- Research article
- Open Access
Pheromone gland development and monoterpenoid synthesis specific to oviparous females in the pea aphid
© The Author(s). 2018
Received: 14 November 2017
Accepted: 3 April 2018
Published: 11 May 2018
Aphids display “cyclic parthenogenesis,” in which parthenogenetically and sexually reproducing morphs seasonally alternate in the aphid annual life cycles. There are various characteristics that differ between asexual viviparous and sexual oviparous females. In oviparous females, swollen cuticular structures (~ 10 μm in diameter), called “scent plaques,” are scattered on the surface of hind tibias, and secrete monoterpenoid sex pheromones. However, the developmental processes of the pheromone glands and the biosynthetic pathways of monoterpenoid pheromones have yet to be elucidated.
Comparisons of the developmental processes that form hind tibias between sexual and parthenogenetic females revealed that, in sexual females, the epithelial tissues in proximal parts of hind tibias become columnar in fourth instar nymphs, and circular pheromone glands with Class 1 gland cells appear in adults, although they do not appear in parthenogenetic females. Furthermore, by comparing the expression levels of genes involved in the mevalonate pathway, which is required for monoterpenoid synthesis, we show that genes that encode the downstream enzymes in the pathway are highly expressed in hind tibias of sexual females.
Glandular tissues of scent plaque are differentiated from the fourth instar in sexual females, while parthenogenetic females lack the glandular cells. Only the downstream steps of the mevalonate pathway appear to occur in scent plaques on hind tibias of sexual females, although the upstream steps may occur somewhere in other body parts.
Sexual aphid females are known to secrete sex pheromones to attract males [9–12]. In many aphid species, including the pea aphid Acyrthosiphon pisum, two monoterpenoids, (4aS, 7S, 7aR)-nepetalactone and (1R, 4aS, 7S, 7aR)-nepetalactol, have been identified as sex pheromones [13–17]. The sex pheromones are thought to be secreted from circular bulge structures, called “scent plaques,” scattered on their hind tibias, which was supported by the findings that oviparous females whose hind legs were mutilated attracted fewer males [9, 10, 18, 19]. Since scent plaques are the sex pheromone glands of aphids, the epithelial cells under the scent plaques should be differentiated into specific gland cells for secreting sex pheromones, although the detailed histological structures of gland cells and the developmental process remain unknown.
Generally, terpenoids can be synthesized by several biosynthetic pathways. The mevalonate pathway is the most well-known, and is essential to isoprenoid synthesis in eukaryotes, in which acetyl-CoA is converted to isopentenyl diphosphate (IPP) and dimethylallyl diphosphate (DMAPP) through several enzymatic reactions . In some insects, monoterpenoid pheromones have been shown to be synthesized through the mevalonate pathway [21, 22]. We hypothesized that the mevalonate pathway is also utilized for sex pheromone synthesis in aphids.
In the present study, hind-tibia structure and development were compared between oviparous and parthenogenetic females, and detailed scent plaque structures were histologically observed to examine whether the scent plaque possesses characteristics of a pheromone gland. In addition, although the place of secretion is thought to be scent plaques, the places of reactions in the mevalonate pathway may not necessarily be scent plaques. In order to examine from which step of the mevalonate pathway was localized in hind tibias of oviparous aphids, expression levels of genes that encode enzymes in the mevalonate pathway were compared by realtime quantitative PCR (qPCR). Based on the KEGG pathway database (http://www.genome.jp/kegg/pathway.html), seven major enzymes were chosen: acetoacetyl-CoA thiolase (AACT), HMG-CoA (3-hydroxy-3-methylglutaryl-coenzyme A) synthetase (HMGS), HMG-CoA reductase (HMGR), mevalonate kinase (MevK), phosphomevalonate kinase (MevPK), mevalonate 5-diphosphate decarboxylase (MevPPD), and isopentenyl pyrophosphate isomerase (IPPI) [23–25]. In addition, farnesyl diphosphate synthetase (FPPS) was also included, as it converts the final products in the mevalonate pathway, i.e., IPP and DMAPP, into geranyl diphosphate (GPP), which is a monoterpenoid precursor, as shown in the bark beetle Ips pini [22, 24, 26].
