- Research article
- Open Access
Identification and functional analyses of sex determination genes in the sexually dimorphic stag beetle Cyclommatus metallifer
© Gotoh et al. 2016
Received: 12 August 2015
Accepted: 24 February 2016
Published: 22 March 2016
Genes in the sex determination pathway are important regulators of sexually dimorphic animal traits, including the elaborate and exaggerated male ornaments and weapons of sexual selection. In this study, we identified and functionally analyzed members of the sex determination gene family in the golden metallic stag beetle Cyclommatus metallifer, which exhibits extreme differences in mandible size between males and females.
We constructed a C. metallifer transcriptomic database from larval and prepupal developmental stages and tissues of both males and females. Using Roche 454 pyrosequencing, we generated a de novo assembled database from a total of 1,223,516 raw reads, which resulted in 14,565 isotigs (putative transcript isoforms) contained in 10,794 isogroups (putative identified genes). We queried this database for C. metallifer conserved sex determination genes and identified 14 candidate sex determination pathway genes. We then characterized the roles of several of these genes in development of extreme sexual dimorphic traits in this species. We performed molecular expression analyses with RT-PCR and functional analyses using RNAi on three C. metallifer candidate genes – Sex-lethal (CmSxl), transformer-2 (Cmtra2), and intersex (Cmix). No differences in expression pattern were found between the sexes for any of these three genes. In the RNAi gene-knockdown experiments, we found that only the Cmix had any effect on sexually dimorphic morphology, and these mimicked the effects of Cmdsx knockdown in females. Knockdown of CmSxl had no measurable effects on stag beetle phenotype, while knockdown of Cmtra2 resulted in complete lethality at the prepupal period. These results indicate that the roles of CmSxl and Cmtra2 in the sex determination cascade are likely to have diverged in stag beetles when compared to Drosophila. Our results also suggest that Cmix has a conserved role in this pathway. In addition to those three genes, we also performed a more complete functional analysis of the C. metallifer dsx gene (Cmdsx) to identify the isoforms that regulate dimorphism more fully using exon-specific RNAi. We identified a total of 16 alternative splice variants of the Cmdsx gene that code for up to 14 separate exons. Despite the variation in RNA splice products of the Cmdsx gene, only four protein isoforms are predicted. The results of our exon-specific RNAi indicated that the essential CmDsx isoform for postembryonic male differentiation is CmDsxB, whereas postembryonic female specific differentiation is mainly regulated by CmDsxD.
Taken together, our results highlight the importance of studying the function of highly conserved sex determination pathways in numerous insect species, especially those with dramatic and exaggerated sexual dimorphism, because conservation in protein structure does not always translate into conservation in downstream function.
From the colorful tail feathers of peacocks to the enormous antlers of deer, exaggerated sexual structures are ubiquitous across the animal kingdom [1–6]. Among the most extreme examples of sexually-selected traits are the weapons of beetles. Many species of scarab beetles have horns jutting out from the head and thorax and many stag beetles have elongated mandibles [6–8]. All are used as weapons for male-male combat over access to reproduction. The shapes and sizes of beetle weapons differ widely among families, genera, and sometimes even between populations of the same species, and most of these weapons are sexually dimorphic [9, 10].
Although much research has focused on the function and evolution of beetle weapons (e.g. [11–14]), the proximate mechanisms regulating their development are also beginning to be explored. Recent advances in next-generation sequencing technologies and the successful utilization of RNA interference (RNAi) have enabled researchers to delve into the genetic mechanisms underlying the growth, expression, and condition-dependence of these traits. Several recent studies focusing on horns in scarab beetles have revealed many of the genetic mechanisms regulating horn development, growth, and condition-dependence [15–17]. Because of their sex-specific expression, the sex determination genes of insects, which are highly conserved across the arthropods and animals in general , are particularly important to understanding the development of sexually selected weapon traits .
Despite the overall conservation in sexual differentiation genes in arthropods, the initial cues of the sex determination cascade are highly diverse both at the chromosomal level (e.g. XX-XY in Drosophila, ZW-ZZ in Lepidoptera, XX-XO in aphids, and haplodiploidy in Hymenoptera) and at the level of the molecular signal (e.g. sex-dependent activation of the Sex-lethal (Sxl) gene in Drosophila [18, 19], transformer (tra) in Ceratitis [20, 21], and sex-specific expression of piRNA fem in Bombyx ). Diversity among species is also found in the maternal transfer of tra mRNA in Nasonia  and heterozygosity in the complementary sex determiner (csd) gene’s activation of the gene feminizer (fem) in honeybees . Although the relative importance of these upstream signals appears to vary from species to species, as does the default activation state of the tra and fem genes (ON in females versus OFF in males), all sex determination mechanisms known for insects thus far integrate expression of this same network of genes , and they all result in the sex-specific expression of alternative splice variants of the sex determination gene doublesex (dsx), which, in turn, appears to trigger sex-specific patterns of growth of dimorphic body parts [18, 25–27].
Recently several authors, including our group, have shown that dsx regulates sex-specific trait expression in beetles, including species with horns and elongated mandibles [28–31]. However, very little is known about the initial cues of sex determination in these species, or about pathway members acting upstream of dsx in the sex determination cascade.
The gene Sxl is activated in Drosophila females based on the dose of X chromosome [18, 32]. Once activated, female splice forms of Sxl appear to control their own expression by a positive feedback splicing mechanism, resulting in the regulation and splicing of downstream genes such as tra and dsx in a sex-specific manner . The Sxl gene is not activated in Drosophila males, and, in the absence of this signal, male-specific tra and dsx splice variants are expressed. The role of Sxl in sexual differentiation appears to be specific to the Drosophila lineage, though the functions of Sxl outside of Drosophila are not well known [18, 34].
