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Identification and functional analyses of sex determination genes in the sexually dimorphic stag beetle Cyclommatus metallifer



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 [16]. 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 [68]. 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. [1114]), 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 [1517]. Because of their sex-specific expression, the sex determination genes of insects, which are highly conserved across the arthropods and animals in general [18], are particularly important to understanding the development of sexually selected weapon traits [17].

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 [22]). Diversity among species is also found in the maternal transfer of tra mRNA in Nasonia [23] and heterozygosity in the complementary sex determiner (csd) gene’s activation of the gene feminizer (fem) in honeybees [24]. 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 [18], 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, 2527].

Recently several authors, including our group, have shown that dsx regulates sex-specific trait expression in beetles, including species with horns and elongated mandibles [2831]. 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.

In this study, we tested the role of several genes in the sex determination pathway known to regulate sex-specific trait expression in other animals in the sexually dimorphic and male dimorphic golden metallic stag beetle Cyclommatus metallifer (Fig. 1). This species (and many other species of Lucanidae) possess extremely elongated mandibles, which are expressed in a sex and condition dependent manner [31]. We constructed a transcriptome for the critical prepupal period of development when all adult structures are determined and differentiating in C. metallifer in order to rigorously test the hypothesis that the effector genes for sexual differentiation are conserved with other insects. We queried this transcriptome database for 24 sex-determination cascade genes known in Drosophila, and identified 14 putative homologs in C. metallifer. From these 14 sex-determination cascade gene homologs, we selected three central sex-determination cascade genes for further analyses; tra, tra2, and ix, which have significant roles during sexual differentiation in many insects. Also, we analyzed Sxl, which is recruited as a key upstream switch for sex determination in the Drosophila lineage.

Fig. 1
figure 1

The golden metallic stag beetle, Cyclommatus metallifer. Female (left) and large male (right) are shown. Only males possess enlarged mandibles, which are used for male-male competition over resources. Scale bar indicates 20 mm

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 [33]. 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 [37] 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

We performed Roche 454-pyrosequencing of an mRNA-enriched RNA library prepared from head and prothorax tissue of C. metallifer larvae and prepupae - three each of small and large individuals of both sexes. This stage was chosen as the most likely developmental period when differentiation of adult structures is occurring [31, 39]. A total of 1,223,516 raw reads consisting of 391,313,087 bp were sequenced. Raw sequence data was submitted to NCBI’s Sequence Read Archive (SRA) under the Bioproject Accession number PRJNA311137. Assembled and annotated transcriptome files available upon request. After adapter trimming and quality filtering, the reads were assembled into a de novo transcriptome using Newbler 2.9. This transcriptome produced 17,894 contigs, incorporating 353,508,757 base pairs and 1,106,461 reads. 10,468 contigs were larger than 500 bp (labeled as “large contigs” by the Newbler assembler, Fig. 2a, Table 1). The average “large” contig size was 1,267 bp, with a max contig size of 16,848 bp (Fig. 2a). The average read depth was 87.84, and the N50 was 1479 (Table 1). The assembled contigs were used to generate putative isoform variants, or isotigs. 14,565 isotigs were generated, with an average size of 1,592 bp (Fig. 2b). The average number of contigs per isotig was 2.1, with 8,863 isotigs deriving from single contigs (Fig. 2c). The isotigs were grouped into 10,793 unique isogroups (or an estimate of the total number of genes expressed in C. metallifer across the tissue libraries). A total of 8,933 isogroups only contained a single isotig, with an average of 1.4 isotigs per isogroup (Fig. 2d).

Fig. 2
figure 2

Summary of Cyclommatus metallifer transcriptome assembly. a Length distribution of contig sequences. X-axis indicates sequence length of contig and Y-axis indicates number of contigs. b Length distribution of isotig sequences. X-axis indicates sequence length of isotig and Y-axis indicates number of isotigs (c) Number of contigs used in the assembly of individual isotigs. X-axis indicates number of contigs per each isotig and Y-axis indicates number of isotigs (d) Number of isotigs used in the assembly of individual isogroups. X-axis indicates number of isotigs per each isogroup and Y-axis indicates number of isogroups

Table 1 Newbler 2.9 assmbly metrics

After assembly, we performed BLASTx searches of the isotigs against the NCBI non-redundant protein database [40], using an e-value cut-off of e−5. After the BLASTx search, isotigs from each isogroup were filtered by best BLAST hit. If no members of an isogroup had a BLAST hit, one isotig from that isogroup was randomly selected to represent that isogroup. This left a total of 10,772 isotigs. 8,774 isotigs (78.38 %) had significant homology with proteins in the database. Many of them (49.2 %) have e-values < e−100 (Fig. 3a). Most of the isotigs (80.77 %) had the highest similarity with sequences of red flour beetle Tribolium castaneum (Fig. 3b). Considering their phylogenetic closeness (both Cyclommatus and Tribolium are in the order Coleoptera) and the completeness of Tribolium genome sequence and its annotation, this result is expected. Gene Ontology (GO) terms were assigned to each isotig using Blast2GO (Fig. 3c-e).