The ApL strain of the pea aphid, Acrthosiphon pisum, which was suitable to the induction of sexual generation, was used in this study. This strain was originally obtained from Sapporo, Hokkaido (referred to as Sap05Ms2 in ) and used as the ApL strain in the following studies . The A. pisum genome was sequenced and is publicly available . During parthenogenetic generations, aphids were reared in plastic tubes (diameter: 2.5 cm, height: 10 cm), in each of which a vetch seedling (Vicia faba) was placed on wet vermiculite. The tubes were kept under long-day and room-temperature conditions (16 h light: 8 h dark; 20 °C) .
Induction of parthenogenetic and oviparous females
The induction of viviparous and oviparous females in the ApL strain was performed following the protocols previously described . Briefly, a first-instar aphid produced by a single viviparous parthenogenetic female was isolated and reared in a rectangular plastic case (width: 6 cm, height: 10 cm, depth: 2.5 cm) with a vetch seedling at 15 °C under short-day conditions (8 h light: 16 h dark). After becoming an adult, the parthenogenetic female produced approximately 30 first-instar nymphs, defined as Short-day Generation 1 (“SG1”), during the first 16 days after the onset of larviposition . The SG1 parthenogenetic females were again reared under short-day and low-temperature conditions (8 h light: 16 h dark; 15 °C) until larviposition. SG1 females produced exclusively sexual oviparous females during the first six days after the onset of larviposition, which were collected and reared in rectangular cases (100 mm × 65 mm × 28 mm) under the same day-length conditions (8 h light: 16 h dark; 15 °C).
Scanning electron microscopy (SEM)
To examine the detailed morphological features of hind legs in third, fourth, and fifth (adult) instars of female aphids, the surface structures were observed by SEM. Specimens for SEM observations were prepared as previously described . Briefly, aphid samples were fixed using a microwave oven, transferred into increasing concentrations of ethanol, and then transferred into hexamethyldisilazane. Subsequently, samples were transferred into t-butanol and freeze-dried using a freeze dryer (ES-2030; Hitachi, Tokyo, Japan), and legs were dissected from the bodies. Then, the legs were coated with gold ion using an ion sputter (E-1010; Hitachi, Tokyo, Japan). Detailed morphological characteristics of legs were then observed by SEM (JSM-5510LV; JEOL, Tokyo, Japan).
To determine inner morphological features of hind legs, female legs of third, fourth, and fifth instars were observed using paraffin sections. As described previously , aphid legs fixed in FAA fixative (formalin-acetic acid-alcohol fixative: formalin: acetic acid: ethanol = 6:1:16) were dehydrated in increasing concentrations of ethanol and embedded in paraplast (Sigma-Aldrich, St. Louis, MO, USA). Sections (5-μm thick) were processed using a microtome. Sections were stained with hematoxylin and eosin. Tissues were observed under a light microscope (BZ-9000E; Keyence, Osaka, Japan).
Transmission electron microscopy (TEM)
To determine inner ultrastructure of epidermal tissue from hind tibias of fourth and fifth instars of female aphids, TEM observations were performed. Specimens were prepared following the protocols described in . Aphid hind tibias were initially fixed in 2% glutaraldehyde and 2% paraformaldehyde in 0.2 M cacodylate sodium buffer (pH 7.4) at 4 °C for 5 h, postfixed in 2% osmium tetroxide in the same buffer for 2 h at 4 °C, and then dehydrated in increasing concentrations of acetone. Subsequently, the hind tibias were embedded in EPON 812 resin and polymerized. Ultra-thin sections were made using an ultramicrotome EM UC7 (Leica, Tokyo, Japan) with a diamond knife cryo 35-(wet) (DiATOME, Biel, Switzerland). The sections were then stained with 1% potassium permanganate solution for 2 min and 2% lead citrate for 3 min. The prepared sections were then observed by TEM (H-7500; Hitachi, Tokyo, Japan).
