced by male Queensland fruit flies during sexual advertisement. Sci. Rep. 2018, 8, 16503. [Google Scholar] [CrossRef] [PubMed]
  69. Noushini, S.; Park, S.J.; Jamie, I.M.; Jamie, J.F.; Taylor, P.W. Sampling technique biases in the analysis of fruit fly pheromones: A case study of Queensland fruit fly. Sci. Rep. 2020, in press. [Google Scholar]
  70. Booth, Y.K.; Schwartz, B.D.; Fletcher, M.; Lambert, L.K.; Kitching, W.; De Voss, J.J. A diverse suite of spiroacetals, including a novel branched representative, is released by female Bactrocera tryoni (Queensland fruit fly). Chem. Commun. 2006, 42, 3975–3977. [Google Scholar] [CrossRef]
  71. Lin, Y.; Jin, T.; Zeng, L.; Lu, Y. Cuticular penetration of β-cypermethrin in insecticide-susceptible and resistant strains of Bactrocera dorsalis. Pestic. Biochem. Physiol. 2012, 103, 189–193. [Google Scholar] [CrossRef]
  72. Sultana, S.; Baumgartner, J.B.; Dominiak, B.C.; Royer, J.E.; Beaumont, L.J. Potential impacts of climate change on habitat suitability for the Queensland fruit fly. Sci. Rep. 2017, 7, 13025. [Google Scholar] [CrossRef]
  73. Weldon, C.W.; Taylor, P.W. Desiccation resistance of adult Queensland fruit flies Bactrocera tryoni decreases with age. Physiol. Èntomol. 2010, 35, 385–390. [Google Scholar] [CrossRef]
  74. Weldon, C.W.; Yap, S.; Taylor, P. Desiccation resistance of wild and mass-reared Bactrocera tryoni (Diptera: Tephritidae). Bull. Èntomol. Res. 2013, 103, 690–699. [Google Scholar] [CrossRef] [PubMed]
  75. Dominiak, B.C.; Westcott, A.E.; Barchia, I.M. Release of sterile Queensland fruit fly, Bactrocera > tryoni (Froggatt) (Diptera: Tephritidae), at Sydney, Australia. Aust. J. Exp. Agric. 2003, 43, 519–528. [Google Scholar] [CrossRef]
  76. Meats, A.; Duthie, R.; Clift, A.D.; Dominiak, B.C. Trials on variants of the Sterile Insect Technique (SIT) for suppression of populations of the Queensland fruit fly in small towns neighbouring a quarantine (exclusion) zone. Aust. J. Exp. Agric. 2003, 43, 389–395. [Google Scholar] [CrossRef]
  77. Reynolds, O.L.; Smallridge, C.J.; Cockington, V.G.; Penrose, L.D. The effect of release method and trial site on recapture rates of adult sterile Queensland fruit fly, Bactrocera tryoni (Froggatt) (Diptera: Tephritidae). Aust. J. Èntomol. 2011, 51, 116–126. [Google Scholar] [CrossRef]
  78. Noushini, S.; Pérez, J.; Park, S.J.; Holgate, D.; Alvarez, V.M.; Jamie, I.M.; Jamie, J.F.; Taylor, P.W. Attraction and Electrophysiological Response to Identified Rectal Gland Volatiles in Bactrocera frauenfeldi (Schiner). Molecules 2020, 25, 1275. [Google Scholar] [CrossRef]
  79. El-Sayed, A.M.; Heppelthwaite, V.J.; Manning, L.-A.; Gibb, A.R.; Suckling, D.M.; El-Sayed, A.M. Volatile Constituents of Fermented Sugar Baits and Their Attraction to Lepidopteran Species. J. Agric. Food Chem. 2005, 53, 953–958. [Google Scholar] [CrossRef]
  80. Pino, J.A.; Mesa, J.; Muñoz, Y.; Martí, M.P.; Marbot, R. Volatile Components from Mango (Mangifera indica L.) Cultivars. J. Agric. Food Chem. 2005, 53, 2213–2223. [Google Scholar] [CrossRef]
  81. Peppard, T.L. Volatile flavor constituents of Monstera deliciosa. J. Agric. Food Chem. 1992, 40, 257–262. [Google Scholar] [CrossRef]
  82. Silva, D.B.; Pott, A.; Oliveira, D.C.R. Analyses of the Headspace Volatile Constituents of Aerial Parts (leaves and stems), Flowers and Fruits of Bidens gardneri Bak. and Bidens sulphurea (Cav.) Sch.Bip. Using Solid-Phase Microextraction. J. Essent. Oil Res. 2010, 22, 560–563. [Google Scholar] [CrossRef]
  83. Isidorov, V.; Krajewska, U.; Dubis, E.N.; Jdanova, M. Partition coefficients of alkyl aromatic hydrocarbons and esters in a hexane-acetonitrile system. J. Chromatogr. A 2001, 923, 127–136. [Google Scholar] [CrossRef]
