RESEARCH ARTICLE
Structure and Development of Male Pheromone Gland of Longicorn Beetles and its Phylogenetic Relationships Within the Tribe Clytini KEITA HOSHINO*, SATOSHI NAKABA, HIROKI INOUE, AND KIKUO IWABUCHI Faculty of Agriculture, Tokyo University of Agriculture and Technology, Fuchu, Tokyo, Japan
ABSTRACT
J. Exp. Zool. (Mol. Dev. Evol.) 324B:68–76, 2015
The male sex pheromone of the longicorn beetle, Xylotrechus pyrrhoderus pyrrhoderus Bates (Cerambycidae: Tribe Clytini) plays an important role in attracting females. This pheromone is produced by the pheromone gland located in the prothorax. However, the detailed structure and underlying developmental process of this gland are still unknown. We investigated the gland structure by using histological analysis and confirmed that the gland consists of the following parts: gland cell mass, a unique spherical space in the cuticle layer, and ductules connecting the gland cells with the spherical space and conducting canals to the outer opening. The gland structure first appeared male-specific in the late pupal stage, during which the epidermal cells began depositing the exocuticle; the development of the gland was completed after adult emergence. Furthermore, we verified the structural equivalents of the X. p. pyrrhoderus male pheromone gland in 11 species of 2 tribes, Clytini and Anaglyptini. The glands of these insects could be classified into four types on the basis of the absence or presence of the spherical space and the division of the gland cell mass layer. Most noteworthy, all the species with the spherical space and division-type gland were restricted to the Xylotrechus clade, as inferred from the molecular phylogenetic analysis. These results suggest that Clytini and Anaglyptini species share a fundamental process of male pheromone gland development, and that the Japanese Xylotrechus species might have established their current status by developing distinct structural features in the male pheromone gland. J. Exp. Zool. (Mol. Dev. Evol.) 324B:68–76, 2015. © 2014 Wiley Periodicals, Inc. How to cite this article: Hoshino K, Nakaba S, Inoue H, Iwabuchi K. 2015. Structure and development of male pheromone gland of longicorn beetles and its phylogenetic relationships within the tribe Clytini. J. Exp. Zool. (Mol. Dev. Evol.) 324B:68–76.
Longicorn beetles are a principal group in the order Coleoptera, which consists of diverse species of phytophagous borers, including many important arboreal pests. In these beetles, chemical communication systems are utilized during mating similar to those found in most other insect species. Since earlier studies have reported the presence of pheromones in longicorn beetles (Galford, '77), pheromone-mediated communication in cerambycid beetles has been investigated. Female volatile pheromones have been identified in the species belonging to subfamilies of Prioninae, Lepturinae, (Rodstein et al., 2011; Ray
Grant sponsor: Japan Society for the Promotion of Science (JSPS); grant number: 25870210. Conflict of interest: None. Correspondence to: Keita Hoshino, Faculty of Agriculture, Tokyo University of Agriculture and Technology, Fuchu, Tokyo 183-8509, Japan. E-mail: [email protected] Received 26 September 2014; Accepted 18 October 2014 DOI: 10.1002/jez.b.22606 Published online in Wiley Online Library (wileyonlinelibrary.com). .
© 2014 WILEY PERIODICALS, INC.
STRUCTURE AND DEVELOPMENT OF MALE PHEROMONE GLAND IN LONGICORN BEETLES et al., 2011, 2012a, 2012b), as well as Vesperinae (uncertain affiliation with the family Cerambycidae; Leal et al., '94). Thus far, male-produced sex or aggregation pheromones have been identified in 18 species of the subfamilies Cerambycinae, Lamiinae, and Spondylidinae (listed in Lacey et al., 2004; Silk et al., 2007; Fonseca et al., 2010; Teale et al., 2011; Fierke et al., 2012; Hanks and Millar, 2013; Pajares et al., 2013). Maleproduced pheromones have been found especially in the subfamily Cerambycinae, and chemical and ecological studies have been extensively performed in the tribe Clytini within this subfamily (Iwabuchi, '99). These pheromones are emitted from the surface of the prothorax, as indicated by the chemical analysis of individual body parts (Iwabuchi, '86) and the morphological analysis of male-specific pores in the subfamily Cerambycinae (Ray et al., 2006). Pheromone glands are located beneath the pores on the prothorax in 5 species of the subfamily Cerambycinae (Iwabuchi, '86; Nakamuta et al., '94; Noldt et al., '95; Hanks et al., 2007; Lacey et al., 2007). These structural analyses of pheromone glands, however, have not been performed in detail, and more thorough analysis will enhance the understanding of these glands. Pheromone glands have a specialized glandular function; generally, they consist of gland cells with conducting canals in the cuticle layer (Chapman, '98; Neville, '75). Female pheromone glands have been studied in detail, both morphologically and developmentally, mainly in lepidopteran insects (Percy-Cunningham and MacDonald, '87). In contrast, studies on male pheromone glands are rare, but have been reported for mecopteran, neuropteran, dipteran, and coleopteran insects (Crossley and Waterhouse, '69; Wattebled et al., '78; Nardi et al., '96; Spiegel et al., 2002, 2004). Developmental studies on male pheromone glands, in particular, will provide valuable information for understanding how male-specific developmental processes have evolved; however, no such studies have yet been conducted in male longicorn beetles. Furthermore, male pheromones of tribe Clytini are indicated to be similar at the chemical structure level (Ray et al., 2012b; Hanks and Millar, 2013); thus, a comparative analysis of the morphology of the gland structures within the tribe will aid in determining the phylogenetic relationships in the male pheromone gland evolution. Molecular-based phylogenetic analysis might enhance the accuracy of the comparative study of the glands, since phylogenetic relationships among genera of tribe Clytini are not well understood (Ohbayashi and Niisato, 2007) and have never been confirmed by molecular data. In this study, we investigated the detailed morphological and developmental processes of the male pheromone gland of Xylotrechus pyrrhoderus pyrrhoderus Bates, a clytine species that is known to have male sex pheromone (Iwabuchi, '82). We also conducted a comparative study of the male pheromone glands among another 11 species of tribe Clytini and 1 species of Anaglyptini (closely related tribe within the subfamily Ceram-
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bycinae; Ohbayashi and Niisato, 2007). Finally, on the basis of the result of molecular-based phylogenetic analysis, we discuss the development and evolution of male pheromone glands in tribe Clytini.
MATERIALS AND METHODS Insects X. p. pyrrhoderus was obtained by collecting pruned grape vine shoots with wood-boring larvae in March. The grape vine shoots were placed upright in outdoor insect cages (120 120 200 cm in height). The development of the larvae was periodically monitored. In late June, the final instar larvae (weight, >170 mg) were collected from the shoots and maintained individually in 35-mm plastic dishes. A good environment for the larvae was maintained by placing a small cutting of the grape vine shoot and wet filter paper within each dish. Dishes were maintained in the laboratory at 25 °C with a light:dark (L:D) photoperiod of 16:8. Pupae were sexed according to morphological characteristics in the apical region of the abdomen, as described by Duffy, ('53). The pupal periods and days after emergence were counted and recorded daily. Thirteen species belonging to the tribes Clytini and Anaglyptini in the subfamily Cerambycinae (Clytini: X. p. pyrrhoderus; Xylotrechus chinensis chinensis (Chevrolat); Xylotrechus rufilius Bates; Xylotrechus emaciatus Bates; Xylotrechus cuneipennis (Kraatz); Clytus melaenus Bates; Cyrtoclytus caproides caproides (Bates); Plagionotus christophi (Kraatz); Demonax notabilis notabilis (Pascoe); Rhaphuma xenisca (Bates); Rhabdoclytus acutivittis inscriptus (Bates); and Chlorophorus quinquefasciatus (Castelnau et Gory); Anaglyptini: Paraclytus excultus Bates) were used for histological analysis of the pheromone glands. Two species of different subfamilies (Lepturinae: Leptura ochraceofasciata ochraceofasciata (Motschulsky) and Necydalinae: Necydalis solida Bates) were also used for histological analysis. All specimens used for this analysis were collected and transferred alive to the laboratory. Sex was determined for all specimens on the basis of genital morphology and other accompanying sexual characters, including the shapes of the antenna and abdomen. Collection localities and numbers of all specimens used for the histological analysis were listed in supplementary materials Table S1. For the phylogenetic study, besides the above-mentioned 13 clytine and anaglyptine species, 13 species (Clytini: Xylotrechus villioni (Villard); Xylotrechus atronotatus angulithorax Gressitt; Xylotrechus grayli grayii (White); Perissus kiusiuensis kiusiuensis Ohbayashi; Clytus auripilis Bates; Brachyclytus singularis Kraatz; Epiclytus yokoyamai (Kano); Demonax transilis Bates; Chlorophorus japonicus (Chevrolat); Chlorophorus muscosus (Bates), Anaglyptini: Anaglyptus niponensis Bates; Anaglyptus matsushitai Hayashi, Callidiini: Phymatodes maakii Kraatz) and 3 additional species from a different J. Exp. Zool. (Mol. Dev. Evol.)