Identification of the genes in the A. pisum genome
List of enzyme genes involved in the monoterpenoid synthesis
Reference nucleotide sequence
Reference protein sequence
Mevalonate 5-diphosphate decarboxylase
Farnesyl diphospate synthetase
For real-time quantitative RT-PCR, total RNA was extracted by ISOGEN (Nippon Gene, Tokyo, Japan) with recombinant DNase1 (RNase-free) (TaKaRa, Shiga, Japan) from more than 40 hind tibias (20 aphid individuals) and three whole bodies of adult aphids. For biological replication, three replicated samples were prepared for each of the following four categories: (1) hind tibias of adult oviparous females, (2) hind tibias of adult parthenogenetic females, (3) whole bodies of adult oviparous females, and (4) whole bodies of adult parthenogenetic females. For each sample, extracted total RNA was reverse-transcribed with the High Capacity cDNA Reverse Transcription Kit based on the manufacturer’s instructions (Applied Biosystems, Foster City, CA, USA).
Primers for the target enzyme genes were designed using Primer Express software v3.0.1 (Applied Biosystems) (Additional file 1: Table S1). For AACT1 and AACT2, the primers were designed to cover the conserved region shared by the two paralogs, because the DNA sequences of the two genes are too similar to design primers specific to each paralog.
Quantitative RT-PCR (qRT-PCR) was performed using a SYBR Green I Chemistry System and Sequence Detection System ABI PRISM 7500 (Applied Biosystems, Foster City, CA, USA). To determine the endogenous control of constitutive expression, the suitability of different putative reference genes was evaluated among 4 candidate reference genes, glyceraldehyde-3-phosphate dehydrogenase (GAPDH), beta-actin, elongation factor 1 alpha (EF1a), and ribosomal protein L32 (rpL32), by three statistical software programs, geNorm , BestKeeper , and Normfinder . As the results, the expression level of GAPDH was shown to be the most stable so that it was used as the reference gene.
Data acquisition and analyses were performed by ABI Prism 7500 v2.0.1 (Applied Biosystems, Foster City, CA, USA). Baselines and Ct values (threshold cycle) were set automatically. The relative standard curve method was used for qRT-PCR quantification, as described in User Bulletin 2 for the ABI Prism 7700 Sequence Detection System (Applied Biosystems, Foster City, CA, USA). To evaluate the significance of expression differences among the categories, Student’s t-tests (p < 0.01) were performed.
Scanning electron microscopy on the hind tibia surface
By contrast, in oviparous females, numerous circular bulge structures called scent plaques were scattered on the hind tibia surface, especially in proximal and medial parts (Fig. 2j–l). Most scent plaques ranged from 7 to 10 μm in diameter. Most occurred separate from one another, although some were attached, forming “∞” shapes (Fig. 2k). On each scent plaque, many small pores, called epicuticular pores, were observed (Fig. 2n). The surface structures on hind femurs were similar to those on parthenogenetic females, where scent plaques were absent (Fig. 2m). In addition, scent plaques were absent on the surfaces of fore and mid tibias, which had smooth surfaces, as seen in parthenogenetic females (Fig. 2f and o).
Furthermore, hind tibias were also observed in nymphs. In the fourth instar (last nymphal instar), no structural differences were found between the two morphs; i.e., scent plaques were not found on any leg part in either morph (Fig. 2g–i, p–r). The tibia surfaces were smooth, which was also observed in parthenogenetic adult females. Also, scent plaques were not found on hind tibias in any other nymphal instars (data not shown).
Histological structures of hind tibial epidermis
To understand the associated developmental processes, nymphal hind tibias were also observed. In the third-instar nymphs of both morphs, epithelial cell shape in hind tibias was columnar, and a newly formed cuticle was observed between the outer cuticle and epithelial cell layer (Fig. 3b and f). The structural differences between oviparous and parthenogenetic females were first observed in fourth-instar nymphs. In future oviparous females, the epithelial cell layers were more developed than those of asexual females (Fig. 3c and g).
Epidermal cell ultrastructure
In the fourth-instar nymphs of oviparous females, the epithelial tissues of hind tibias did not possess secretory vesicles, although epicuticular pores were already formed on newly formed cuticle (Fig. 4e). In parthenogenetic female nymphs, no epicuticular pores were observed on newly formed cuticle (data not shown).