  84. Robinson, A.L.; Adams, D.O.; Boss, P.K.; Heymann, H.; Solomon, P.S.; Trengove, R. Influence of Geographic Origin on the Sensory Characteristics and Wine Composition of Vitis vinifera cv. Cabernet Sauvignon Wines from Australia. Am. J. Enol. Vitic. 2012, 63, 467–476. [Google Scholar] [CrossRef]
  85. Palmeira, S.F.; Moura, F.D.S.; Alves, V.D.L.; De Oliveira, F.M.; Bento, E.S.; Conserva, L.M.; Andrade, E.D.A. Neutral components from hexane extracts ofCroton sellowii. Flavour Fragr. J. 2004, 19, 69–71. [Google Scholar] [CrossRef]
  86. Pino, J.A.; Almora, K.; Marbot, R. Volatile components of papaya (Carica papaya L., Maradol variety) fruit. Flavour Fragr. J. 2003, 18, 492–496. [Google Scholar] [CrossRef]
  87. Palmeira, S.F.; Conserva, L.M.; Andrade, E.H.D.A.; Guilhon, G.M.S.P. Analysis by GC-MS of the hexane extract of the aerial parts of Aristolochia acutifolia Duchtr. Flavour Fragr. J. 2001, 16, 85–88. [Google Scholar] [CrossRef]
  88. Demyttenaere, J.C.; Martínez, J.I.S.; Verhé, R.; Sandra, P.; De Kimpe, N.; Sanchezmartinez, J. Analysis of volatiles of malt whisky by solid-phase microextraction and stir bar sorptive extraction. J. Chromatogr. A 2003, 985, 221–232. [Google Scholar] [CrossRef]
  89. Andriamaharavo, N.R. Retention Data: NIST Mass Spectrometry Data Center. 2014. Available online: (accessed on 10 September 2020).
  90. Tuberoso, C.; Barra, A.; Angioni, A.; Sarritzu, E.; Pirisi, F.M. Chemical Composition of Volatiles in Sardinian Myrtle (Myrtus communis L.) Alcoholic Extracts and Essential Oils. J. Agric. Food Chem. 2006, 54, 1420–1426. [Google Scholar] [CrossRef]
  91. Kenig, F.; Damsté, J.S.S.; Dalen, A.K.-V.; Rijpstra, W.C.; Huc, A.Y.; De Leeuw, J.W. Occurrence and origin of mono-, di-, and trimethylalkanes in modern and Holocene cyanobacterial mats from Abu Dhabi, United Arab Emirates. Geochim. Cosmochim. Acta 1995, 59, 2999–3015. [Google Scholar] [CrossRef]
  92. Nelson, D.R.; Charlet, L.D. Cuticular hydrocarbons of the sunflower beetle, Zygogramma exclamationis. Comp. Biochem. Physiol. Part B Biochem. Mol. Boil. 2003, 135, 273–284. [Google Scholar] [CrossRef]
  93. Carlson, D.; Geden, C.; Bernier, U. Identification of Pupal Exuviae of Nasonia vitripennis and Muscidifurax raptorellus Parasitoids Using Cuticular Hydrocarbons. Boil. Control. 1999, 15, 97–106. [Google Scholar] [CrossRef]
  94. Lockey, K.H. Cuticular hydrocarbons of adult Cylindrinotus laevioctostriatus (Goeze) and Phylan gibbus (Fabricius) (Coleoptera: Tenebrionidae). Insect Biochem. 1981, 11, 549–561. [Google Scholar] [CrossRef]
  95. Katritzky, A.R.; Chen, K.; Maran, U.; Carlson, D.A. QSPR correlation and predictions of GC retention indexes for methyl-branched hydrocarbons produced by insects. Anal. Chem. 2000, 72, 101–109. [Google Scholar] [CrossRef] [PubMed]
  96. Lockey, K.H. The adult cuticular hydrocarbons of Tenebrio molitor L. and Tenebrio obscurus F. (Coleoptera: Tenebrionidae). Insect Biochem. 1978, 8, 237–250. [Google Scholar] [CrossRef]
  97. Bernier, U.R.; Carlson, D.A.; Geden, C.J. Gas chromatography/mass spectrometry analysis of the cuticular hydrocarbons from parasitic wasps of the genus Muscidifurax. J. Am. Soc. Mass Spectrom. 1998, 9, 320–332. [Google Scholar] [CrossRef]
  98. Ekanayake, D. [Leave for Adam-Compressed File] The Mating System and Courtship Behaviour of the Queensland Fruit Fly, Bactrocera tryoni (Froggatt) (Diptera: Tephritidae); Queensland University of Technology: Brisbane, Australia, 2017; p. 271. [Google Scholar]
  99. Blomquist, G.J.; Tittiger, C.; Jurenka, R. Cuticular Hydrocarbons and Pheromones of Arthropods. In Hydrocarbons, Oils and Lipids: Diversity, Origin, Chemistry and Fate. Available online: (accessed on 10 September 2020).