70 subfamilies (Disteniinae: Distenia gracilis gracilis (Blessig), Prioninae: Prionus insularis insularis Motschulsky, Lamiinae: Dolichoprosopus yokoyamai (Gressitt)) were used. In all, samples from 29 species were stored at 20°C with or without 100% ethanol until use. The data of all specimens used for the phylogenetic analysis were listed in Table S2. Preparation of Histological Samples A razor and forceps were used to obtain a prothoracic sample from each living specimen; the sample was trimmed within half a minute, and the left region was used. The prothoracic piece was immediately placed in a 2-mL plastic tube containing 500 mL of fixative solution (3% paraformaldehyde (WAKO, Osaka, Japan), 2% glutaraldehyde (TAAB, Berks, UK) in 0.1 M phosphate buffer (PB), pH 7.0) and was refrigerated for 2 hr at 4 °C. After fixation, the sample was washed by replacing the buffer solution with fresh one twice. The sample was then transferred to the post-fixative solution (2% osmium tetroxide (WAKO) in 0.1 M PB, pH 7.0) and refrigerated for another 2 hr at 4 °C. After post-fixation, the sample was washed with the buffer and dehydrated with increasing concentrations of diluted ice-cold ethanol (50, 60, 70, 80, 90, and 95%). Samples were incubated in each ethanol solution for 15 min at room temperature, except for 30 min in 95% ethanol. The samples were further dehydrated in absolute ethanol containing molecular sieves (3 A; Sigma–Aldrich, St Louis, MO, USA). After the dehydration process was repeated with absolute ethanol three times, the ethanol was replaced with a mixture containing 50% ethanol and 50% 1-butoxy 1, 3epoxypropan (QY-1; EM Japan, Tokyo). The mixture was gradually replaced with QY-1 by exchanging the medium four times and allowing the sample to stabilize for 10 min after each exchange. After the solution was replaced with 100% QY-1, QY-1 was gradually replaced with Epon resin (solution A: Epon 812 (TAAB) 580 mL and dodecanyl succinic anhydride (DDSA) (TAAB) 925 mL; solution B: Epon 812 (1,850 mg) and methyl nadic anhydride (MNA) (TAAB) 1670 mg; A:B ¼ 3:7) through serial exchanges with solutions containing an increasing proportion of Epon resin. Samples were adapted for more than 30 min after each exchange, and the final exchange solution contained only Epon resin. After the samples were placed in the resin overnight, the processed specimens were then transferred to a mold filled with the prepared resin to which DMP-30 (TAAB) was added. The specimens were then heated in an oven according to Luft ('61) and completely embedded for the next sectioning step. Thin Sectioning and Staining The embedded specimens were roughly cut out of the surplus resin surrounding the tissue and trimmed into a right trapezoidal shape. The direction of the specimen was adjusted by cutting from the longer base of the trapezoid. Approximately 1-mm sections were cut from the specimen by using an ultramicrotome J. Exp. Zool. (Mol. Dev. Evol.)
HOSHINO ET AL. (Ultracut N; Reihert, Vienna, Austria). For thin sectioning, glass knives were placed in a container filled with water, and knives were exchanged every 5–10 cuts. Each section was checked for quality by examining its brightness, picked up using a loop of a platinum–palladium wire attached to a wooden shaft, and placed in a pool of water. The sections were transferred to a staining solution of 10% azure B, 1% safranin, or 1% toluidine blue and stained for 30 min at 60 °C, 30 min at room temperature, or 5 min at room temperature, respectively. The sections were then washed by transferring to water pools 3 times and finally transferred onto glass slides. The glass slides were dried in an oven at 60 °C. Observation and Comparative Study of the Male Pheromone Gland Structure Detailed morphological analysis of the male pheromone gland was conducted for X. p. pyrrhoderus. Approximately 200 serial sections were cut and examined. By integration of sharp and well-stained microscopic images, each structure for male pheromone gland was defined and represented schematically, followed by referring to the general criteria (Neville, '75; Chapman, '98). The development of pheromone glands was elucidated by examining the various developmental stages of male X. p. pyrrhoderus at 1–2 day intervals in the period before and after adult emergence. Approximately 50 sections per stage were observed, and developmental changes were detected. For taxonomic comparison of male pheromone glands across different species, the presence of structural equivalent of male pheromone glands of X. p. pyrrhoderus was determined. Sections were sampled at every 50-mm scraping and, when the gland was detected, the structural features, especially the presence of spherical space and division or layered type of gland cell mass, were examined. For females, 8 species (X. p. pyrrhoderus; X. c. chinensis; X. emaciatus; X. cuneipennis; C. melaenus; C. c. caproides; P. christophi; and R. xenisca) were used for histological analysis. All specimens were observed under a microscope (DM LB; Leica) with an optical mode at a magnification of 200. Nucleotide Sequencing DNA was extracted from insect forelegs by using a DNeasy blood and tissue kit (QIAGEN, Hilden, Germany). Samples were extracted from frozen specimens or specimens that had been submerged in 100% ethanol. The DNA extraction was conducted according to the manufacturer's instructions, and the eluted DNA solution was washed and concentrated using the ethanol precipitation method. The concentration of the samples was measured, and the ratio of absorbance at 260 and 280 nm was determined using Nanodrop (Thermo Fisher Scientific, Wilmington, DE, USA). Samples were stored at 80°C until use. The three partial regions of 2 mitochondrial genes encoding NADH dehydrogenase subunit 5 (ND5) and cytochrome oxidase subunit 1 (CO1) and a nuclear 28 S ribosomal DNA (rDNA) were
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amplified using polymerase chain reactions (PCRs) by using Quick Taq HS Dyemix (Toyobo Co. Ltd., Tokyo, Japan). All primers used for the amplification of each target were newly designed from alignment of related genes from GenBank: CerND5-FW (50 -GGAGTTTATTTAYTRATTCG-30 ) and Cer-ND5-RV (50 -CCTAAAGTAGAAACAAYWGG-30 ) for mitochondrial ND5, Cer-CO1-FW (50 -CCCGGATTTGGRATAATYTC-30 ) and Cer-CO1RV (50 -TCAGAATATCTRTGTTCDGC-30 ) for mitochondrial CO1, and Cer-28S-rRNA-FW (50 -CTAGGATTCCCTCAGTAGCT-30 ) and Cer-28S-rRNA-RV (50 -GCTCWTCCCTKTTCGCTCGCA-30 ) for nuclear 28S-rDNA. The PCRs were performed according to the manufacturer's instructions. PCR conditions were as follows: 2 min at 94 °C, followed by 35 cycles of 30 sec at 94 °C, 30 sec at 50 °C, and 1 min at 68 °C. The amplicons were purified using agarose gel electrophoresis and then extracted from the gels using a QIAEX gel extraction kit 2 (QIAGEN). The purified DNA fragments were directly sequenced using the ABI Prism Big Dye Terminator cycle sequence kit version 3.1 (Applied Biosystems (ABI), Foster City, CA, USA) and the ABI Prism 3130 genetic analyzer. The nucleotide sequences of the amplified fragments were determined using a cycle sequencing method. The purified PCR amplicons were sequenced using a dye terminator system (ABI). The sequence data obtained were checked for signal strength and electropherogram quality by using an analysis tool (ABI) and processed for phylogenetic analysis. Phylogenetic Analysis The phylogenetic relationships among the 22 clytine and 3 anaglyptine species were analyzed by comparing the nucleotide sequences encoding mitochondrial CO1 and ND5 and nuclear rDNA. The sequence data have been submitted to GenBank, and the accession numbers are listed in Table S2. Two partial gene sequences of CO1 and ND5 were concatenated and used as mitochondrial sequences. The mitochondrial and nuclear sequences were aligned using the ClustalX program (Thompson et al., '94). The best-fitting nucleotide substitution model was determined as GTR þ G þ I model by the program MrModeltest (Posada and Crandall, 2001) executed in PAUP* 4.0b10 (Swofford, 2003). The phylogenetic dendrogram was inferred using Bayesian method by using MrBayes version 3.2 (Ronquist and Huelsenbeck, 2003).