The ix gene is also necessary for female development in Drosophila and has been shown to act in concert with dsx in the sex determination pathway [35, 36]. tra and tra2 are required for female sexual development in many insect groups including Drosophila, Apis and Tribolium, and function by regulating the alternative splicing of dsx [21, 33, 37, 38]. tra is expressed as different splice variants in a sex-specific manner  and, as with Sxl in Drosophila, female specific splice forms of tra in some insects contain an autoregulatory domain that appears to result in self-activation of expression, with the resulting positive feedback contributing to sex differences in expression and splice forms of downstream genes in the pathway [21, 37]. What roles these genes may have in regulating sexual dimorphisms in animals with sexually-selected weapons has yet to be explored.
We examined the expression pattern of C. metallifer tra, ix, Sxl and tra2 (hereafter, Cmtra, Cmix, CmSxl and Cmtra2, respectively) using RT-PCR. Also, we analyzed the function of Cmix, CmSxl and Cmtra2 by RNA interference (RNAi). Functional analyses revealed that of these, Cmix is necessary for female differentiation, while CmSxl is not necessary for sexual trait differentiation in either males or females. Knockdown of Cmtra2 resulted in death at the prepupal stage in both sexes, suggesting that Cmtra2 likely has an additional function during development besides its known role in sex determination. In this study we also report the functional characterization of male and female specific isoforms of the sex determination gene C. metallifer doublesex (Cmdsx), which was initially identified in a previous study by our group. Functional analyses via RNAi for isoform-specific Cmdsx genes revealed that two have critical roles in sex-specific weapon trait differentiation in C. metallifer.
Sequencing and assembly
Newbler 2.9 assmbly metrics
Number of aligned reads
1106461 (90.43 %)
353508757 (91.32 %)
“Large” contigs (>500 bp)
10468 (58.5 %)
Isotigs with BLASTx results (e-5)
12056 (82.77 %)
Isotigs with homology to Tribolium (e-5)
9492 (78.73 %)
Identification of sex-determination cascade genes
Summary of database search for candidate genes
Candidate gene name (ab)
Accession No. of Drosophila homolog for query
Accession No. of Tribolium homolog for query
Isotig number of the homolog or highest similarity (Drosophila query)
Isotig number of the homolog or highest similarity (Tribolium query)
e-value (Drosophila query)
e-value (Tribolium query)
Longest hit length and the percentage of amino acid identity (Drosophila query)
Longest hit length and the percentage of amino acid identity (Tribolium query)
Identified as Cyclommatus homolog of
321/355 (90 %)
323/348 (93 %)
133/157 (85 %)
181/192 (94 %)
85/98 (87 %)
99/106 (93 %)
39/69 (57 %)
194/248 (78 %)
97/125 (78 %)
124/156 (79 %)
62/80 (78 %)
81/126 (64 %)
89/98 (91 %)
122/126 (97 %)
female lethal d (fl(2)d)
55/63 (87 %)
116/142 (82 %)
female lethal d (fl(2)d)
73/118 (62 %)
109/132 (83 %)
59/107 (55 %)
111/120 (93 %)
29/47 (62 %)
108/143 (76 %)
63/128 (49 %)
68/100 (68 %)
40/70 (57 %)
88/112 (79 %)
38/107 (36 %)
41/107 (38 %)
Not found in OrthoDB
11/34 (32 %)
Not found in OrthoDB
12/27 (44 %)
E3 ubiquitin-protein ligase
Not found in OrthoDB
14/24 (58 %)
ovarian tumor (otu)
16/46 (35 %)
11/37 (30 %)
Hypothetical protein/terribly reduced optic lobes (trol)
Not found in OrthoDB
11/34 (32 %)
Brain-enriched guanylate kinase-associated protein
10/27 (37 %)
17/41 (41 %)
mitochondrial ribosomal protein L1/ transformer (tra)
14/33 (42 %)
11/18 (61 %)
ribosomal protein L4e/egalitarian (egl)
10/19 (53 %)
16/31 (52 %)
NSFL1 cofactor p47/Ubiquinol-cytochrome c reductase core protein 1
Not found in OrthoDB
17/45 (38 %)
Not found in OrthoDB
11/30 (37 %)
High mobility group protein DSP1
Expression analyses of CmSxl, Cmix, and Cmtra2
Functional analyses of Sxl, ix and tra2
Identification of undescribed dsx variants by RACE
Functional analyses of splice variant specific knockdown of Cmdsx on sex-specific growth of mandibles
The developmental function of Cmdsx was previously tested using RNAi against a common region (exon 1) to knockdown all CmDsx isoforms . In this study, we more precisely determined the specific function of each Cmdsx splice variant using exon-specific RNAi. We designed dsRNA against Cmdsx exon 4, exon 6–7, exon 8a–8b (hereafter exon 8) and exon 9a–9b (hereafter exon 9) which enabled us to knock down each isoform specifically.