  100. Vanickova, L.; Svatoš, A.; Kroiss, J.; Kaltenpoth, M.; Nascimento, R.R.D.; Hoskovec, M.; Břízová, R.; Kalinová, B. Cuticular Hydrocarbons of the South American Fruit Fly Anastrepha fraterculus: Variability with Sex and Age. J. Chem. Ecol. 2012, 38, 1133–1142. [Google Scholar] [CrossRef]
  101. Lucas, C.; Pho, D.; Fresneau, D.; Jallon, J. Hydrocarbon circulation and colonial signature in Pachycondyla villosa. J. Insect Physiol. 2004, 50, 595–607. [Google Scholar] [CrossRef]
  102. Soroker, V.; Vienne, C.; Hefetz, A. Hydrocarbon dynamics within and between nestmates in Cataglyphis niger (Hymenoptera: Formicidae). J. Chem. Ecol. 1995, 21, 365–378. [Google Scholar] [CrossRef]
  103. Bagnères, A.-G.; Trabalon, M.; Blomquist, G.J.; Schulz, S. Waxes of the social spider Anelosimus eximius (Araneae, Theridiidae): Abundance of noveln-propyl esters of long-chain methyl-branched fatty acids. Arch. Insect Biochem. Physiol. 1997, 36, 295–314. [Google Scholar] [CrossRef]
  104. Chinta, S.P.; Goller, S.; Uhl, G.; Schulz, S. Identification and Synthesis of Branched Wax-type Esters, Novel Surface Lipids from the Spider Argyrodes elevates (Araneae: Theridiidae). Chem. Biodivers. 2016, 13, 1202–1220. [Google Scholar] [CrossRef]
  105. Gu, P.; Welch, W.H.; Blomquist, G.J. Methyl-branched fatty acid biosynthesis in the German cockroach, Blatella germanica: Kinetic studies comparing a microsomal and soluble fatty acid synthetase. Insect Biochem. Mol. Boil. 1993, 23, 263–271. [Google Scholar] [CrossRef]
  106. Wang, H.-L.; Brattström, O.; Brakefield, P.M.; Francke, W.; Löfstedt, C. Identification and Biosynthesis of Novel Male Specific Esters in the Wings of the Tropical Butterfly, Bicyclus martius sanaos. J. Chem. Ecol. 2014, 40, 549–559. [Google Scholar] [CrossRef]
  107. Morgan, E. Biosynthesis in Insects; Royal Society of Chemistry (RSC): Cambridge, UK, 2006. [Google Scholar]
  108. Gibbs, A.; Pomonis, J. Physical properties of insect cuticular hydrocarbons: The effects of chain length, methyl-branching and unsaturation. Comp. Biochem. Physiol. Part B Biochem. Mol. Boil. 1995, 112, 243–249. [Google Scholar] [CrossRef]
  109. Foley, B.R.; Telonis-Scott, M. Quantitative genetic analysis suggests causal association between cuticular hydrocarbon composition and desiccation survival in Drosophila melanogaster. Heredity 2010, 106, 68–77. [Google Scholar] [CrossRef] [PubMed]
  110. Gefen, E.; Talal, S.; Brendzel, O.; Dror, A.; Fishman, A. Variation in quantity and composition of cuticular hydrocarbons in the scorpion Buthus occitanus (Buthidae) in response to acute exposure to desiccation stress. Comp. Biochem. Physiol. Part A Mol. Integr. Physiol. 2015, 182, 58–63. [Google Scholar] [CrossRef] [PubMed]
  111. Ferveur, J.-F.; Cortot, J.; Rihani, K.; Cobb, M.; Everaerts, C. Desiccation resistance: Effect of cuticular hydrocarbons and water content in Drosophila melanogaster adults. PeerJ 2018, 6, 4318. [Google Scholar] [CrossRef] [PubMed]