In whole bodies, AACT1&2, HMGS1, and MevPPD did not show significant expression differences, but MevPK, IPPI1, and FPPS showed significantly higher expression levels in oviparous females (approximately two times higher; Student’s t-test, P < 0.01) (Additional file 2: Figure S3). The expression levels of the other genes (AACT3–5, HMGS2, HMGR, MevK, MevPPD2, and IPPI2) were not detected, probably due to very low levels of expression.
The present study describes the internal structures of the scent plaques associated with epidermal gland cells in the pea aphid. Epidermal gland cells are generally classified into three types, as follows: (1) Class 1 epidermal gland cells are monocellular and contact the glandular cuticle; (2) Class 2 epidermal gland cells do not contact the glandular cuticle, and transport secretory materials into adjacent Class 1 cells; and (3) Class 3 epidermal gland cells are multicellular, and consist of secretory cells and duct cells [37, 38]. In Class 1 epidermal glands, the glandular cuticle is often thin to facilitate transport of secretory molecules [37, 39]. In addition, enlarged pore canals, epicuticular filaments, and epicuticular pores are often formed in the glandular cuticle of Class 1 cells to facilitate diffusion of large molecules or retain secretory materials [37, 38, 40–42]. Based on these criteria, scent plaques in the pea aphid are suggested to be composed of glandular cuticle, and the epithelial cells under the scent plaques are classified as Class 1 gland cells, suggesting that sex pheromones are synthesized in the gland cells under the scent plaques. Thus, in the hind tibias of adult oviparous females, the sex pheromones are first transported and contained in invaginated tubular structures, and are secreted through the glandular cuticle (i.e., scent plaques) (Fig. 4f).
In general, the epidermis of adult insects is flattened, as adult insects generally do not synthesize new cuticles [38, 43]. Hind, fore, and mid tibias of adult parthenogenetic females have general adult features but the hind tibias lack scent plaques, which indicates that the epithelial cells in parthenogenetic females are incapable of secreting sex pheromones.
Monoterpene biosynthesis is essential for plants to synthesize defense oil and resin . In animals, however, monoterpene biosynthesis was thought to be rare . In contrast, recent studies revealed that several beetles (I. pini, Dendroctonus jeffreyi, Phaedon cochleariae, and Gastrophysa viridula) used the mevalonate pathway to synthesize monoterpenes [21, 44–49]. In addition, the FPPS protein in the peach-potato aphids, Myzus persicae, was reported to convert the final products of mevalonate pathway to GPP [50, 51]. Therefore, the qRT-PCR results (Fig. 5) indicated that the downstream enzymes (MevPK, MevPPD, and IPPI) and FPPS were responsible for monoterpene biosynthesis in oviparous females (Figs. 2–4), which is consistent with the fact that only oviparous females possess Class 1 gland cells on hind tibias. These results indicate that oviparous females synthesize monoterpenoid sex pheromones by up-regulating gene expression of downstream enzyme genes in the Class 1 epidermal gland cells of hind tibias. In contrast, the expression levels of all paralogous genes of AACT, HMGR, and MevK were not detected from hind tibias. However, the expression of AACT were detected in whole bodies of both morphs (Additional file 2: Figure S3). Based on these results, we suggest that the upstream substrates are also synthesized through the mevalonate pathway somewhere in the body, and the products (precursors of terpenoids) are transported to the gland cells of hind tibias to facilitate the downstream cascades of terpenoid synthesis.
As for gene expression in whole bodies, the expression levels of mevalonate-pathway genes were relatively high even in parthenogenetic females. This suggests that the pathway is also important for parthenogenetic females. For example, it is known that the mevalonate pathway is also crucial for juvenile hormone (JH) biosynthesis in insects, which suggests that expression levels in whole bodies may be caused by JH biosynthesis in the corpora allata. Juvenile hormone is reported to play important roles in the production of parthenogenetic females , and some JH-related genes are down-regulated under short-day length, when sexual females are produced . Although these JH actions are only known to affect the next generations, JH should also be required for the postembryonic development (i.e., moultings) both in the parthenogenetic and sexual females, so that expressions of the mevalonate-pathway genes were detected in whole bodies.