In males, only Cmdsx-exon 9 knockdown affected male-specific morphology, including mandible length (Fig. 7). There was no significant difference in the scaling relationship between Cmdsx-exon 9 RNAi males and exon 1 RNAi males in mandible size (P = 0.661 slope and P = 0.057 intercept, ANCOVA; Fig. 7), while Cmdsx-exon 9 RNAi males had significantly shorter mandibles compared to GFP RNAi controls in our observed body size range (P = 0.0004, ANCOVA; Fig. 7). Normal development of other male-specific traits such as the tibial spines and body color were also affected by Cmdsx-exon 9 RNAi. In contrast, knockdown of Cmdsx-exon 4, exon 6–7 and exon 8 did not affect male mandible scaling relationships (P > 0.05 in both slope and intercept for all three treatments against GFP RNAi; ANCOVA), or tibial spines or body color at detectable levels (Fig. 7). In females, Cmdsx-exon 4 and exon 6 knockdown did not affect morphology (Fig. 7). There were no significant differences in mandible size compared with GFP RNAi control females (P > 0.05 in both slope and intercept, ANCOVA). In contrast, Cmdsx-exon 8 knockdown transformed females to an intersex morphology (Fig. 7); mandibles were relatively longer than those of GFP RNAi control females (P = 0.0244, ANCOVA). Knockdown of Cmdsx-exon 9 in females also slightly, but significantly, increased mandible length. There was no significant difference for the regression slopes between GFP and Cmdsx-exon 9 RNAi females (P = 0.0847, ANCOVA), but the intercepts of these regression lines differed significantly (P = 0.0098, ANCOVA).
Transcriptome assembly and annotation
The stag beetle C. metallifer transcriptomic database we have constructed is a valuable new resource for molecular screening and candidate gene identification in comparative studies of insect biology, particularly those involving exaggerated, sexually selected and sexually dimorphic structures. Our transcriptomic database covers more than 50 % of the total genes in the genome of C. metallifer, based on the number of genes predicted based on the Tribolium castaneum genome (16,404 genes ) and other holometabolous insect genomes (approximately 10,000 – 19,000 genes ). Accordingly, more than half (14 out of 24 candidates) of the genes in the Drosophila sex determination cascade were contained in our assembled database, suggesting that our database is a useful platform for molecular screening and candidate gene identification in this species. In this study, we extracted RNAs from larval and prepupal heads and thoraces of both sexes. Although this time period is critical to organ growth, and was selected to investigate exaggerated mandible growth [31, 39], future studies may wish to focus on other time periods (embryo, larval, pupal and adult period) and other body parts (for example, the whole embryo, whole thorax and abdomen), in order to recover additional genes expressed in this species.
Insights for the evolution of insect sex-determination mechanisms and development of sexually dimorphic weapons
Expression and functional analyses of three C. metallifer genes in the Drosophila sex determination pathway revealed both diversifications of function (CmSxl and Cmtra2) as well as conservation (Cmix) at least in postembryonic sexual character development. Our results are the first report of expression and functional analysis of the Sxl gene in a beetle species. Sxl acts as one of the earliest selector genes for sex determination in Drosophila [19, 43, 44]. Functional Drosophila Sxl is produced only in females, which splice downstream tra mRNA to the functional female isoforms [43, 44]. In addition to controlling the sex-specific splicing of tra transcripts, Sxl also promotes its own expression, using an auto-regulatory splicing loop, and activates downstream genes such as tra2 and dsxF to establish female identity . Since Sxl also regulates female-specific dosage compensation, loss-of-function mutants of Sxl are lethal only in females [34, 46]. However, outside of flies, the function of Sxl in other insects is not well known . We found uniform expression and no detectable defect in knockdown animals suggesting that CmSxl does not have a significant role in postembryonic sexual character development in male or female C. metallifer. These results are consistent with previous expression and functional studies in non-Drosophilid flies and Lepidopteran insects, suggesting that the Sxl sex determination function is not a common mechanism among insects  and likely evolved in the Drosophilid lineage. Although CmSxl did not have a phenotype when knocked down early in the prepupal period of C. metallifer, to fully understand its function it will be necessary to investigate additional developmental stages, especially in early embryos when the initial sex-determination signal is likely to function.
Knockdown of the RNA binding protein Tra2 had lethal effects in C. metallifer, so we could not examine its effects on adult sexually dimorphic traits. In Drosophila, Tra2 functions as a co-factor with Tra for splicing dsx transcripts to female-specific dsx mRNA [35, 48, 49]. When Tra2 is not present, Tra fails to splice dsx into the female specific transcript, resulting in a male-like phenotype. This sex-specific splicing function of Tra2 is conserved in other insects such as Apis mellifera  and Tribolium castaneum . Based on these data, we predicted that Cmtra2 RNAi would transform females to a more male-like phenotype in the stag beetle. However, contrary to our prediction, all of the Cmtra2 RNAi male and female larvae died during the prepupal periods without molting into pupae. Similar non-sex specific lethality of tra2 RNAi was also reported from the red flour beetle Tribolium castaneum  and the honeybee Apis mellifera . Shukla and Palli (2013) have suggested that Tra2 and Tra inhibit the formation of the dosage compensation complex in females and Tra2 is necessary for the formation of the dosage compensation complex in males . However, this hypothesis has not been tested. Additional details of the role of tra2 during development need to be examined to understand the multiple functions of this gene in insects.
In addition to Cmtra2, we also identified a putative homolog of tra from our transcriptome. In many previously studied insects, including Tribolium, tra is known as an up-stream regulating factor of dsx-mediated sex-specific alternative splicing [37, 52]. Only females produce functional Tra protein, which results in production of the female-type Dsx isoform. The functional Tra protein also splices the tra transcript itself. By this positive auto-splicing feedback, female sexuality is fully established . On the other hand, in males, non-functional Tra protein is produced, and Dsx is consequently expressed in the male-specific isoform. Considering that CmTra also has two sex-specific isoforms and only the female-specific isoform has a complete auto-regulation domain, female-specific auto-splicing feedback might be conserved in Cyclommatus sex-determination. Further investigations of Cmtra function via RNAi and expression analyses of splice patterns of dsx and tra transcripts in Cmtra RNAi knockdown individuals are necessary for full confirmation of this possibility.