RESULTS Morphological Observation of the Male Pheromone Glands The pheromone glands of X. p. pyrrhoderus were observed only in male samples (Fig. 1) and are represented schematically and terminologically in Figure 2. In one field of microscopic view, a single gland unit consisted of approximately 20 gland cells that were associated with the “end apparatus,” which is the terminus of an extended ductule within the gland cell cavity (Figs. 1B and 2; Chapman, '98). Immediately exterior to the gland cells, a
FIGURE 1. Micrographs of the transverse sections of adult X. p. pyrrhoderus prothorax 2 weeks after emergence. The densely stained, wavy horizontal line is the cuticle layer. The upper side of the cuticle layer is the outer space and, below this layer, is the hemocoel. (A–C): male; D: female. (A) The ovoid, multicellular tissue visible under the cuticle layer is a pheromone gland (stained blue with azure B). (B) An axial section of gland cells stained with safranin. The gland cells have a cavity (arrow, surrounded by whity region) from which the ductules (arrowhead) extend toward the spherical space. (C) A partial image of the spherical space in the cuticle layer from a section stained with toluidine blue. A bundle of gathered ductules from gland cells extended to the spherical space. (D) None of the pheromone gland structures, including gland cell mass and ductules in the cuticle layer, are found in female-derived sections stained with safranin. Bar indicates 50 mm.
cuticle consisting of two distinct layers (exocuticle and endocuticle) was visible under light microscopy (Fig. 1A, D). The exocuticle was the outer layer and had a wavy appearance. The endocuticle consisted of 7–9 layers with a striped pattern, J. Exp. Zool. (Mol. Dev. Evol.)
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FIGURE 2. Schematic representation and terminology of X. p. pyrrhoderus male pheromone gland structure. The cuticle is the uppermost part in the figure, and the hemocoel is beneath the cuticle. The colors represent exocuticle (red), endocuticle (blue), and gland cells (yellow). A spherical space is formed by exocuticle evagination. Gland cell mass consists of many individual gland cells of which some representatives, i.e., three axial sectional and one transectional images, are described. Abbreviations represented in this figure are used for Figure 3 and supplementary materials Fig. S1 and S2.
corresponding to the structure termed “balken” and known to be a distinctive feature of coleopteran insects (Wigglesworth, '72; De Camp and Greven, 2010). Observation of serial sections showed spherical structures in the endocuticle layer. This structure was merely designated as a “spherical space,” since this was previously indicated to be a reservoir (Iwabuchi, '86) of unknown function. From the gland cell mass located under each spherical space, a bundle of ductules extended to the inner part of the spherical space (Figs. 1C and 2). Observation of serial sections showed that only the superior region of the spherical space was fragile. These findings suggested the presence of some conducting canals to the outer openings from the spherical space, as indicated by previous scanning electron microscopic (SEM) observations (Iwabuchi, '86; Ray et al., 2006), but they are probably not visible because their size is less than 1 mm. Developmental Changes During the Formation of the Male Pheromone Glands The chronological process of male pheromone gland formation is represented schematically in comparison with that in females in Figure 3. At 10 days after pupation (4 days before adult emergence), spherical spaces appeared in the epidermal cell layer, during which the exocuticle layer was deposited (Supplementary materials Fig. S1). The spherical spaces were formed by local invaginations of the exocuticle into the epidermal cell layer, which was located under the depressions of the cuticular surface (Fig. 3). After the spherical space formed, a bundle of ductules and gland cells appeared beneath each spherical space (Fig. 3 and S1). J. Exp. Zool. (Mol. Dev. Evol.)
After adult emergence, there was no change in the structure of the gland cells, except for some increase in size. On the other hand, the endocuticle grew at a rate of one layer per day, and epidermal cells gradually degenerated. These changes ceased within 7–9 days after adult emergence. Pheromone glands were observed only in males, but cuticle growth and dermal gland formation were detected in both the sexes and followed the same developmental course. Taxonomic Comparisons of Male Pheromone Glands Across 11 Longicorn Species Of the 12 species, 11 had typical male pheromone glands that were similar to those of X. p. pyrrhoderus (Fig. S2). On the other hand, no pheromone glands were found in either sex in X. cuneipennis (data not shown) and two more distantly related species of subfamilies Lepturinae and Necydalinae (Fig. S2). In all the species with pheromone glands, the gland cells exhibited some common structural features such as end apparatuses and filamentous ductules. However, two obvious differences were detected, and we classified the glands on the basis of these differences. The primary classification was made on the basis of the absence or presence of a spherical space (type spherical space (SS)-positive: “ þ SS” and SS-negative: “ SS”), and the subdivision on the basis of the division of the gland cell mass layer with the basal lamina (type “division” and “layer”; see Fig. 4). The gland types of each species were as follows: þSSdivision (X. p. pyrrhoderus, X. c. chinensis, X. emaciatus, X. rufilius, and C. melaenus), þSS-layer (R. xenisca, P. christophi,
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FIGURE 3. Schematic representation of the transverse sections of male and female X. p. pyrrhoderus at each developmental stage, illustrating the development of the male pheromone gland. The numbers represent the number of days before ( ) or after (þ) adult emergence (AE). Abbreviations are as in Figure 2. Colors are as in Figure 2; grey and brown represent epidermal cell layer and pupal skin, respectively. Deposited exocuticle evaginated to form the spherical space, beneath which the swollen epidermal cells differentiated into ductules and gland cells (yellow region) in the males. Endocuticle growth and dermal gland formation occurred in both the sexes after AE. ps, pupal skin; ec, epidermal cells.
and C. c. caproides), SS-division (C. quinquefasciatus), and – SS-layer (P. excultus and R. acutivittis inscriptus). Molecular Phylogeny and its Relation to the Morphology of the Pheromone Glands The sequence alignment data were analyzed using Bayesian methods, and the obtained consensus trees for mitochondrial
gene are shown in Figure 5. Xylotrechus species formed the clade with those of the near-related genera Clytus and Perissus (Ohbayashi and Niisato, 2007), and Cyrtoclytus, including Brachyclytus species, formed the Xylotrechus clade. All other Clytini species were included in the sister group of Xylotrechus clade, whereas Anaglyptini and Callidiini were positioned between Xylotrechus clade and the outgroups. Species with
FIGURE 4. Classification of male pheromone glands detected in tribes Clytini and Anaglyptini, according to two structural characteristics: spherical space (SS) and gland cell mass layer. Types “ þ SS” and “ SS” indicate the presence and absence of spherical space, respectively. Types “division” and “layer” indicate divided gland cell mass layer by basal lamina and non-divided gland cell mass layer, respectively. The characteristics are represented in combinations as types þSS-division, þSS-layer, –SS-division, and SS-layer. Schematic diagrams represent the outlines of the gland types on the basis of Figure 2, and individual gland cells of the divided or only layered gland cell mass are not shown.
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FIGURE 5. Dendrograms showing the relationships of 25 longicorn beetle species in the tribes Clytini and Anaglyptini (subfamily Cerambycinae), as deduced from the molecular data. The nucleotide sequences of partial mitochondrion genes CO1 and ND5 were used for Bayesian analysis. All branches are labeled with Bayesian posterior probabilities. The phylogenetic areas of tribe Clytini and Xylotrechus clade are shaded. Each species' pheromone gland types (refer to Fig. 4) are mentioned after the species' names.
spherical space-positive glands were intensively distributed in the Xylotrechus clade, which was also closely linked with the subtype of the division, whereas the remaining types were scattered across the remaining taxonomic groups (Fig. 5). In contrast, the nuclear gene tree showed no significant resolution because of high gene conservation (data not shown).
DISCUSSION The principal male pheromone gland structure consists of gland cells and cuticular ducts that are similar to those found in many other typical pheromone glands (Percy-Cunningham and MacDonald, '87). The gland cells of X. p. pyrrhoderus were similar to those of the pheromone glands of coleopteran species (Nardi et al., '96) and the male-specific gland of the phlebotomine fly (Spiegel et al., 2002, 2004). However, the pheromone glands of X. p. pyrrhoderus differed from those of the abovementioned species in that (1) the ductules from each gland cell gathered to form a bundle of ductules that penetrated the endocuticle and reached the spherical space, and (2) the gland cells were clustered together. Such a bundle of ducts has also been observed in two North American clytine species, Xylotrechus nauticus and Neoclytus acuminatus acuminatus (Hanks et al., 2007; Lacey et al., 2007), but they lack the spherical spaces in the endocuticle J. Exp. Zool. (Mol. Dev. Evol.)