Parthenogenesis is thought to have evolved in aphid life cycles about 200 million years ago [2, 54–57]. In addition, scent plaques are found in the most Aphididae species [9, 10, 58], and it was reported that two monoterpenoid components are used as sex pheromones in many aphid species [19, 59]. These suggest that hind tibias with scent plaques is an ancestral characteristic in Aphididae. Therefore, as the result of the evolution of parthenogenesis, the developmental mechanism that forms sex-pheromone glands is repressed in parthenogenetic females.
This study reveals that glandular tissues of scent plaque are differentiated from the fourth instar in sexual oviparous females, while parthenogenetic females lack the glandular cells. Furthermore, it was shown that the downstream steps of the mevalonate pathway are up-regulated in scent plaques on hind tibias of sexual females, although the upstream steps might occur somewhere in other body parts. In the aphid life cycle, oviparous sexual females only appear once a year, while parthenogenetic females occur throughout the rest of the annual life cycle [2–4]. Therefore, the adaptive switching mechanism that leads the development of sexual characters with functional metabolic pathways in response to environmental signals is suggested to have evolved, accompanying the evolution of cyclic parthenogenesis.
We are grateful to D. Watanabe for his technical advice. We also thank T. Funatsu and T. Terada for their assistance in aphid stock maintenance.
This work was supported by a Grant-in-Aid for Scientific Research (A) (No. 25251041) to TM from the Ministry of Education, Culture, Sports, Science and Technology of Japan.
Availability of data and materials
The nucleotide sequence data used in this study can be found in the DDBJ/EMBL/GenBank database. The accession numbers are listed in Table 1.
KM and TM designed the study project; KM, KO and TK carried out the experiments. KM and TM interpreted the obtained data and wrote the manuscript. All the authors read and approved the final manuscript.
Ethics approval and consent to participate
Laboratory-maintained insects were used in all the experiments. Ethical approval and consent to participate were not required for this work.
The authors declare that they have no competing interests.
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.
- Hebert PDN. Genotypic characteristics of cyclic parthenogens and their obligately asexual derivatives. In: Stearns SC, editor. The evolution of sex and its consequences. Experientia Supplementum, vol 55, vol. 55. Basel: Birkhäuser; 1987. p. 175–95.View ArticleGoogle Scholar
- Moran NA. The evolution of aphid life cycle. Annu Rev Entomol. 1992;37:321–48.View ArticleGoogle Scholar
- Simon JC, Rispe C, Sunnucks P. Ecology and evolution of sex in aphids. Trends Ecol Evol. 2002;17:34–9.View ArticleGoogle Scholar
- Miura T, Braendle C, Shingleton A, Sisk G, Kambhampati S, Stern DL. A comparison of parthenogenetic and sexual embryogenesis of the pea aphid Acyrthosiphon pisum (Hemiptera: Aphidoidea). J Exp Zool B: Mol Dev Evol. 2003;295:59–81.View ArticleGoogle Scholar
- Marcovitch S. Plant lice and light exposure. Science. 1923;58:537–8.View ArticlePubMedGoogle Scholar
- Blackman RL. Reproduction, cytogenetics and development. In: Minks AK, Harrewijn P, editors. Aphids: their biology, natural enemies and control. Amsterdam:Elsevier; 1987. p. 163–95.Google Scholar
- West-Eberhard MJ. Developmental plasticity and evolution. New York: Oxford University Press; 2003.Google Scholar