Our data support the conserved function of the Ix protein as a co-factor with female-specific Dsx protein [36, 53]. In Drosophila females, ix mutant phenotypes are highly similar to dsx mutants, while in males, ix mutants do not show any detectable morphological defects . Knockdown of the ix gene in the milkweed bug Oncopeltus fasciatus affected both male and female genital morphological structures . In C. metallifer, Cmix RNAi affected sex specific trait expression in females only. Interestingly, female Cmix RNAi external morphological phenotypes were not detectably different from Cmdsx RNAi phenotypes. Based on this, we suggest that CmIx acts together with CmDsxF to specify female differentiation. Although we cannot detect any morphological differences between Cmix RNAi and wild type males, we did not evaluate the reproductive abilities of these knockdown animals. RT-PCR indicated that Cmix is expressed in male tissues, suggesting that it may have some function in males that we did not detect. Further analyses on the non-morphological traits such as fertility of males, as well as perturbations at additional developmental periods (e.g. early embryos), will be needed to more fully characterize the role of Cmix in males.
Sexual differentiation is regulated by sex-specific expression of Doublesex isoforms
We identified additional splice variants of the Cmdsx gene. All of the new transcripts are predicted to encode one of the four previously described isoforms (CmDsxA, CmDsxB, CmDsxC or CmDsxD; ). In males, neither Cmdsx-exon 4 RNAi nor exon 6-7 RNAi, which are predicted to knockdown the male biased isoform CmDsxA, affected male morphology. These results suggest that CmDsxA may not have a significant role in male differentiation even though this transcript has a male-biased expression pattern . In females, Cmdsx-exon 6-7 RNAi did not cause any detectable phenotypes although it is predicted to be included in female specific isoform CmDsxC.
In contrast to knockdown of CmDsxA and CmDsxC, knockdown of CmDsxB and CmDsxD did influence sex-specific morphogenesis in males and females. In males, Cmdsx-exon 9 RNAi (predicted to knockdown male specific isoform CmDsxB) resulted in a more female-like phenotype (Fig. 7). These results suggest that CmDsxB is essential for male differentiation. The amino acid sequence of CmDsxB also has high similarity to male-specific Dsx proteins from other beetles [28, 30]. In females, C. metallifer CmDsxD specific knockdown (Cmdsx-exon 8 RNAi) resulted in a more male-like phenotype (Fig. 7). Thus, in the stag beetle, only CmDsxB and CmDsxD appear critical for generating sex specific morphologies during male and female differentiation. These results indicate a redundancy of CmDsx isoforms in postembryonic sexual differentiation in this species.
Redundancy of Dsx isoforms has also been reported in other beetle species. For example, in the Asian rhinoceros beetle T. dichotomus, two different female specific isoforms of Dsx (DsxF-Short and DsxF-Long) have been reported . However, the T. dichotomus DsxF-Short specific knockdown did not cause any detectable deformation phenotype in either sex . Also, Kijimoto et al. (2012) reported that Onthophagus taurus has several Dsx isoforms and knockdown of one of them (OtDsxb) did not produce any noticeable morphological defects . It is possible that these isoforms are not redundant, but play critical roles in sex determination during embryonic development or in reproductive trait development such as functional spermatogenesis and oocyte development, but these have yet to be measured. Another possibility is that these isoforms have functions in non-morphological sexual characters such as mating behavior. For example, it is known that the sex-determination cascade regulates expression and splice patterns of the gene fruitless (fru) in Drosophila . Coordinated actions of fru and dsx are necessary for male-specific neural development for adult male sexual behaviors [56, 57]. Future functional analyses of CmDsxA and CmDsxC focusing on other non-morphological traits will be needed to understand the dsx molecular evolution.
The transcriptome we generated for the prepupal development period in the stag beetle is a useful and important resource for identifying genes involved in morphological evolution including the regulation of sex determination and sexual dimorphism. The ability to identify and functionally screen entire suites of candidate genes, such as those in the sex determination pathway, is a powerful step toward a deeper understanding of both the diversification as well as the conservation of developmental mechanisms in the evolution of complex traits.
The golden metallic stag beetle Cyclommatus metallifer was purchased from the beetle pet store Hercules-Hercules in Sapporo, Japan. Adults were reared and bred in the laboratory as described previously . Induction of large and small males was also described in Gotoh et al. 2011. Briefly, we kept larvae individually starting from the second instar in 430 mL (large males) or 120 mL (small males) plastic cups filled with decaying wood flakes to induce large and small males.
Construction of the transcriptome database
Six mRNA libraries were constructed for sequencing from individuals sampled at three stages of larval development (3rd instar larval stage, early pre-pupal stage, and late pre-pupal stage). Libraries at each stage included heads from large males, prothoraces from large males, heads from small males, prothoraces from small males, heads from females, and prothoraces from females. In total, we sampled from 102 individuals evenly spread across the treatments.
Tissues were dissected and stored in RNALater before being transferred to TRiZOL® (Invitrogen, USA) and homogenized using a hand held tissue tearer and left overnight at −80 °C. RNA extraction was then carried out using TRIZoL following the manufacturer’s guidelines, followed by treatment with Ambion® DNA-free™ DNase, before assaying quantity with a NanoDrop (Thermo Scientific, USA) and quality with a BioAnalyzer 2100 (Agilent Inc., USA). For quality control, two aliquots of RNA were taken per extraction, one was left at 37 °C for two hours, while the other was stored at 4 °C. Both aliquots were run on an Agilent RNA nano-chip (Agilent Inc., USA) and the resulting electropherograms inspected for signs of degradation. If any degradation was observed, the extraction was discarded. It should be noted that the RIN value system  is not applicable to beetle species since, as in other insect species, the 28S ribosomal peak collapses into the 18S ribosomal peak .