layer. Consequently, to our knowledge, this study confirmed the detailed structures of the pheromone glands for the first time (Fig. 2) and revised and renewed the previous representation by Iwabuchi ('86), especially, with regard to the presence of a spherical space in the endocuticle layer but made of exocuticle, and that the ductules extended from the end apparatus that showed characteristics of class-3 gland cells defined by Noirot and Quennedey, ('74). Initial gland formation was observed as the appearance of spherical spaces in the epidermal cell layer. Thereafter, the spherical space was strongly linked with the differentiation of the male pheromone gland; that is, gland cells and ductules arose in the swollen regions of the epidermis by the formation of the spherical space (Fig. 4). In the swollen region, imaginal epidermal cells differentiated into the male pheromone gland, probably because of the initiation of mitotic division that occurred along the vertical and oblique axes, as suggested by Corbet and Laifook ('77) and Percy-Cunningham and MacDonald ('87). There were very few pheromone gland openings on the surface compared to the number of gland cells, indicating that the spherical spaces served as the connecting point between ductules and conducting canals. Therefore, the spherical spaces might allow large-scale pheromone production by increasing the
STRUCTURE AND DEVELOPMENT OF MALE PHEROMONE GLAND IN LONGICORN BEETLES number of pheromone glands per opening pore. The dermal glands also generally consist of plural cells and are classified as class-3 gland cells; during their morphogenetic process, the conducting canal and gland cells are connected via a saccule for product storage and chemical reaction (Lai-Fook, '70). Additionally, dermal glands were common to both the sexes, and SEM observation indicated that the pores of male dermal glands and the pheromone glands within a single depression differed in number but were similar in size and other properties (Iwabuchi, '86). Therefore, the canals and pores might be formed through the same process; however, after formation, the ductules connect to the canals via saccules in the dermal glands and spherical spaces in the pheromone glands. Such similarities suggest that the spherical spaces might induce the differentiation of the male pheromone glands and might function as receptacles for the final conversion and storage of the male sex pheromones. Pheromone glands appeared before the formation of dermal glands, suggesting that male pheromone glands might be derived from a heterochronical shift of the universal gland developmental program. The structure of pheromone glands of Hylotrupes bajulus belonging to the tribe Hylotrupini and subfamily Cerambycinae is similar to that of X. p. pyrrhoderus (Noldt et al., '95). This is also reminiscent of the slight developmental changes reflected in the phylogeny. However, further investigations at the ultrastructural level are needed to clarify the developmental and phylogenetic similarities. Male pheromone glands of the 11 clytine species and 1 anaglyptine species were structurally similar to those of X. p. pyrrhoderus. The pheromone glands of these species could be classified into four types (see Fig. 4), but the other structural features were common, suggesting that the glands had diverged from a prototype, and this might explain the similarity of the chemical motif found in male pheromones of tribe Clytini (Ray et al., 2012b; Hanks and Millar, 2013). By using molecular-based phylogenetic analysis, we determined the accurate phylogenetic relationships among the genera of tribe Clytini and showed the distribution patterns of the gland types in the tribe. All the species with þ SS-division-type gland were included in the robust Xylotrechus clade, unlike the distribution of the other type species that were extended to the distant anaglyptine species clade. The Xylotrechus clade included some species with a remarkably narrow range of host plant species and living conditions (X. p. pyrrhoderus, X. c. chinensis, X. villioni, etc.), and the majority of the species of this clade preferred freshly fallen trees (Ohbayashi and Niisato, 2007; Kojima and Nakamura, 2011). However, P. christophi that have strict host preference for freshly fallen trees and showed the þSS but layer-type gland were not included in the Xylotrechus clade. In contrast, X. cuneipennis was included in the Xylotrechus clade, but it has no glands in either sex, possibly because of the secondary loss that seems to be associated with its preference for decayed trees unlike the Japanese Xylotrechus. Therefore, the Japanese Xylotrechus clade species
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might have evolved the spherical space and division-type glands in association with some specific ecological demands. Taken together, our findings suggest that Clytini and Anaglyptini species share a fundamental process of male pheromone gland development, and that Xylotrechus clade species might have established their current status by developing distinct structural features of the gland in agrrement with the phylogenetic relationships within the tribe Clytini. Recently, Ray et al. (2012b) reported that the parsimonious chemical motif was extended to subfamily Prioninae and that the conversion of the sexes that produce pheromones could occur. Accordingly, further studies over an extended range of longicorn species are needed to reveal the earlier evolutionary processes of male pheromones in longicorn beetles.
ACKNOWLEDGMENTS The authors thank Mr. Nazumi Mandokoro for providing the specimen of X. villioni larva and Miss Ami Ogura for her help with the field collection of X. p. pyrrhoderus larvae. We are also thankful to Prof. Ryo Funada and the members of the Laboratory of Morphogenesis of Plant Resources of Tokyo University of Agriculture and Technology for facilitating this research. We also thank Prof. Kazuhiko Sakurai of Seijo University for his field research. This work was partially supported by Japan Society for the Promotion of Science (JSPS) KAKENHI Grant Number 25870210 to KH.
LITERATURE CITED Chapman RF. 1998. The insects. 4th edition. New York: Cambridge University Press. Corbet SA, Lai-Fook J. 1977. The hairpencils of the flour moth, Ephestia kushucella. J Zool 181:377–394. Crossley ACS, Waterhouse DF. 1969. The ultrastructure of a pheromone-secreting gland in the male scorpion fly Harpobittacus australis (Bittacidae, Mecoptera). Tissue Cell 1:525–554. De Kamp T, Greven H. 2010. On the architecture of beetle elytra. Entomologie heute 22:191–204. Duffy EAJ. 1953. A monograph of the immature stages of British and imported timber beetles (Cerambycidae). London: The British Museum (Natural History). Fierke MK, Skabeikis DD, Millar JG, et al. 2012. Identification of a maleproduced aggregation pheromone for Monochamus scutellatus scutellatus and an attractant for the congener Monochamus notatus (Coleoptera: Cerambycidae). J Econ Entomol 105:2029–2034. Fonseca MG, Vidal DM, Zarbin PHG. 2010. Male-produced sex pheromone of the cerambycid beetle Hedypathes betulinus: chemical identification and biological activity. J Chem Ecol 36:1132–1139. Galford JR. 1977. Evidence for a pheromone in the locust borer. USDA Forest Service Research Note NE-240 1–3. Hanks LM, Millar JG. 2013. Field bioassays of cerambycid pheromones reveal widespread parsimony of pheromone structures, enhancement by host plant volatiles, and antagonism by components from heterospecifics. Chemoecol 23:21–44. J. Exp. Zool. (Mol. Dev. Evol.)
76 Hanks LM, Millar JG, Moreira JA, et al. 2007. Using generic pheromone lures to expedite identification of aggregation pheromones for the cerambycid beetles Xylotrechus nauticus, Phymatodes lecontei, a nd Neoclytus modestus modestus. J Chem Ecol 33:889–907. Iwabuchi K. 1982. Mating behavior of Xylotrechus pyrrhoderus Bates (Coleoptera: Cerambycidae) I. Behavioral sequence and existence of the male sex pheromone. Appl Entomol Zool 17:494–500. Iwabuchi K. 1986. Mating behavior of Xylotrechus pyrrhoderus Bates (Coleoptera: Cerambycidae) III. Pheromone secretion by male. Appl Entomol Zool 21:606–612. Iwabuchi K. 1999. Sex pheromones of cerambycid beetles. In: Hidaka T, Matsumoto Y, Honda K, Honda H, Tatsuki S, editors. Environmental Entomology. Tokyo: University of Tokyo Press. p 436–451. Kojima K, Nakamura S. 2011. Food plants of Cerambycid beetles (Cerambycidae: Coleoptera) in Japan (revised and enlarged edition). Hiroshima, Japan: Hiba Society of Natural History. Lacey ES, Ginzel MD, Millar JG, Hanks LM. 2004. Male-produced aggregation pheromone of the cerambycid beetle Neoclytus acuminatus acuminatus. J Chem Ecol 30:1493–1507. Lacey ES, Ray AM, Hanks LM. 2007. Calling behavior of the cerambycid beetle Neoclytus acuminatus acuminatus (F.). J Insect Behav 20:117–128. Lai-Fook J. 1970. The fine structure of developing type “B” dermal glands in Rhodnius prolixus. Tissue Cell 2:119–138. Leal WS, Bento JMS, Vilela EF, Lucia TMCD. 1994. Female sex pheromone of the longhorn beetle Migdolus fryanus Westwood: N(20 S)-methylbutanoyl 2-methylbutylamine. Experientia 50:853–856. Luft JH. 1961. Improvements in epoxy resin embedding method. J Biophys Biochem Cytol 9:409–414. Nakamuta K, Sato H, Nakashima T. 1994. Behavioral and morphological evidence for a male-produced sex pheromone in the cryptomeria twig borer, Anaglyptus subfasciatus PIC (Coleoptera, Cerambycidae). Jpn J Ent 62:371–376. Nardi JB, Dowd PF, Bartelt RJ. 1996. Fine structure of cells specialized for secretion of aggregation pheromone in a nitidulid beetle Carpophilus freemeni (Coleoptera: Nitidulidae). Tissue Cell 28:43–52. Neville AC. 1975. Biology of the arthropod cuticle. Berlin: SpringerVerlag. Noirot C, Quennedey A. 1974. Fine structure of insect epidermal glands. Annu Rev Entomol 19:61–80. Noldt U, Fettköther R, Dettner K. 1995. Structure of sex pheromoneproducing prothoracic glands of the male old house borer, Hylotrupes bajulus (L.) (Coleoptera: Cerambycidae). Intl J Insect Morphol Embryol 24:223–234. Ohbayashi N, Niisato T. 2007. Longicorn beetles of Japan. Hadano, Japan: Tokai University Press. Pajares JA, Alvarez G, Hall DR, et al. 2013. 2-(Undecyloxy)-ethanol is a major component of the male-produced aggregation pheromone of Monochamus sutor. Entomol Exp Appl 149:118–127. Percy-Cunningham JE, MacDonald JA. 1987. Biology and ultrastructure of sex pheromone-producing glands. In: Prestwich GD, J. Exp. Zool. (Mol. Dev. Evol.)