- Ogawa K, Miura T. Aphid polyphenisms: trans-generational developmental regulation through viviparity. Frontiers Physiol. 2014;5:1.View ArticleGoogle Scholar
- Petterson J. An aphid sex attractant. 1. Biological studies. Scandinavian Entomol. 1970;1:63–73.View ArticleGoogle Scholar
- Marsh D. Sex pheromone in the aphid Megoura viciae. Nature. 1972;238:31–2.View ArticleGoogle Scholar
- Marsh D. Responses of male aphids to the female sex pheromone in Megoura viciae Buckton. J Entomol (A). 1975;50:43–64.Google Scholar
- Hardie J, Peace L, Pickett JA, Smiley DWM, Stoper JR, Wadhams LJ. Sex pheromone stereochemistry and purity affect field catches of male aphids. J Chem Ecol. 1997;23:2547–54.View ArticleGoogle Scholar
- Dawson GW, Griffiths DC, Janes NF, Mudd A, Pickett JA, Wadhams LJ, Woodcocs CM. Identification of an aphid sex pheromone. Nature. 1987;325:614–6.View ArticleGoogle Scholar
- Dawson GW, Griffiths DC, Janes NF, Mudd A, Pickett JA, Wadhams L, Woodcocs CM. Aphid semiochemicals, a review, and recent advances on the sex pheromone. J Chem Ecol. 1990;16:3019–30.View ArticlePubMedGoogle Scholar
- Hardie J, Holyoak M, Nicholas J, Nottingham SF, Pickett JA, Wadhams LJ, Woodcock CM. Aphid sex pheromone components: age dependent release by females and species-specific male response. Chemoecol. 1990;1:63–8.View ArticleGoogle Scholar
- Hardie J, Pickett JA, Pow EM, Aphids SDWM. In: Hardie RJ, Minks AK, editors. Pheromones of non-lepidopteran insects associated with agricultural plants. Wallingford: CAB international; 1999. p. 250–77.Google Scholar
- Birkett MA, Picket JA. Aphid sex pheromones: from discovery to commercial production. Phytochemistory. 2003;62:651–6.View ArticleGoogle Scholar
- Stroyan HLG 1957. Introduction and subgenus Sappaphis sensu stricto. In: Stroyan HLG, editor. A revision of the British species of Sappaphis Matsumara. London: Ministry of Agriculture, Fisheries and Food; 1957. p. 24–27.Google Scholar
- Petterson J. An aphid sex attractant. 2. Histological, ethological and comparative studies. Scandinavian Entomol. 1971;2:81–93.Google Scholar
- Bellés X, Martín D, Piulachs MD. The mevalonate pathway and the synthesis of juvenile hormone in insects. Annu Rev Entomol. 2005;50:181–99.View ArticlePubMedGoogle Scholar
- Blomquist GJ, Figueroa-Teran R, Aw M, Song M, Gorzalski A, Abbott NL, Chang E, Tittiger C. Pheromone production in bark beetles. Insect Biochem Mol Biol. 2010;40:699–712.View ArticlePubMedGoogle Scholar
- Gilg AB, Bearfield JC, Tittiger C, Welch WH, Blomquist GJ. Isolation and functional expression of an animal geranyl diphosphate synthase and its role in bark beetle pheromone biosynthesis. Proc Natl Acad Sci U S A. 2005;102:9760–5.View ArticlePubMedPubMed CentralGoogle Scholar
- Goldstein JL, Brown MS. Regulation of the mevalonate pathway. Nature. 1990;343:425–30.View ArticlePubMedGoogle Scholar
- Cane DE. Isoprenoid biosynthesis overview. In: Cane DE, editor. Comprehensive natural products chemistry, vol. 2. Oxford:Pergamon; 1999. p. 1–13.Google Scholar
- Kuzuyama T. Mevalonate and nonmevalonate pathways for the biosynthesis of isoprene units. Biosci Biotechnol Biochem. 2002;66:1619–27.View ArticlePubMedGoogle Scholar
- Poulter CD, Rilling HC. Prenyltransferases and isomerase. In: Porter JW, Spurgeon SL, editors. Biosynthesis of isoprenoid compounds, vol. 1. New York: John Wiley & Sons; 1981. p. 161–224.Google Scholar