  112. Pike, N.; Wang, W.Y.; Meats, A. The likely fate of hybrids of Bactrocera tryoni and Bactrocera neohumeralis. Heredity 2003, 90, 365–370. [Google Scholar] [CrossRef] [PubMed]
  113. Popa-Báez, Á.-D.; Catullo, R.; Lee, S.F.; Yeap, H.L.; Mourant, R.G.; Frommer, M.; Sved, J.A.; Cameron, E.C.; Edwards, O.R.; Taylor, P.W.; et al. Genome-wide patterns of differentiation over space and time in the Queensland fruit fly. Sci. Rep. 2020, 10, 1–13. [Google Scholar] [CrossRef]
  114. Fedina, T.Y.; Kuo, T.-H.; Dreisewerd, K.; Dierick, H.A.; Yew, J.Y.; Pletcher, S.D. Dietary Effects on Cuticular Hydrocarbons and Sexual Attractiveness in Drosophila. PLoS ONE 2012, 7, e49799. [Google Scholar] [CrossRef]
  115. Otte, T.; Hilker, M.; Geiselhardt, S. The effect of eietary fatty acids on the cuticular hydrocarbon phenotype of an herbivorous insect and consequences for mate recognition. J. Chem. Ecol. 2015, 41, 32–43. [Google Scholar] [CrossRef]
  116. Otte, T.; Hilker, M.; Geiselhardt, S. Phenotypic Plasticity of Cuticular Hydrocarbon Profiles in Insects. J. Chem. Ecol. 2018, 44, 235–247. [Google Scholar] [CrossRef]
  117. Drew, R.A.I. The tropical fruit flies (Diptera: Tephritidae: Dacinae) of the Australian and Oceanian regions. Mem Queensl. Mus. 1989, 26, 521. [Google Scholar]
  118. Pérez-Staples, D.; Prabhu, V.; Taylor, P.W. Post-teneral protein feeding enhances sexual performance of Queensland fruit flies. Physiol. Èntomol. 2007, 32, 225–232. [Google Scholar] [CrossRef]
  119. Kováts, E. Gas-chromatographische Charakterisierung organischer Verbindungen. Teil 1: Retentionsindices aliphatischer Halogenide, Alkohole, Aldehyde und Ketone. Helvetica Chim. Acta 1958, 41, 1915–1932. [Google Scholar] [CrossRef]
  120. McCarthy, E.D.; Han, J.; Calvin, M. Hydrogen atom transfer in mass spectrometric fragmentation patterns of saturated aliphatic hydrocarbons. Anal. Chem. 1968, 40, 1475–1480. [Google Scholar] [CrossRef]
  121. Nelson, D.R.; Sukkestad, D.R. Normal and branched aliphatic hydrocarbons from the eggs of the tobacco hornworm. Biochem. 1970, 9, 4601–4611. [Google Scholar] [CrossRef]
  122. Nelson, D.R.; Sukkestad, D.R.; Zaylskie, R.G. Mass spectra of methyl-branched hydrocarbons from eggs of the tobacco hornworm. J. Lipid Res. 1972, 13, 413–421. [Google Scholar]
Sample Availability: Samples of the compounds are not available from the authors.
Figure 1. Typical chromatograms of hexane extracts of immature and mature female and male B. tryoni. (A) immature female; (B) immature male; (C) mature female; (D) mature male.
Figure 1. Typical chromatograms of hexane extracts of immature and mature female and male B. tryoni. (A) immature female; (B) immature male; (C) mature female; (D) mature male.
Molecules 25 04185 g001
Figure 2. Representative chromatogram sections. (A) Non-alkanes in a chromatogram of a mature female B. tryoni, which includes all compounds found in mature and immature males and females; (B) Hydrocarbons (alkanes) section of chromatogram from a mature female B. tryoni. Note that compounds in B are qualitatively identical in immature and mature females and males.