HOSHINO ET AL. Blomquist GJ, editors. Pheromone biochemistry. New York: Academic Press. p 27–75. Posada D, Crandall KA. 2001. Selecting the best-fit model of nucleotide substitution. System Biol 50:580–601. Ray AM, Lacey ES, Hanks LM. 2006. Predicted taxonomic patterns in pheromone production by longhorned beetles. Naturwissenschaften. 93:543–550. Ray AM, Zuni A, Alten RL, et al. 2011. Cis-Vaccenyl acetate, a femaleproduced sex pheromone component of Ortholeptura valida, a longhorned beetle in the subfamily Lepturinae. J Chem Ecol 37:173–178. Ray AM, Swift IP, McElfresh JS, Alten RL, Millar JG. 2012a. (R)desmolactone, a female-produced sex pheromone component of the cerambycid beetle Desmocerus californicus californicus (subfamily Lepturinae). J Chem Ecol 38:157–167. Ray AM, Barbour JD, McElfresh JS, et al. 2012b. B 2,3-Hexanediols as sex attractants and a female-produced sex pheromone for cerambycid beetles in the prionine genus Tragosoma. J Chem Ecol 38:1151–1158. Rodstein J, Millar JG, Barbour JD, et al. 2011. Determination of the relative and absolute configurations of the female-produced sex pheromone of the cerambycid beetle Prionus californicus. J Chem Ecol 37:114–124. Ronquist F, Huelsenbeck JP. 2003. MRBAYES 3: Bayesian phylogenetic inference under mixed models. Bioinformatics 19:1572–1574. Silk PJ, Sweeney J, Wu J, et al. 2007. Evidence for a male-produced pheromone in Tetropium fuscum (F.) and Tetropium cinnamopterum (Kirby) (Coleoptera: Cerambycidae). Naturwissenschaften 94:697–701. Spiegel CN, Brazil RP, Soares MJ. 2002. Ultrastructure of male sex pheromone glands in abdominal tergites of five Lutzomyia sandfly species (Diptera: Psychodidae). Arth Struct Dev 30:219–227. Spiegel CN, Brazil RP, Soares MJ. 2004. Ultrastructural cytochemistry of the sex pheromone glands of Lutzomyia cruzi male sand flies (Diptera: Psychodidae: Phlebotominae). Arth Struct Dev 33: 399–404. Swofford DL. 2003. PAUP*. Phylogenetic Analysis Using Parsimony (*and Other Methods) 4.0 Beta. Sunderland, MA: Sinauer Associates. Teale SA, Wickham JD, Zhang F, et al. 2011. A male-produced aggregation pheromone of Monochamus alternatus (Coleoptera: Cerambycidae), a major vector of pine wood nematode. J Econ Entomol 104:1592–1598. Thompson JD, Higgins DG, Gibson TJ. 1994. CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, positions-specific gap penalties and weight matrix choice. Nucleic Acids Res 22:4673–4680. Wattebled S, Bitsch J, Rousset A. 1978. Ultrastructure of pheromoneproducing eversible vesicles in males of Chrysopa perla L. (Insecta. Neuroptera). Cell Tiss Res 194:481–496. Wigglesworth VB. 1972. The principles of insect physiology. London: Chapman and Hall.
Structure and Development of Male Pheromone Gland of Longicorn Beetles and its Phylogenetic Relationships Within the Tribe Clytini KEITA HOSHINO*, SATOSHI NAKABA, HIROKI INOUE, AND KIKUO IWABUCHI Faculty of Agriculture, Tokyo University of Agriculture and Technology, Fuchu, Tokyo, Japan
ABSTRACT
J. Exp. Zool. (Mol. Dev. Evol.) 324B:68–76, 2015
The male sex pheromone of the longicorn beetle, Xylotrechus pyrrhoderus pyrrhoderus Bates (Cerambycidae: Tribe Clytini) plays an important role in attracting females. This pheromone is produced by the pheromone gland located in the prothorax. However, the detailed structure and underlying developmental process of this gland are still unknown. We investigated the gland structure by using histological analysis and confirmed that the gland consists of the following parts: gland cell mass, a unique spherical space in the cuticle layer, and ductules connecting the gland cells with the spherical space and conducting canals to the outer opening. The gland structure first appeared male-specific in the late pupal stage, during which the epidermal cells began depositing the exocuticle; the development of the gland was completed after adult emergence. Furthermore, we verified the structural equivalents of the X. p. pyrrhoderus male pheromone gland in 11 species of 2 tribes, Clytini and Anaglyptini. The glands of these insects could be classified into four types on the basis of the absence or presence of the spherical space and the division of the gland cell mass layer. Most noteworthy, all the species with the spherical space and division-type gland were restricted to the Xylotrechus clade, as inferred from the molecular phylogenetic analysis. These results suggest that Clytini and Anaglyptini species share a fundamental process of male pheromone gland development, and that the Japanese Xylotrechus species might have established their current status by developing distinct structural features in the male pheromone gland. J. Exp. Zool. (Mol. Dev. Evol.) 324B:68–76, 2015. © 2014 Wiley Periodicals, Inc. How to cite this article: Hoshino K, Nakaba S, Inoue H, Iwabuchi K. 2015. Structure and development of male pheromone gland of longicorn beetles and its phylogenetic relationships within the tribe Clytini. J. Exp. Zool. (Mol. Dev. Evol.) 324B:68–76.
Longicorn beetles are a principal group in the order Coleoptera, which consists of diverse species of phytophagous borers, including many important arboreal pests. In these beetles, chemical communication systems are utilized during mating similar to those found in most other insect species. Since earlier studies have reported the presence of pheromones in longicorn beetles (Galford, '77), pheromone-mediated communication in cerambycid beetles has been investigated. Female volatile pheromones have been identified in the species belonging to subfamilies of Prioninae, Lepturinae, (Rodstein et al., 2011; Ray
Grant sponsor: Japan Society for the Promotion of Science (JSPS); grant number: 25870210. Conflict of interest: None. Correspondence to: Keita Hoshino, Faculty of Agriculture, Tokyo University of Agriculture and Technology, Fuchu, Tokyo 183-8509, Japan. E-mail: [email protected] Received 26 September 2014; Accepted 18 October 2014 DOI: 10.1002/jez.b.22606 Published online in Wiley Online Library (wileyonlinelibrary.com). .
© 2014 WILEY PERIODICALS, INC.