- Kanbe T, Akimoto S. Allelic and genotypic diversity in long-term asexual populations of the pea aphid, Acyrthosiphon pisum in comparison with sexual populations. Mol Ecol. 2009;18:801–16.View ArticlePubMedGoogle Scholar
- Ishikawa A, Ogawa K, Gotoh H, Walsh TK, Tagu D, Brisson JA, Rispe C, Jaubert-Possamai S, Kanbe T, Tsubota T, Shiotsuki T, Miura T. Juvenile hormone titre and related gene expression during the change of reproductive modes in the pea aphid. Insect Mol Biol. 2012;21:49–60.View ArticlePubMedGoogle Scholar
- The International Aphid Genomics Consortium. Genome sequence of the pea pahid Acyrthosiphon pisum. PLoS Biol. 2010;8:e1000313.View ArticlePubMed CentralGoogle Scholar
- Wilkinson TL. Ishikawa H 2000. Injection of essential amino acids substitutes for bacterial supply in aposymbiotic pea aphids (Acyrthosiphon pisum). Entomol Exp Appl. 2000;94:85–91.View ArticleGoogle Scholar
- Ogawa K, Ishikawa A, Kanbe T, Akimoto S, Miura T. Male-specific flight apparatus development in Acyrthosiphon pisum (Aphididae, Hemiptera, Insecta): comparison with female wing polyphenism. Zoomorphology. 2012;131:197–207.View ArticleGoogle Scholar
- Kaji T, Tsukagoshi A. Origin of the novel chemoreceptor Aesthetasc “Y” in Ostracoda: morphogenetical thresholds and evolutionary innovation. Evol Dev. 2008;10:228–40.View ArticlePubMedGoogle Scholar
- Kumar S, Tamura K, Nei M. MEGA: molecular evolutionary genetics analysis software for microcomputers. Comp Appl Biosci. 1994;10:189–91.PubMedGoogle Scholar
- Vandesompele J, De Preter K, Pattyn F, Poppe B, Van Roy N, De Paepe A, Speleman F. Accurate normalization of real-time quantitative RT-PCR data by geometric averaging of multiple internal control genes. Genome Biol. 2002;3:RESEARCH0034.View ArticlePubMedPubMed CentralGoogle Scholar
- Pfaffl MW, Tichopad A, Prgomet C, Neuvians TP. Determination of stable housekeeping genes, differentially regulated target genes and sample integrity: BestKeeper–excel-based tool using pair-wise correlations. Biotechnol Lett. 2004;26:509–15.View ArticlePubMedGoogle Scholar
- Andersen CL, Jensen JL, Ørntoft TF. Normalization of real-time quantitative reverse transcription-PCR data: a model-based variance estimation approach to identify genes suited for normalization, applied to bladder and Colon Cancer data sets. Cancer Res. 2004;64:5245–50.View ArticlePubMedGoogle Scholar
- Quennedey A. Insect epidermal gland cells: ultrastructure and morphogenesis. In: Harrisson FW, Locke M, editors. Microscopic anatomy of invertebrates, vol. 11A. New York: Wiley Liss; 1998. p. 177–207.Google Scholar
- Chapman RF. The insects: structure and function. 5th ed. London: Cambridge University Press; 2013.Google Scholar
- Quennedey A. Morphology and ultrastructure of termite defense glands. In: Hermann HR, editor. Defensive mechanisms in social insects. New York: Academic Press; 1984. p. 151–200.Google Scholar
- Locke M. Pore canals and related structures in insect cuticle. Journal Biophys Biochem Cytol. 1961;10:589–618.View ArticleGoogle Scholar
- Locke M. The structure and formation of the cuticulin layer in the epicuticle of an insect, Calpodes ethlius (Lepidoptera, Hesperiidae). J Morphol. 1966;118:461–94.View ArticlePubMedGoogle Scholar
- Filshie BK. Fine structure of the cuticle of insects and other arthropods. In: King RC, Akai H, editors. Insect ultrastructure, vol. 1. New York: Plenum Press; 1982. p. 281–312.View ArticleGoogle Scholar
- Snodgrass RE. Principles of insect morphology. Mc Graw-Hill: New York and London; 1935.Google Scholar