Figure 2. Representative chromatogram sections. (A) Non-alkanes in a chromatogram of a mature female B. tryoni, which includes all compounds found in mature and immature males and females; (B) Hydrocarbons (alkanes) section of chromatogram from a mature female B. tryoni. Note that compounds in B are qualitatively identical in immature and mature females and males.
Molecules 25 04185 g002
Figure 3. Internal standard-normalized peak areas of N-(3-methylbutyl)acetamide (A1), N-(2-methylbutyl)propanamide (A3), N-(3-methylbutyl)propenamide (A4) and N-(3-methylbutyl)isobutyramide (A6) in section A in Figure 2 in female and male B. tryoni. Standardized peak areas were obtained by dividing the peak area of a compound by the peak area of the n-hexadecane internal standard. Error bars represent standard errors. The results of t-test comparisons between the sexes are shown (ns is not significant; * p < 0.05, ** p < 0.01).
Figure 3. Internal standard-normalized peak areas of N-(3-methylbutyl)acetamide (A1), N-(2-methylbutyl)propanamide (A3), N-(3-methylbutyl)propenamide (A4) and N-(3-methylbutyl)isobutyramide (A6) in section A in Figure 2 in female and male B. tryoni. Standardized peak areas were obtained by dividing the peak area of a compound by the peak area of the n-hexadecane internal standard. Error bars represent standard errors. The results of t-test comparisons between the sexes are shown (ns is not significant; * p < 0.05, ** p < 0.01).
Molecules 25 04185 g003
Figure 4. Internal standard-normalized peak areas of individual cuticular hydrocarbons (the alkanes in section B in Figure 2) in the two sexes of B. tryoni. Standardized peak areas were obtained by dividing the peak area of a compound by the peak area of the n-hexadecane internal standard. B28 and B29 co-eluted and hence the sum of their amounts are presented together. Error bars represent standard errors. The results of t-test comparisons between the sexes are shown (ns is not significant; * p < 0.05; **p < 0.01; *** p < 0.001).
Figure 4. Internal standard-normalized peak areas of individual cuticular hydrocarbons (the alkanes in section B in Figure 2) in the two sexes of B. tryoni. Standardized peak areas were obtained by dividing the peak area of a compound by the peak area of the n-hexadecane internal standard. B28 and B29 co-eluted and hence the sum of their amounts are presented together. Error bars represent standard errors. The results of t-test comparisons between the sexes are shown (ns is not significant; * p < 0.05; **p < 0.01; *** p < 0.001).
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Table 1. The compounds identified in hexane washes of B. tryoni that eluted early in chromatograms (section A in Figure 2).
Table 1. The compounds identified in hexane washes of B. tryoni that eluted early in chromatograms (section A in Figure 2).
NoIdentityMMKIRef.KI (Ref)Characteristic/Diagnostic EI Ions
A1 *N-(3-Methylbutyl)acetamide129.1211311137 [78]129 (M+), 114, 86, 73, 60 (CH3COHNH2+)
A22,8-Dimethyl-1,7-dioxaspiro[5,5]undecane184.1511401147 [78]184 (M+), 140, 115/112 (M-C5H8/C5H8O), 97, 69
A3 *N-(2-Methylbutyl)propanamide143.131198 143 (M+), 114, 86, 74, 57
A4 *N-(3-Methylbutyl)propanamide143.131204 143 (M+), 128 (M-CH3), 114 (M-C2H5), 100, 87, 74, 57