STRUCTURE AND DEVELOPMENT OF MALE PHEROMONE GLAND IN LONGICORN BEETLES et al., 2011, 2012a, 2012b), as well as Vesperinae (uncertain affiliation with the family Cerambycidae; Leal et al., '94). Thus far, male-produced sex or aggregation pheromones have been identified in 18 species of the subfamilies Cerambycinae, Lamiinae, and Spondylidinae (listed in Lacey et al., 2004; Silk et al., 2007; Fonseca et al., 2010; Teale et al., 2011; Fierke et al., 2012; Hanks and Millar, 2013; Pajares et al., 2013). Maleproduced pheromones have been found especially in the subfamily Cerambycinae, and chemical and ecological studies have been extensively performed in the tribe Clytini within this subfamily (Iwabuchi, '99). These pheromones are emitted from the surface of the prothorax, as indicated by the chemical analysis of individual body parts (Iwabuchi, '86) and the morphological analysis of male-specific pores in the subfamily Cerambycinae (Ray et al., 2006). Pheromone glands are located beneath the pores on the prothorax in 5 species of the subfamily Cerambycinae (Iwabuchi, '86; Nakamuta et al., '94; Noldt et al., '95; Hanks et al., 2007; Lacey et al., 2007). These structural analyses of pheromone glands, however, have not been performed in detail, and more thorough analysis will enhance the understanding of these glands. Pheromone glands have a specialized glandular function; generally, they consist of gland cells with conducting canals in the cuticle layer (Chapman, '98; Neville, '75). Female pheromone glands have been studied in detail, both morphologically and developmentally, mainly in lepidopteran insects (Percy-Cunningham and MacDonald, '87). In contrast, studies on male pheromone glands are rare, but have been reported for mecopteran, neuropteran, dipteran, and coleopteran insects (Crossley and Waterhouse, '69; Wattebled et al., '78; Nardi et al., '96; Spiegel et al., 2002, 2004). Developmental studies on male pheromone glands, in particular, will provide valuable information for understanding how male-specific developmental processes have evolved; however, no such studies have yet been conducted in male longicorn beetles. Furthermore, male pheromones of tribe Clytini are indicated to be similar at the chemical structure level (Ray et al., 2012b; Hanks and Millar, 2013); thus, a comparative analysis of the morphology of the gland structures within the tribe will aid in determining the phylogenetic relationships in the male pheromone gland evolution. Molecular-based phylogenetic analysis might enhance the accuracy of the comparative study of the glands, since phylogenetic relationships among genera of tribe Clytini are not well understood (Ohbayashi and Niisato, 2007) and have never been confirmed by molecular data. In this study, we investigated the detailed morphological and developmental processes of the male pheromone gland of Xylotrechus pyrrhoderus pyrrhoderus Bates, a clytine species that is known to have male sex pheromone (Iwabuchi, '82). We also conducted a comparative study of the male pheromone glands among another 11 species of tribe Clytini and 1 species of Anaglyptini (closely related tribe within the subfamily Ceram-
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bycinae; Ohbayashi and Niisato, 2007). Finally, on the basis of the result of molecular-based phylogenetic analysis, we discuss the development and evolution of male pheromone glands in tribe Clytini.
MATERIALS AND METHODS Insects X. p. pyrrhoderus was obtained by collecting pruned grape vine shoots with wood-boring larvae in March. The grape vine shoots were placed upright in outdoor insect cages (120 120 200 cm in height). The development of the larvae was periodically monitored. In late June, the final instar larvae (weight, >170 mg) were collected from the shoots and maintained individually in 35-mm plastic dishes. A good environment for the larvae was maintained by placing a small cutting of the grape vine shoot and wet filter paper within each dish. Dishes were maintained in the laboratory at 25 °C with a light:dark (L:D) photoperiod of 16:8. Pupae were sexed according to morphological characteristics in the apical region of the abdomen, as described by Duffy, ('53). The pupal periods and days after emergence were counted and recorded daily. Thirteen species belonging to the tribes Clytini and Anaglyptini in the subfamily Cerambycinae (Clytini: X. p. pyrrhoderus; Xylotrechus chinensis chinensis (Chevrolat); Xylotrechus rufilius Bates; Xylotrechus emaciatus Bates; Xylotrechus cuneipennis (Kraatz); Clytus melaenus Bates; Cyrtoclytus caproides caproides (Bates); Plagionotus christophi (Kraatz); Demonax notabilis notabilis (Pascoe); Rhaphuma xenisca (Bates); Rhabdoclytus acutivittis inscriptus (Bates); and Chlorophorus quinquefasciatus (Castelnau et Gory); Anaglyptini: Paraclytus excultus Bates) were used for histological analysis of the pheromone glands. Two species of different subfamilies (Lepturinae: Leptura ochraceofasciata ochraceofasciata (Motschulsky) and Necydalinae: Necydalis solida Bates) were also used for histological analysis. All specimens used for this analysis were collected and transferred alive to the laboratory. Sex was determined for all specimens on the basis of genital morphology and other accompanying sexual characters, including the shapes of the antenna and abdomen. Collection localities and numbers of all specimens used for the histological analysis were listed in supplementary materials Table S1. For the phylogenetic study, besides the above-mentioned 13 clytine and anaglyptine species, 13 species (Clytini: Xylotrechus villioni (Villard); Xylotrechus atronotatus angulithorax Gressitt; Xylotrechus grayli grayii (White); Perissus kiusiuensis kiusiuensis Ohbayashi; Clytus auripilis Bates; Brachyclytus singularis Kraatz; Epiclytus yokoyamai (Kano); Demonax transilis Bates; Chlorophorus japonicus (Chevrolat); Chlorophorus muscosus (Bates), Anaglyptini: Anaglyptus niponensis Bates; Anaglyptus matsushitai Hayashi, Callidiini: Phymatodes maakii Kraatz) and 3 additional species from a different J. Exp. Zool. (Mol. Dev. Evol.)
70 subfamilies (Disteniinae: Distenia gracilis gracilis (Blessig), Prioninae: Prionus insularis insularis Motschulsky, Lamiinae: Dolichoprosopus yokoyamai (Gressitt)) were used. In all, samples from 29 species were stored at 20°C with or without 100% ethanol until use. The data of all specimens used for the phylogenetic analysis were listed in Table S2. Preparation of Histological Samples A razor and forceps were used to obtain a prothoracic sample from each living specimen; the sample was trimmed within half a minute, and the left region was used. The prothoracic piece was immediately placed in a 2-mL plastic tube containing 500 mL of fixative solution (3% paraformaldehyde (WAKO, Osaka, Japan), 2% glutaraldehyde (TAAB, Berks, UK) in 0.1 M phosphate buffer (PB), pH 7.0) and was refrigerated for 2 hr at 4 °C. After fixation, the sample was washed by replacing the buffer solution with fresh one twice. The sample was then transferred to the post-fixative solution (2% osmium tetroxide (WAKO) in 0.1 M PB, pH 7.0) and refrigerated for another 2 hr at 4 °C. After post-fixation, the sample was washed with the buffer and dehydrated with increasing concentrations of diluted ice-cold ethanol (50, 60, 70, 80, 90, and 95%). Samples were incubated in each ethanol solution for 15 min at room temperature, except for 30 min in 95% ethanol. The samples were further dehydrated in absolute ethanol containing molecular sieves (3 A; Sigma–Aldrich, St Louis, MO, USA). After the dehydration process was repeated with absolute ethanol three times, the ethanol was replaced with a mixture containing 50% ethanol and 50% 1-butoxy 1, 3epoxypropan (QY-1; EM Japan, Tokyo). The mixture was gradually replaced with QY-1 by exchanging the medium four times and allowing the sample to stabilize for 10 min after each exchange. After the solution was replaced with 100% QY-1, QY-1 was gradually replaced with Epon resin (solution A: Epon 812 (TAAB) 580 mL and dodecanyl succinic anhydride (DDSA) (TAAB) 925 mL; solution B: Epon 812 (1,850 mg) and methyl nadic anhydride (MNA) (TAAB) 1670 mg; A:B ¼ 3:7) through serial exchanges with solutions containing an increasing proportion of Epon resin. Samples were adapted for more than 30 min after each exchange, and the final exchange solution contained only Epon resin. After the samples were placed in the resin overnight, the processed specimens were then transferred to a mold filled with the prepared resin to which DMP-30 (TAAB) was added. The specimens were then heated in an oven according to Luft ('61) and completely embedded for the next sectioning step. Thin Sectioning and Staining The embedded specimens were roughly cut out of the surplus resin surrounding the tissue and trimmed into a right trapezoidal shape. The direction of the specimen was adjusted by cutting from the longer base of the trapezoid. Approximately 1-mm sections were cut from the specimen by using an ultramicrotome J. Exp. Zool. (Mol. Dev. Evol.)