Total RNA for each library was then pooled and three rounds of mRNA enrichment were carried out using the Oligotex® mRNA purification kit (Qiagen Inc., USA). Again quantity, quality, and confirmation of substantial mRNA enrichment were performed using a NanoDrop (Thermo Scientific, USA), and quality, with a BioAnalyzer 2100 (Agilent Inc., USA).
Samples were then taken to the Washington State University Molecular Biology Core Laboratory for sequencing library preparation and subsequent sequencing using the 454 GS FLX Titanium Series (Roche, USA), following manufacturer’s protocols. Briefly, mRNA was fragmented using ZnCl2 and heat prior to cDNA synthesis. Unique molecular identifiers were ligated to each library and titrated to achieve 8–10 % enrichment and the sequencing beads for each library were made in separate reactions. Sample balance was achieved by loading the picotiter plate with an equal number of enriched beads, determined by coulter counter, for each library.
Prior to analysis, basic statistics on the sequenced reads were obtained through scripts provided by Dr. David Rosenkranz (Johannes Gutenberg University, Mainz). All adapter nucleotides were trimmed and the 454 sequences were de novo assembled using the -cdna flag to signify transcriptome assembly in the Newbler 2.9 software package using default parameters (454 Life Sciences/Roche). The resulting isotigs were annotated using a BLASTx search  against the GenBank non-redundant database using an e-value threshold of e −5 to identify putative homology. Blast2GO [60, 61] was used to assign GO (Gene Ontology) terms and KEGG (Kyoto Encyclopedia of Genes and Genomes) based metabolic pathways [62, 63]. Final GO assignments were defined based on a 10 % filter for all three processes profiled at level 2. All other settings for the analysis were maintained at their defaults.
Database search against sex-determination pathway genes
The 24 candidate genes of the sex-determination pathway were selected from the genes categorized as “sex determination (BSID: 492283)” in BioSystems database at NCBI (http://www.ncbi.nlm.nih.gov/biosystems/) . All of those genes were identified in Drosophila melanogaster. We searched candidate genes against the database using the tBLASTn algorithm. We used the Drosophila and Tribolium homologs of those candidate genes (Table 2) as query sequences for the tBLASTn searches. For the Tribolium homologs, we used the annotated gene sequences from OrthoDB7 (http://cegg.unige.ch/orthodb7). To further confirm the orthology of detected genes, we made multiple alignments of candidate genes with the orthologs from other insect species, and constructed unrooted trees of mRNA sequences using ClustalX program  (http://www.clustal.org/) (Additional file 1: Figure S1-S14). We used the conserved 25 amino acid auto-regulation domain  from previously studied insect tra genes for constructing the NJ tree for tra, using the ClustalX program.
Identify full-length Cmdsx gene by RACE-PCR
Rapid amplification of cDNA ends (RACE) -PCR was performed to obtain the full length of undescribed Cmdsx transcripts sequence using the 3’-RACE primer (5’-CTC GAA GAT TGC CAT AAG CTC CTG GAA AGG-3’) and the SMART RACE cDNA Amplification Kit (Clontech, Palp Alto, CA). Also, we performed 5’-RACE for Cmtra2 and Cmtra using 5’-RACE primer (Cmtra2: 5’- GCG TAG GCG TGT GAG CCC TCT CTG TG -3’, Cmtra: 5’- CCG CAA CTT TGA TGG TTT TGA GTT CCT G -3’). The amplified cDNA fragments were subcloned and sequenced as described in Gotoh et al. (2014).
Expression analyses of candidate genes using RT-PCR
The expression patterns of our candidate genes in males and females were examined during the prepupal period using RT-PCR. Head, wings and legs were dissected from St-3 prepupae (prepupae which had just undergone the gut purge, see Gotoh et al. 2011 for a detailed definition of the prepupal stages) of each sex. Dissected tissues were preserved at −20 °C in RNA later (Ambion Inc, Austin, TX) until extraction. RNA was extracted from dissected tissues with the GeneJET RNA purification kit (Thermo Fisher Scientific Inc, Waltham, MA) according to the manufacturer’s protocol. For each sample, 2000 ng of total RNA was reverse transcribed with the iScript cDNA Synthesis kit (Bio-Rad, Hercules, CA). These synthesized cDNAs were used as templates for RT-PCR. See Additional file 2: Table S1 for primer sequences. The PCR step was performed under the following conditions: 94 °C for 2 min; 37 (for Cmix, CmSxl and Cmtra2) or 40 (Cmdsx) cycles of 94 °C for 30s, 54 °C for 30s and 72 °C for 3 min; and 72 °C for 10 min, using Go Taq Master Mix (Promega, Madison, WI). For Cmtra, the PCR program was: 94 °C for 2 min; 40 cycles of 94 °C for 30s, 54 °C for 30s and 72 °C for 4 min; and 72 °C for 10 min, using AmpliTaq360 DNA polymerase (Applied Biosystems). GAPDH was used as a positive control.