A5 #2-Ethyl-8-methyl-1,7-dioxaspiro[5,5]undecane198.1612301237 [78]198 (M+), 169, 129/126 (M-C5H8/C5H8O), 115/112 (C6H10/C6H10O), 97, 83, 69, 55
A6 *N-(3-Methylbutyl)isobutyramide157.151233 157 (M+), 142, 101, 71, 57
A7Ethyl dodecanoate (ethyl laurate)228.3815911593 [79]228 (M+), 183, 157, 115, 101, 88, 73, 70, 60 (CH3CO=OH+)
A8 #Ethyl 6-methyldodecanoate242.221662 242 (M+), 213, 199, 185, 157, 143, 101, 88, 83, 70, 55
A9Propyl dodecanoate242.2216801685 [80]242 (M+), 201, 183, 157, 143, 115, 102, 61 (C3H7OH2+, base peak)
A10Ethyl tridecanoate242.2216911695 [81]242 (M+), 199, 197, 157, 101, 88
A11Methyl tetradecanoate242.2217221724 [82]242 (M+), 157, 143, 101, 87, 74
A12Ethyl (E)-9-tetradecenoate (ethyl myristolaidate)254.221769 254 (M+), 208/209 (loss of EtOH/EtO), 166, 124, 88, 55
A13Ethyl (Z)-9-tetradecenoate (ethyl myristoleate)254.221778 254 (M+), 208/209 (loss of EtOH/EtO), 166, 124, 88, 55
A14Ethyl tetradecanoate (ethyl myristate)256.4317901793 [80]256 (M+), 213, 157, 101, 88
A15 #Ethyl 4-methyltetradecanoate270.261836 270 (M+), 213, 101 (M-C12H25, base peak), 88
A16 #Ethyl 12-methyltetradecanoate270.261862 270 (M+), 227, 213, 157, 101, 88
A17 #Propyl tetradecanoate270.2618871893 [83]270 (M+), 229, 211, 172, 129, 102, 61 (C3H7OH2+, base peak)
A18Ethyl pentadecanoate270.2618901897 [84]270 (M+), 227, 199, 157, 101, 88
A19Methyl (Z)-9-hexadecenoate268.4419021909 [78]268 (M+), 236/237 (loss of MeOH/MeO), 194, 152, 96, 74, 55
A20Methyl hexadecanoate270.2619231927 [85]270 (M+), 227, 143, 87, 74
A21Ethyl (E)-9-hexadecenoate (ethyl palmitelaidate)282.261965 282 (M+), 236/237 (loss of EtOH/EtO), 194, 152, 96, 88, 69, 55
A22Ethyl (Z)-9-hexadecenoate (ethyl plamitoleate)282.2619701975 [86] 282 (M+), 236/237 (loss of EtOH/EtO), 194, 152, 96, 88, 69, 55
A23Ethyl hexadecanoate (ethyl palmitate)284.2719901993 [80]284 (M+), 241, 157, 101, 88
A24 #Ethyl 15-methylhexadecanoate298.292029 298 (M+), 255, 157 (M-C10H21), 101, 88
A25 #Ethyl 4-methylhexadecanoate298.292035 298 (M+), 241 (M-C4H9), 101 (base peak), 88
A26 #Ethyl 14-methylhexadecanoate298.292062 298 (M+), 269, 255, 241, 199, 157, 101, 88
A27 #Propyl 9-hexadecenoate296.272067 296 (M+) 281, 237, 194 (M-C3H7COOCH2)
A28Ethyl (Z,Z)-octadeca-9,12-dienoate (ethyl linoleate)308.2721582155 [87]308 (M+), 262/263 (loss of EtOH/EtO), 178, 135, 95, 81
A29Ethyl (Z)-9-octadecenoate (ethyl oleate)310.2921682168 [88]310 (M+), 264/265 (loss of EtOH/EtO), 222, 180, 97, 55
A30Ethyl (E)-9-octadecenoate (ethyl elaidate)310.2921712174 [89]310 (M+), 264/265 (loss of EtOH/EtO), 222, 180, 97, 55
A31Ethyl octadecanoate312.5421902191 [90]312 (M+), 269, 157, 101, 88
A32 #Ethyl 11-eicosenoate338.572366 338 (M+), 292/293 (M-EtOH/EtO), 250, 208, 97, 55
MM: molecular mass, KI: Kovats’ retention index, Ref.KI (Ref): reference KI if available for a similar column type, active phase and temperature conditions, with references in parentheses; * indicates that the compound is present in both sexes of mature adults, all other compounds being mature female specific; # indicates the compound was only tentatively identified.
Table 2. The tentatively identified cuticular hydrocarbons found in n-hexane washes of B. tryoni (section B in Figure 1).
Table 2. The tentatively identified cuticular hydrocarbons found in n-hexane washes of B. tryoni (section B in Figure 1).