HOSHINO ET AL. (Ultracut N; Reihert, Vienna, Austria). For thin sectioning, glass knives were placed in a container filled with water, and knives were exchanged every 5–10 cuts. Each section was checked for quality by examining its brightness, picked up using a loop of a platinum–palladium wire attached to a wooden shaft, and placed in a pool of water. The sections were transferred to a staining solution of 10% azure B, 1% safranin, or 1% toluidine blue and stained for 30 min at 60 °C, 30 min at room temperature, or 5 min at room temperature, respectively. The sections were then washed by transferring to water pools 3 times and finally transferred onto glass slides. The glass slides were dried in an oven at 60 °C. Observation and Comparative Study of the Male Pheromone Gland Structure Detailed morphological analysis of the male pheromone gland was conducted for X. p. pyrrhoderus. Approximately 200 serial sections were cut and examined. By integration of sharp and well-stained microscopic images, each structure for male pheromone gland was defined and represented schematically, followed by referring to the general criteria (Neville, '75; Chapman, '98). The development of pheromone glands was elucidated by examining the various developmental stages of male X. p. pyrrhoderus at 1–2 day intervals in the period before and after adult emergence. Approximately 50 sections per stage were observed, and developmental changes were detected. For taxonomic comparison of male pheromone glands across different species, the presence of structural equivalent of male pheromone glands of X. p. pyrrhoderus was determined. Sections were sampled at every 50-mm scraping and, when the gland was detected, the structural features, especially the presence of spherical space and division or layered type of gland cell mass, were examined. For females, 8 species (X. p. pyrrhoderus; X. c. chinensis; X. emaciatus; X. cuneipennis; C. melaenus; C. c. caproides; P. christophi; and R. xenisca) were used for histological analysis. All specimens were observed under a microscope (DM LB; Leica) with an optical mode at a magnification of 200. Nucleotide Sequencing DNA was extracted from insect forelegs by using a DNeasy blood and tissue kit (QIAGEN, Hilden, Germany). Samples were extracted from frozen specimens or specimens that had been submerged in 100% ethanol. The DNA extraction was conducted according to the manufacturer's instructions, and the eluted DNA solution was washed and concentrated using the ethanol precipitation method. The concentration of the samples was measured, and the ratio of absorbance at 260 and 280 nm was determined using Nanodrop (Thermo Fisher Scientific, Wilmington, DE, USA). Samples were stored at 80°C until use. The three partial regions of 2 mitochondrial genes encoding NADH dehydrogenase subunit 5 (ND5) and cytochrome oxidase subunit 1 (CO1) and a nuclear 28 S ribosomal DNA (rDNA) were
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amplified using polymerase chain reactions (PCRs) by using Quick Taq HS Dyemix (Toyobo Co. Ltd., Tokyo, Japan). All primers used for the amplification of each target were newly designed from alignment of related genes from GenBank: CerND5-FW (50 -GGAGTTTATTTAYTRATTCG-30 ) and Cer-ND5-RV (50 -CCTAAAGTAGAAACAAYWGG-30 ) for mitochondrial ND5, Cer-CO1-FW (50 -CCCGGATTTGGRATAATYTC-30 ) and Cer-CO1RV (50 -TCAGAATATCTRTGTTCDGC-30 ) for mitochondrial CO1, and Cer-28S-rRNA-FW (50 -CTAGGATTCCCTCAGTAGCT-30 ) and Cer-28S-rRNA-RV (50 -GCTCWTCCCTKTTCGCTCGCA-30 ) for nuclear 28S-rDNA. The PCRs were performed according to the manufacturer's instructions. PCR conditions were as follows: 2 min at 94 °C, followed by 35 cycles of 30 sec at 94 °C, 30 sec at 50 °C, and 1 min at 68 °C. The amplicons were purified using agarose gel electrophoresis and then extracted from the gels using a QIAEX gel extraction kit 2 (QIAGEN). The purified DNA fragments were directly sequenced using the ABI Prism Big Dye Terminator cycle sequence kit version 3.1 (Applied Biosystems (ABI), Foster City, CA, USA) and the ABI Prism 3130 genetic analyzer. The nucleotide sequences of the amplified fragments were determined using a cycle sequencing method. The purified PCR amplicons were sequenced using a dye terminator system (ABI). The sequence data obtained were checked for signal strength and electropherogram quality by using an analysis tool (ABI) and processed for phylogenetic analysis. Phylogenetic Analysis The phylogenetic relationships among the 22 clytine and 3 anaglyptine species were analyzed by comparing the nucleotide sequences encoding mitochondrial CO1 and ND5 and nuclear rDNA. The sequence data have been submitted to GenBank, and the accession numbers are listed in Table S2. Two partial gene sequences of CO1 and ND5 were concatenated and used as mitochondrial sequences. The mitochondrial and nuclear sequences were aligned using the ClustalX program (Thompson et al., '94). The best-fitting nucleotide substitution model was determined as GTR þ G þ I model by the program MrModeltest (Posada and Crandall, 2001) executed in PAUP* 4.0b10 (Swofford, 2003). The phylogenetic dendrogram was inferred using Bayesian method by using MrBayes version 3.2 (Ronquist and Huelsenbeck, 2003).
RESULTS Morphological Observation of the Male Pheromone Glands The pheromone glands of X. p. pyrrhoderus were observed only in male samples (Fig. 1) and are represented schematically and terminologically in Figure 2. In one field of microscopic view, a single gland unit consisted of approximately 20 gland cells that were associated with the “end apparatus,” which is the terminus of an extended ductule within the gland cell cavity (Figs. 1B and 2; Chapman, '98). Immediately exterior to the gland cells, a
FIGURE 1. Micrographs of the transverse sections of adult X. p. pyrrhoderus prothorax 2 weeks after emergence. The densely stained, wavy horizontal line is the cuticle layer. The upper side of the cuticle layer is the outer space and, below this layer, is the hemocoel. (A–C): male; D: female. (A) The ovoid, multicellular tissue visible under the cuticle layer is a pheromone gland (stained blue with azure B). (B) An axial section of gland cells stained with safranin. The gland cells have a cavity (arrow, surrounded by whity region) from which the ductules (arrowhead) extend toward the spherical space. (C) A partial image of the spherical space in the cuticle layer from a section stained with toluidine blue. A bundle of gathered ductules from gland cells extended to the spherical space. (D) None of the pheromone gland structures, including gland cell mass and ductules in the cuticle layer, are found in female-derived sections stained with safranin. Bar indicates 50 mm.
cuticle consisting of two distinct layers (exocuticle and endocuticle) was visible under light microscopy (Fig. 1A, D). The exocuticle was the outer layer and had a wavy appearance. The endocuticle consisted of 7–9 layers with a striped pattern, J. Exp. Zool. (Mol. Dev. Evol.)
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FIGURE 2. Schematic representation and terminology of X. p. pyrrhoderus male pheromone gland structure. The cuticle is the uppermost part in the figure, and the hemocoel is beneath the cuticle. The colors represent exocuticle (red), endocuticle (blue), and gland cells (yellow). A spherical space is formed by exocuticle evagination. Gland cell mass consists of many individual gland cells of which some representatives, i.e., three axial sectional and one transectional images, are described. Abbreviations represented in this figure are used for Figure 3 and supplementary materials Fig. S1 and S2.
corresponding to the structure termed “balken” and known to be a distinctive feature of coleopteran insects (Wigglesworth, '72; De Camp and Greven, 2010). Observation of serial sections showed spherical structures in the endocuticle layer. This structure was merely designated as a “spherical space,” since this was previously indicated to be a reservoir (Iwabuchi, '86) of unknown function. From the gland cell mass located under each spherical space, a bundle of ductules extended to the inner part of the spherical space (Figs. 1C and 2). Observation of serial sections showed that only the superior region of the spherical space was fragile. These findings suggested the presence of some conducting canals to the outer openings from the spherical space, as indicated by previous scanning electron microscopic (SEM) observations (Iwabuchi, '86; Ray et al., 2006), but they are probably not visible because their size is less than 1 mm. Developmental Changes During the Formation of the Male Pheromone Glands The chronological process of male pheromone gland formation is represented schematically in comparison with that in females in Figure 3. At 10 days after pupation (4 days before adult emergence), spherical spaces appeared in the epidermal cell layer, during which the exocuticle layer was deposited (Supplementary materials Fig. S1). The spherical spaces were formed by local invaginations of the exocuticle into the epidermal cell layer, which was located under the depressions of the cuticular surface (Fig. 3). After the spherical space formed, a bundle of ductules and gland cells appeared beneath each spherical space (Fig. 3 and S1). J. Exp. Zool. (Mol. Dev. Evol.)
After adult emergence, there was no change in the structure of the gland cells, except for some increase in size. On the other hand, the endocuticle grew at a rate of one layer per day, and epidermal cells gradually degenerated. These changes ceased within 7–9 days after adult emergence. Pheromone glands were observed only in males, but cuticle growth and dermal gland formation were detected in both the sexes and followed the same developmental course. Taxonomic Comparisons of Male Pheromone Glands Across 11 Longicorn Species Of the 12 species, 11 had typical male pheromone glands that were similar to those of X. p. pyrrhoderus (Fig. S2). On the other hand, no pheromone glands were found in either sex in X. cuneipennis (data not shown) and two more distantly related species of subfamilies Lepturinae and Necydalinae (Fig. S2). In all the species with pheromone glands, the gland cells exhibited some common structural features such as end apparatuses and filamentous ductules. However, two obvious differences were detected, and we classified the glands on the basis of these differences. The primary classification was made on the basis of the absence or presence of a spherical space (type spherical space (SS)-positive: “ þ SS” and SS-negative: “ SS”), and the subdivision on the basis of the division of the gland cell mass layer with the basal lamina (type “division” and “layer”; see Fig. 4). The gland types of each species were as follows: þSSdivision (X. p. pyrrhoderus, X. c. chinensis, X. emaciatus, X. rufilius, and C. melaenus), þSS-layer (R. xenisca, P. christophi,
STRUCTURE AND DEVELOPMENT OF MALE PHEROMONE GLAND IN LONGICORN BEETLES
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FIGURE 3. Schematic representation of the transverse sections of male and female X. p. pyrrhoderus at each developmental stage, illustrating the development of the male pheromone gland. The numbers represent the number of days before ( ) or after (þ) adult emergence (AE). Abbreviations are as in Figure 2. Colors are as in Figure 2; grey and brown represent epidermal cell layer and pupal skin, respectively. Deposited exocuticle evaginated to form the spherical space, beneath which the swollen epidermal cells differentiated into ductules and gland cells (yellow region) in the males. Endocuticle growth and dermal gland formation occurred in both the sexes after AE. ps, pupal skin; ec, epidermal cells.