Functional analyses of candidate genes via RNAi
Functional analysis of our three target sex-determination genes (Cmix, CmSxl, Cmtra2) and each Cmdsx splice variant was accomplished with RNA interference (RNAi) during prepupal development. To silence candidate transcripts, double-stranded RNA (dsRNA) against either each target gene (Cmix, CmSxl, Cmtra2) or each target exon (Cmdsx exon 4, 6–7, 8 and 9) was synthesized. Using primers with the T7 sequence in the 5’-end, partial sequences of target sequences were amplified by PCR. Primer sequences are described in Additional file 3: Table S2. dsRNA synthesis were performed with MEGAscript RNAi Kit (Ambion, Austin, TX) according to the manufacturer’s protocol. For Cmdsx exon 1 and GFP, we used dsRNAs which were used in our previous study on Cmdsx function . All dsRNA was diluted with PBS. One μg (which is sufficient to strongly knock down gene expression in C. metallifer ) of dsRNA in 5 μl of PBS was injected into the dorsal prothorax of late 3rd instar larvae using a microliter syringe (Hamilton, Reno, NV) under a stereomicroscope. Individuals used for subsequent morphological analyses were those that successfully eclosed into adults. Sample sizes are described in Fig. 7. For statistical tests of RNAi effects on mandible size, analysis of covariance (ANCOVA) were performed with body size as a covariate using R 3.0.2 software.
We thank Dr. J. Shinji for experimental support for 5’ RACE of Cmtra2 gene. We also thank Dr. M. Lavine for his advice and critical comments on the manuscript. We would like to thank M. Wildung and D. Pouchnik for their advice on sequencing preparation methods. This work was supported by KAKENHI (No. 26650151) to T. Miura, Grant-in-Aid for Scientific Research on Innovative Areas “Genetic Bases for the Evolution of Complex Adaptive Traits”. (Grant Number: 25128706) to T. Niimi, NSF IOS 0642407 and USDA Hatch funds to L. Lavine, NSF IOS 0642409 to D. Emlen and NSF IOS 0920142 to I. Dworkin and a Grant for Basic Science Research Projects from the Sumitomo Foundation (No. 120165) to H. Gotoh. H. Gotoh was also supported by a Japanese Society for the Promotion of Sciences Research Fellowship for Young Scientists.
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- Darwin C. The Descent of Man. 1871. J. Murray.Google Scholar
- Andersson M. Sexual Selection. Princeton Univ. Press: Princeton; 1994.Google Scholar
- Emlen DJ, Nijhout HF. The development and evolution of exaggerated morphologies in insects. Annu Rev Entomol. 2000;45:661–708.View ArticlePubMedGoogle Scholar
- Panhuis TM, Butlin R, Zuk M, Tregenza T. Sexual selection and speciation. Trends Ecol Evol. 2001;16:364–71.View ArticlePubMedGoogle Scholar
- Ritchie MG. Sexual selection and speciation. Annu Rev Ecol Evol Syst. 2007;38:79–102.View ArticleGoogle Scholar
- Emlen DJ. The evolution of animal weapons. Annu Rev Ecol Evol Syst. 2008;39:387–413.View ArticleGoogle Scholar
- Kawano K. Sexual dimorphism and the making of oversized male characters in Beetles (Coleoptera). Ann Entomol Soc America. 2006;99:327–41.View ArticleGoogle Scholar
- Kodric-Brown AK, Sibly RM, Brown JH. The allometry of ornaments and weapons. Proc Natl Acad Sci U S A. 2006;103:8733–8.View ArticlePubMedPubMed CentralGoogle Scholar
- Moczek AP, Nijhout HF. Developmental mechanisms of threshold evolution in a polyphenic beetle. Evol Dev. 2002;4:252–64.View ArticlePubMedGoogle Scholar
- Emlen DJ, Marangelo J, Ball B, Cunningham CW. Diversity in the weapons of sexual selection: horn evolution in the beetle genus onthophagus (coleoptera: scarabaeidae). Evolution. 2005;59:1060–84.View ArticlePubMedGoogle Scholar
- Siva-Jothy MT. Mate securing tactics and the cost of fighting in the Japanese horned beetle, Allomyrina dichotoma L. (Scarabaeidae). J Ethol. 1987;5:165–72.View ArticleGoogle Scholar
- Eberhard WG. The functions of horns in Podischnus agenor Dynastinae and other beetles. In: Blum MS, Blum NA, editors. Sexual Selection and Reproductive Competition in Insects. New York: Academic; 1979. p. 231–58.Google Scholar
- Otronen M. Intra- and intersexual interactions at breeding burrows in the horned beetle, Coprophanaeus ensifer. Anim Behav. 1988;36:741–8.View ArticleGoogle Scholar
- Hongo Y. Evolution of male dimorphic allometry in a population of the Japanese horned beetle Trypoxylus dichotomus septentrionalis. Behav Ecol Sociobiol. 2007;62:245–53.View ArticleGoogle Scholar
- Moczek AP, Rose DJ. Differential recruitment of limb patterning genes during development and diversification of beetle horns. Proc Natl Acad Sci U S A. 2009;106:8992–7.View ArticlePubMedPubMed CentralGoogle Scholar
- Emlen DJ, Warren IA, Johns A, Dworkin I, Lavine LC. A mechanism of extreme growth and reliable signaling in sexually selected ornaments and weapons. Science. 2012;337:860–4.View ArticlePubMedGoogle Scholar