NoIdentityMMKIRef.KI (Ref)Characteristic/Diagnostic EI Ions
B111-; 13-; 15-MeC29422.8229292932 [91]280/281,168/169; 252/253, 196/197; 224/225(s)
B27-MeC29422.8229462940 [51]336/337, 112/113
B35-MeC29422.8229522949 [91]364/365, 84/85
B49,13-DiMeC29436.5029662963 [91]322/323, 140/141, 252/253, 210/211
B57,11-DiMeC29436.502970 350/351, 112/113, 280/281, 182/183
B63-MeC29422.8229762973 [91]392/393, 56/57
B75,11-DiMeC29; 5,13-DiMeC29436.5029862983 [91]378/379, 84/85,280/281, 182/183; 378/379, 84/85, 280/281, 210/211
B84, x, 22-TriMeC29 (x = 14 or 16)450.523009 392/393, 84/85, 252/253, 224/225, 336/337, 126/127
B912-Me; 14-MeC30436.5030253031 [92]280/281,182/183; 252/253, 210/211
B108-MeC30436.5030343040 [93]336/337, 126/127
B116-MeC30436.5030413045 [93]364/365, 98/99
B124-MeC30436.5030553065 [93]392/393, 70/71
B138,12-DiMeC30; 8,14-DiMeC30450.5230613064 [94]350/351, 126/127, 280/281, 196/197; 350/351, 126/127, 253/252, 225.224
B146,14-DiMeC30; 6,12-DiMeC30450.503071 378/379, 98/99, 252/253, 224/225; 378/379, 98/99, 280/281, 196/197
B154,12-DiMeC30; 4,14-DiMeC30; 4,20-DiMeC30450.5230883098 [94]406/407, 70/71, 280/281, 196/197; 406/407, 70/71, 225/224, 253/252; 406/407, 70/71, 309/308, 169/168
B16n-C31436.503100 436
B174,8,12-TriMeC30; 4,8,14-TriMeC30; 4,8,20-TriMeC30464.533115 70/71, 420/421, 350/351, 140/141, 280/291, 210/211;
70/71, 420/421, 350/351, 140/141, 252/253, 238/239;
70/71, 420/421, 350/351, 140/141, 322/323, 168/169
B1811-; 13-; 15-MeC31450.5231293130 [92]308/309, 168/169; 280/281, 196/197; 250/251, 224/225
B197-MeC31; 9-MeC31450.5231373140 [93]364/365, 112/113; 336/337, 141/140
B205-MeC31450.5231473150 [93]392/393, 84/85
B2111,15-DiMeC31464.5331533155 [93]322/323, 168/169, 252/253, 238/239
B229,13-DiMeC31; 9,15-DiMeC31; 11,13-DiMeC31; 13,15-DiMeC31464.5331573159 [92]350/351, 140/141, 280/281, 210/211; 350/351, 140/141, 252/253, 238/239; 322/323, 168/169, 280/281, 210/211; 294/295, 196/197, 252/253, 238/239
B237,13-DiMeC31;7,15-DiMeC31464.533164 378/379, 112/113, 280/281, 210/211; 378/379, 112/113, 252/253, 238/239
B243-MeC31450.5231703172 [91]420/421, 56/57
B255,11-DiMeC31; 5,13-DiMeC31464.5331783180 [95]406/407, 84/85, 182/183, 308/309; 406/407, 84/85, 280/281, 210/211
B2612-; 14-; 16-MeC32464.5332233225 [57]308/309, 182/183; 280/281, 210/211; 252/253, 238/239
B278-; 10-MeC32464.5332253225 [57]364/365, 126/127; 336/337, 154/155
B2810,14-DiMeC32478.5532333254 [94]350/351, 154/155, 280/281, 224/225
B298,12-DiMeC32; 8,14-DiMeC32; 8,16-DiMeC32478.5532573263 [96]378/379, 126/127, 308/309, 196/197; 378/379, 126/127, 280/281, 224/225; 378/379, 126/127, 252/253 (s)
B304-MeC32464.5332673265 [97]420/421, 70/71
B316,16-DiMeC32478.553285 406/407, 98/99, 252/253
B329-; 11-MeC33478.5533273335 [93]364/365, 140/141; 336/337, 168/169
B3313-; 15-; 17-MeC33478.5533303335 [93]308/309, 196/197; 280/281, 224/225; 252/253 (s)
B349,23-DiMeC33492.563359 378/379, 140/141, 350/351, 168/169
MM: molecular mass; KI: Kovats retention index; Ref.KI (Ref): reference KI if available with references in parenthesis; (s) indicates ions from symmetrical structures.
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