and C. c. caproides), SS-division (C. quinquefasciatus), and – SS-layer (P. excultus and R. acutivittis inscriptus). Molecular Phylogeny and its Relation to the Morphology of the Pheromone Glands The sequence alignment data were analyzed using Bayesian methods, and the obtained consensus trees for mitochondrial
gene are shown in Figure 5. Xylotrechus species formed the clade with those of the near-related genera Clytus and Perissus (Ohbayashi and Niisato, 2007), and Cyrtoclytus, including Brachyclytus species, formed the Xylotrechus clade. All other Clytini species were included in the sister group of Xylotrechus clade, whereas Anaglyptini and Callidiini were positioned between Xylotrechus clade and the outgroups. Species with
FIGURE 4. Classification of male pheromone glands detected in tribes Clytini and Anaglyptini, according to two structural characteristics: spherical space (SS) and gland cell mass layer. Types “ þ SS” and “ SS” indicate the presence and absence of spherical space, respectively. Types “division” and “layer” indicate divided gland cell mass layer by basal lamina and non-divided gland cell mass layer, respectively. The characteristics are represented in combinations as types þSS-division, þSS-layer, –SS-division, and SS-layer. Schematic diagrams represent the outlines of the gland types on the basis of Figure 2, and individual gland cells of the divided or only layered gland cell mass are not shown.
J. Exp. Zool. (Mol. Dev. Evol.)
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FIGURE 5. Dendrograms showing the relationships of 25 longicorn beetle species in the tribes Clytini and Anaglyptini (subfamily Cerambycinae), as deduced from the molecular data. The nucleotide sequences of partial mitochondrion genes CO1 and ND5 were used for Bayesian analysis. All branches are labeled with Bayesian posterior probabilities. The phylogenetic areas of tribe Clytini and Xylotrechus clade are shaded. Each species' pheromone gland types (refer to Fig. 4) are mentioned after the species' names.
spherical space-positive glands were intensively distributed in the Xylotrechus clade, which was also closely linked with the subtype of the division, whereas the remaining types were scattered across the remaining taxonomic groups (Fig. 5). In contrast, the nuclear gene tree showed no significant resolution because of high gene conservation (data not shown).
DISCUSSION The principal male pheromone gland structure consists of gland cells and cuticular ducts that are similar to those found in many other typical pheromone glands (Percy-Cunningham and MacDonald, '87). The gland cells of X. p. pyrrhoderus were similar to those of the pheromone glands of coleopteran species (Nardi et al., '96) and the male-specific gland of the phlebotomine fly (Spiegel et al., 2002, 2004). However, the pheromone glands of X. p. pyrrhoderus differed from those of the abovementioned species in that (1) the ductules from each gland cell gathered to form a bundle of ductules that penetrated the endocuticle and reached the spherical space, and (2) the gland cells were clustered together. Such a bundle of ducts has also been observed in two North American clytine species, Xylotrechus nauticus and Neoclytus acuminatus acuminatus (Hanks et al., 2007; Lacey et al., 2007), but they lack the spherical spaces in the endocuticle J. Exp. Zool. (Mol. Dev. Evol.)
layer. Consequently, to our knowledge, this study confirmed the detailed structures of the pheromone glands for the first time (Fig. 2) and revised and renewed the previous representation by Iwabuchi ('86), especially, with regard to the presence of a spherical space in the endocuticle layer but made of exocuticle, and that the ductules extended from the end apparatus that showed characteristics of class-3 gland cells defined by Noirot and Quennedey, ('74). Initial gland formation was observed as the appearance of spherical spaces in the epidermal cell layer. Thereafter, the spherical space was strongly linked with the differentiation of the male pheromone gland; that is, gland cells and ductules arose in the swollen regions of the epidermis by the formation of the spherical space (Fig. 4). In the swollen region, imaginal epidermal cells differentiated into the male pheromone gland, probably because of the initiation of mitotic division that occurred along the vertical and oblique axes, as suggested by Corbet and Laifook ('77) and Percy-Cunningham and MacDonald ('87). There were very few pheromone gland openings on the surface compared to the number of gland cells, indicating that the spherical spaces served as the connecting point between ductules and conducting canals. Therefore, the spherical spaces might allow large-scale pheromone production by increasing the
STRUCTURE AND DEVELOPMENT OF MALE PHEROMONE GLAND IN LONGICORN BEETLES number of pheromone glands per opening pore. The dermal glands also generally consist of plural cells and are classified as class-3 gland cells; during their morphogenetic process, the conducting canal and gland cells are connected via a saccule for product storage and chemical reaction (Lai-Fook, '70). Additionally, dermal glands were common to both the sexes, and SEM observation indicated that the pores of male dermal glands and the pheromone glands within a single depression differed in number but were similar in size and other properties (Iwabuchi, '86). Therefore, the canals and pores might be formed through the same process; however, after formation, the ductules connect to the canals via saccules in the dermal glands and spherical spaces in the pheromone glands. Such similarities suggest that the spherical spaces might induce the differentiation of the male pheromone glands and might function as receptacles for the final conversion and storage of the male sex pheromones. Pheromone glands appeared before the formation of dermal glands, suggesting that male pheromone glands might be derived from a heterochronical shift of the universal gland developmental program. The structure of pheromone glands of Hylotrupes bajulus belonging to the tribe Hylotrupini and subfamily Cerambycinae is similar to that of X. p. pyrrhoderus (Noldt et al., '95). This is also reminiscent of the slight developmental changes reflected in the phylogeny. However, further investigations at the ultrastructural level are needed to clarify the developmental and phylogenetic similarities. Male pheromone glands of the 11 clytine species and 1 anaglyptine species were structurally similar to those of X. p. pyrrhoderus. The pheromone glands of these species could be classified into four types (see Fig. 4), but the other structural features were common, suggesting that the glands had diverged from a prototype, and this might explain the similarity of the chemical motif found in male pheromones of tribe Clytini (Ray et al., 2012b; Hanks and Millar, 2013). By using molecular-based phylogenetic analysis, we determined the accurate phylogenetic relationships among the genera of tribe Clytini and showed the distribution patterns of the gland types in the tribe. All the species with þ SS-division-type gland were included in the robust Xylotrechus clade, unlike the distribution of the other type species that were extended to the distant anaglyptine species clade. The Xylotrechus clade included some species with a remarkably narrow range of host plant species and living conditions (X. p. pyrrhoderus, X. c. chinensis, X. villioni, etc.), and the majority of the species of this clade preferred freshly fallen trees (Ohbayashi and Niisato, 2007; Kojima and Nakamura, 2011). However, P. christophi that have strict host preference for freshly fallen trees and showed the þSS but layer-type gland were not included in the Xylotrechus clade. In contrast, X. cuneipennis was included in the Xylotrechus clade, but it has no glands in either sex, possibly because of the secondary loss that seems to be associated with its preference for decayed trees unlike the Japanese Xylotrechus. Therefore, the Japanese Xylotrechus clade species
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might have evolved the spherical space and division-type glands in association with some specific ecological demands. Taken together, our findings suggest that Clytini and Anaglyptini species share a fundamental process of male pheromone gland development, and that Xylotrechus clade species might have established their current status by developing distinct structural features of the gland in agrrement with the phylogenetic relationships within the tribe Clytini. Recently, Ray et al. (2012b) reported that the parsimonious chemical motif was extended to subfamily Prioninae and that the conversion of the sexes that produce pheromones could occur. Accordingly, further studies over an extended range of longicorn species are needed to reveal the earlier evolutionary processes of male pheromones in longicorn beetles.
ACKNOWLEDGMENTS The authors thank Mr. Nazumi Mandokoro for providing the specimen of X. villioni larva and Miss Ami Ogura for her help with the field collection of X. p. pyrrhoderus larvae. We are also thankful to Prof. Ryo Funada and the members of the Laboratory of Morphogenesis of Plant Resources of Tokyo University of Agriculture and Technology for facilitating this research. We also thank Prof. Kazuhiko Sakurai of Seijo University for his field research. This work was partially supported by Japan Society for the Promotion of Science (JSPS) KAKENHI Grant Number 25870210 to KH.
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