- Moczek AP, Kijimoto T. Development and evolution of insect polyphenisms: novel insights through the study of sex determination mechanisms. Curr Opin Insect Sci. 2014;1:52–8.View ArticleGoogle Scholar
- Gempe T, Beye M. Function and evolution of sex determination mechanisms, genes and pathways in insects. Bioessays. 2011;33:52–60.View ArticlePubMedPubMed CentralGoogle Scholar
- Bell LR, Maine EM, Schedl P, Cline TW. Sex-lethal, a Drosophila sex determination switch gene, exhibits sex-specific RNA splicing and sequence similarity to RNA binding proteins. Cell. 1988;55:1037–46.View ArticlePubMedGoogle Scholar
- Pane A, Salvemini M, Bovi PD, Polito C, Saccone G. The transformer gene in Ceratitis capitata provides a genetic basis for selecting and remembering the sexual fate. Development. 2002;129:3715–25.PubMedGoogle Scholar
- Saccone G, Salvemini M, Polito LC. The transformer gene of Ceratitis capitata: a paradigm for a conserved epigenetic master regulator of sex determination in insects. Genetica. 2011;1(139):99–111.View ArticleGoogle Scholar
- Kiuchi T, Koga H, Kawamoto M, Shoji K, Sakai H, Arai Y, et al. A single female-specific piRNA is the primary determiner of sex in the silkworm. Nature. 2014;509:633–6.View ArticlePubMedGoogle Scholar
- Verhulst EC, Beukeboom LW, Van de Zande L. Maternal control of haplodiploid sex determination in the wasp Nasonia. Science. 2010;328:620–3.View ArticlePubMedGoogle Scholar
- Hasselmann M, Gempe T, Schiøtt M, Nunes-Silva CG, Otte M, Beye M. Evidence for the evolutionary nascence of a novel sex determination pathway in honeybees. Nature. 2008;454:519–22.View ArticlePubMedGoogle Scholar
- Williams TM, Carroll SB. Genetic and molecular insights into the development and evolution of sexual dimorphism. Nat Rev Genet. 2009;10:797–804.View ArticlePubMedGoogle Scholar
- Shukla JN, Nagaraju J. Doublesex: a conserved downstream gene controlled by diverse upstream regulators. J Genet. 2010;89:341–56.View ArticlePubMedGoogle Scholar
- Kopp A. Dmrt genes in the development and evolution of sexual dimorphism. Trends Genet. 2012;28:175–84.View ArticlePubMedPubMed CentralGoogle Scholar
- Kijimoto T, Moczek AP, Andrews J. Diversification of doublesex function underlies morph-, sex-, and species-specific development of beetle horns. Proc Natl Acad Sci U S A. 2012;109:20526–31.View ArticlePubMedPubMed CentralGoogle Scholar
- Shukla JN, Palli SR. Doublesex target genes in the red flour beetle, Tribolium castaneum. Sci Rep. 2012b;2:00948.Google Scholar
- Ito Y, Harigai A, Nakata M, Hosoya T, Araya K, Oba Y, et al. The role of doublesex in the evolution of exaggerated horns in the Japanese rhinoceros beetle. EMBO Rep. 2013;14:561–7.View ArticlePubMedPubMed CentralGoogle Scholar
- Gotoh H, Miyakawa H, Ishikawa A, Ishikawa Y, Sugime Y, Emlen DJ, et al. Developmental link between sex and nutrition; doublesex regulates sex-specific mandible growth via juvenile hormone signaling in stag beetles. PLoS Genet. 2014;10(1):e1004098.View ArticlePubMedPubMed CentralGoogle Scholar
- Erickson JW, Quintero JJ. Indirect effects of ploidy suggest X chromosome dose, not the X: a ratio, signals sex in Drosophila. PLoS Biol. 2007;5:e332.View ArticlePubMedPubMed CentralGoogle Scholar
- Salz HK. Sex determination in insects: a binary decision based on alternative splicing. Curr Opin Genet Dev. 2011;21:395–400.View ArticlePubMedPubMed CentralGoogle Scholar
- Traut W, Niimi T, Ikeo K, Sahara K. Phylogeny of the sex-determining gene Sex-lethal in insects. Genome. 2006;49:254–62.View ArticlePubMedGoogle Scholar
- Waterbury JA, Jackson LL, Schedl P. Analysis of the doublesex female protein in Drosophila melanogaster: role in sexual differentiation and behavior and dependence on intersex. Genetics. 1999;152:1653–67.PubMedPubMed CentralGoogle Scholar
- Garrett-Engele CM, Siegal ML, Manoli DS, Williams BC, Li H, Baker BS. intersex, a gene required for female sexual development in Drosophila, is expressed in both sexes and functions together with doublesex to regulate terminal differentiation. Development. 2002;129:4661–75.PubMedGoogle Scholar
- Verhulst EC, van de Zande L, Beukeboom LW. Insect sex determination: it all evolves around transformer. Curr Opin Genet Dev. 2010;20:376–83.View ArticlePubMedGoogle Scholar
- Goralski TJ, Edström JE, Baker BS. The sex determination locus transformer-2 of Drosophila encodes a polypeptide with similarity to RNA binding proteins. Cell. 1989;56:1011–8.View ArticlePubMedGoogle Scholar
- Gotoh H, Cornette R, Koshikawa S, Okada Y, Lavine LC, Emlen DJ, et al. Juvenile hormone regulates extreme mandible growth in male stag beetles. PLoS ONE. 2011;6:e21139.View ArticlePubMedPubMed CentralGoogle Scholar
- Altschul SF, Madden TL, Schäffer AA, Zhang J, Zhang Z, et al. Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res. 1997;25:3389–402.View ArticlePubMedPubMed CentralGoogle Scholar