- Tittiger C, Barkawi LS, Bengoa CS, Blomquist GJ, Seybold SJ. Structure and juvenile hormone-mediated regulation of the HMG-CoA reductase gene from the Jeffrey pine beetle, Dendroctonus jeffreyi. Mol Cell Endocrinol. 2003;199:11–21.View ArticlePubMedGoogle Scholar
- Bearfield JC, Keeling CI, Young S, Blomquist GJ, Tittiger C. Isolation, endocrine regulation and mRNA distribution of the 3-hydroxy-3-methylglutaryl coenzyme a synthase (HMG-S) gene from the pine engraver, Ips pini (Coleoptera: Scolytidae). Insect Mol Biol. 2006;15:187–95.View ArticlePubMedGoogle Scholar
- Tillman JA, Lu F, Goodard LM, Donaldson Z, Dwinell SC, Tittiger C, Hall GM, Strer AJ, Blomquist GJ, Seybold SJ. Juvenile hormone regulates de novo isoprenoid aggregation pheromone biosynthesis in pine bark beetles, Ips spp. (Coleoptera: Scolytidae), through transcriptional control of HMG-CoA reductase. J Chem Ecol. 2004;30:2459–94.View ArticlePubMedGoogle Scholar
- Keeling CI, Blomquist GJ, Tittiger C. Coordinated gene expression for pheromone biosynthesis in the pine engraver beetle, Ips pini (Coleoptera: Scolytidae). Naturwiss. 2004;91:324–8.View ArticlePubMedGoogle Scholar
- Hall GM, Tittiger C, Blomquist GJ, Andrews G, Mastick G, Barkawi LS, Bengoa C, Seybold SJ. Male Jeffrey pine beetle, Dendroctonus jeffreyi, synthesizes the pheromone component frontalin in anterior midgut tissue. Insect Biochem Mol Biol. 2002;32:1525–32.View ArticlePubMedGoogle Scholar
- Burse A, Schmidt A, Frick S, Kuhn J, Gershenzon J, Boland W. Iridoid biosynthesis in Chrysomelina larvae: fat body produces early terpenoid precursors. Insect Biochem Mole Biol. 2007;37:255–65.View ArticleGoogle Scholar
- Lewis MJ, Prosser IM, Mohib A, Field LM. Cloning and characterization of a prenyltransferase from the aphid Myzus persicae with potential involvement in alarm pheromone biosynthesis. Insect Mol Biol. 2008;17:437–43.View ArticlePubMedGoogle Scholar
- Vandermoten S, Charloteaux B, Santini S, Sen SE, Béliveau C, Vandenbol M, Francis F, Brasseur R, Cusson M, Haubruge E. Characterization of a novel aphid prenyltransferase displaying dual geranyl/farnesyl diphosphate synthase activity. FEBS Lett. 2008;582:1928–34.View ArticlePubMedGoogle Scholar
- Corbitt TS, Hardie J. Juvenile hormone effects on polymorphism in the pea aphid, Acyrthosiphon pisum. Entomol Exp Appl. 1985;38:131–5.View ArticleGoogle Scholar
- Le Trionnaire G, Jaubert-Possamai S, Bonhomme J, Gauthier JP, Guernec G, Le Cam A, Legeai F, Monfort J, Tagu D. Transcriptomic profiling of the reproductive mode switch in the pea aphid in response to natural autumnal photoperiod. J Insect Physiol. 2012;58:1517–24.View ArticlePubMedGoogle Scholar
- Heie OE. Palaeontology and phylogeny. In: Minks AK, Harrewijn P, editors. Aphids: their biology, natural enemies and control, vol. 2A. Amsterdam: Elsevier; 1987. p. 367–91.Google Scholar
- Hales DF, Tomiuk J, Wöhrmann K, Sunnucks P. Evolutionary and genetic aspects of aphid biology: a review. Euro J Entomol. 1997;94:1–55.Google Scholar
- Dixon AFG. Aphid ecology. 2nd ed. London: Chapman & Hall; 1998.Google Scholar
- Davis GK. Cyclical parthenogenesis and viviparity in aphids as evolutionary novelties. J Exp Zool B: Mol Dev Evol. 2012;318:448–59.View ArticleGoogle Scholar
- Harrington R. A comparison of the external morphology of ‘scent plaques’ on the hind tibiae of oviparous aphids (Homoptera: Aphididae). Sys Entomol. 1985;10:135–44.View ArticleGoogle Scholar
- Brickett MA, Pickett JA. Aphid sex pheromones: from discovery to commercial duction. Phytochemistry. 2003;62:651–6.View ArticleGoogle Scholar