- Richards S, Gibbs RA, Weinstock GM, Brown SJ, Denell R, Beeman RW, et al. The genome of the model beetle and pest Tribolium castaneum. Nature. 2008;452:949–55.View ArticlePubMedGoogle Scholar
- Gilbert LI. Insect Molecular Biology and Biochemistry. Cambridge: Academic; 2012.Google Scholar
- Clough E, Oliver B. Genomics of sex determination in Drosophila. Brief Funct Genomics. 2012;11:387–94.View ArticlePubMedPubMed CentralGoogle Scholar
- Mullon C, Pomiankowski A, Reuter M. Molecular evolution of Drosophila Sex-lethal and related sex determining genes. BMC Evol Biol. 2012;12:5.View ArticlePubMedPubMed CentralGoogle Scholar
- Nagengast AA, Stitzinger SM, Tseng CH, Mount SM, Salz HK. Sex-lethal splicing autoregulation in vivo: interactions between SEX-LETHAL, the U1 snRNP and U2AF underlie male exon skipping. Development. 2003;130:463–71.View ArticlePubMedGoogle Scholar
- Cline TW. Two closely linked mutations in Drosophila melanogaster that are lethal to opposite sexes and interact with daughterless. Genetics. 1978;90:683–97.PubMedPubMed CentralGoogle Scholar
- Zhang Z, Klein J, Nei M. Evolution of the Sex-lethal gene in insects and origin of the Sex-determination system in Drosophila. J Mol Evol. 2014;78:50–65.View ArticlePubMedGoogle Scholar
- Burtis KC, Baker BS. Drosophila doublesex gene controls somatic sexual differentiation by producing alternatively spliced mRNAs encoding related sex-specific polypeptides. Cell. 1989;56:997–1010.View ArticlePubMedGoogle Scholar
- Tian M, Maniatis T. A splicing enhancer complex controls alternative splicing of doublesex pre-mRNA. Cell. 1993;74:105–14.View ArticlePubMedGoogle Scholar
- Nissen I, Müller M, Beye M. The Am-tra2 gene is an essential regulator of female splice regulation at two levels of the sex determination hierarchy of the honeybee. Genetics. 2012;192:1015–26.View ArticlePubMedPubMed CentralGoogle Scholar
- Shukla JN, Palli SR. Tribolium castaneum Transformer-2 regulates sex determination and development in both males and females. Insect Biochem Mol Biol. 2013;43:1125–32.View ArticlePubMedPubMed CentralGoogle Scholar
- Shukla JN, Palli SR. Sex determination in beetles: production of all male progeny by parental RNAi knockdown of transformer. Sci Rep. 2012;2.Google Scholar
- Chase BA, Baker BS. A genetic analysis of intersex, a gene regulating sexual differentiation in Drosophila melanogaster females. Genetics. 1995;139:1649–61.PubMedPubMed CentralGoogle Scholar
- Aspiras AC, Smith FW, Angelini DR. Sex-specific gene interactions in the patterning of insect genitalia. Dev Biol. 2011;360:369–80.View ArticlePubMedGoogle Scholar
- Ryner LC, Goodwin SF, Castrillon DH, Anand A, Villella A, Baker BS, et al. Control of male sexual behavior and sexual orientation in Drosophila by the fruitless gene. Cell. 1996;87:1079–89.View ArticlePubMedGoogle Scholar
- Kimura KI, Hachiya T, Koganezawa M, Tazawa T, Yamamoto D. Fruitless and doublesex coordinate to generate male-specific neurons that can initiate courtship. Neuron. 2008;59:759–69.View ArticlePubMedGoogle Scholar
- Rideout EJ, Dornan AJ, Neville MC, Eadie S, Goodwin SF. Control of sexual differentiation and behavior by the doublesex gene in Drosophila melanogaster. Nature Neurosci. 2000;13:458–66.View ArticleGoogle Scholar
- Schroeder A, Mueller O, Stocker S, Salowsky R, Leiber M, et al. The RIN: an RNA integrity number for assigning integrity values to RNA measurements. BMC Mol Biol. 2006;7:3.View ArticlePubMedPubMed CentralGoogle Scholar
- Winnebeck EC, Millar CD, Warman GR. Why does insect RNA look degraded? J Insect Sci. 2010;10:159.View ArticlePubMedPubMed CentralGoogle Scholar
- Conesa A, Götz S, García-Gómez JM, Terol J, Talón M, et al. Blast2GO: a universal tool for annotation, visualization and analysis in functional genomics research. Bioinformatics. 2005;21:3674–6.View ArticlePubMedGoogle Scholar
- Götz S, García-Gómez JM, Terol J, Williams TD, Nagaraj SH, et al. High-throughput functional annotation and data mining with the Blast2GO suite. Nucleic Acids Res. 2008;36:3420–35.View ArticlePubMedPubMed CentralGoogle Scholar
- Kanehisa M, Goto S. KEGG: kyoto encyclopedia of genes and genomes. Nucleic Acids Res. 2000;28:27–30.View ArticlePubMedPubMed CentralGoogle Scholar
- Kanehisa M, Araki M, Goto S, Hattori M, Hirakawa M, et al. KEGG for linking genomes to life and the environment. Nucleic Acids Res. 2008;36:D480–4.View ArticlePubMedPubMed CentralGoogle Scholar
- Geer LY, Marchler-Bauer A, Geer RC, Han L, He J, He S et al. The NCBI biosystems database. Nucleic Acids Res. 2010;38:492–96.Google Scholar
- Larkin MA, Blackshields G, Brown NP, Chenna R, McGettigan PA, McWilliam H, et al. Clustal W and Clustal X version 2.0. Bioinformatics. 2007;23:2947–8.View ArticlePubMedGoogle Scholar