Fig. 3
figure 3

Results of BLASTx search and classification of sequences based on GO term analyses. a Distribution of BLASTx top-hits against NCBI non-redundant protein database. E-value cutoff of e-5 was used as threshold. Results with an e-value < e-100 were clustered together. b Distribution of BLASTx results, clustered by species of origin. BLASTx hits that returned an “unknown” species were removed from the analysis. c-e Gene Ontology isotig groups for Biological Process (c), Molecular Function (d), and Cellular Component (e)

Identification of sex-determination cascade genes

Using homologous gene sequences of Drosophila and Tribolium as our query terms (Table 2), we searched for 24 candidate genes from the sex-determination cascade (Table 2) against our C. metallifer database using BLASTx. A total of 14 C. metallifer genes were identified as homologous with known sex determination genes (Table 2) based on e-value (threshold < e−40) and/or gene phylogenies (Additional file 1: Figure S1-S14).

Table 2 Summary of database search for candidate genes

Notably, the isogroup corresponding to Cmtra2 contained nine different isotigs, indicating that Cmtra2 potentially has multiple different splice variants (Fig. 4). Since three splice variants corresponding to isotig 01274, 01278 and 01279 were predicted to lack 5’ sequences, we additionally performed 5’ RACE to identify 5’ sequences of these splice variants. Three of these variants also shared exons 1 and 3 with isotigs 01272, 01276 and 01277. At the protein level, five different putative protein products (isoforms) were encoded (Fig. 4; labeled Isoforms A to E). Isoform A, B and C are similar to each other, that is, they share conserved regions. Differences among the isoforms are only detected in the last several amino acid residues at the 3’ends (Fig. 4). Isoform D and E are shorter than the other three, due to the insertion of stop codons on exon 2 and exon 4. In addition to Cmtra2, two isotigs corresponded to the female lethal d (fl(2)d) homolog,suggesting the existence of splice variants (data not shown).

Fig. 4
figure 4

Predicted gene models of the C. metallifer transformer-2 (Cmtra2) gene. Nine different splice variants were predicted from the database assembly and annotation. A total of five different isoforms were predicted (isoform A-E). The coding sequences are in gray, stop codons are indicated by black arrowheads. The open arrowhead indicates the start codon. Arrows indicate forward and reverse primer locations for RT-PCR and the red bar indicates the region for dsRNA synthesis. The blue dashed box indicates the sequences which were identified by 5’ RACE

Additionally, we found tra-like sequence in our database (isotig 04317) by using the Tribolium tra homolog as a query, although the homology was relatively low (e-value = 2e−8). Expression analyses and subsequent sequencing revealed that gene transcripts and the corresponding putative protein products were different between sexes (Fig. 5a, b). The male-specific isoform was shorter than the female-specific isoform due to an insertion of a stop codon in exon 2 (Fig. 5a). The female-specific isoform possessed a putative auto-regulation domain, which is contained in all tra genes identified in other insects except Drosophila [37]. This auto-regulation domain was only partially conserved in the male-specific isoform (Fig. 5a, c). Phylogenetic analyses identifying female Tra proteins in other insects indicated that this Tra-like putative protein was grouped with Tribolium Tra (Fig. 5d). Thus, we concluded that the tra-like isotig 04317 is the Cyclommatus tra homolog and named it Cmtra.

Fig. 5
figure 5

Gene annotation of isotig 04317. a Predicted gene model of isotig 04317 (Cmtra), and expressed transcripts in females and males. The coding sequences are in gray, stop codons are indicated by black arrowheads. The open arrowhead indicates the start codon. Arrows indicate forward and reverse primer locations for RT-PCR (b) Expression patterns of Cmtra in females and males. Template cDNAs were derived from Stage 3 prepupae. H: head, L: leg, W: wing. GAPDH was used as a positive control. (c) Alignment of conserved 25 amino acid sequences of the putative auto regulation domain [37] of the Tra protein. d NJ tree of Tra protein from the conserved 25 amino acid auto-regulation domain

Expression analyses of CmSxl, Cmix, and Cmtra2

From the 14 candidate C. metallifer sex determination genes (Table 2), we selected CmSxl, Cmix, and Cmtra2 for expression and functional analyses because these three genes have well known functions in the sex determination cascade in Drosophila. In order to describe the expression patterns of those candidate genes, we conducted semi-quantitative RT-PCR between the sexes and across body parts (head, legs, wings) during the prepupal period. Our results showed uniform expression of CmSxl between sexes and among body parts (Fig. 6a). The expression pattern of Cmix was also neither sex nor body-parts specific (Fig. 6a). Since there were several assembled isotigs homologous to Cmtra2 indicating existence of multiple splice variants, we used multiple primers to investigate the expression of specific Cmtra2 splice variants. Only a single product was detected by RT-PCR using the F1-R1 primer set (Fig. 6b). According to its size, this amplified product is probably composed of exon 1, 3 and 5. Thus, isotig 01272 (isoform A), 01276 (isoform B) and 01277 (isoform C) appear to be the most commonly expressed forms. Products of the isotigs corresponding to isoform D and E were not amplified to a detectable level. In addition, RT-PCR using F2-R2 primers also amplified only a single product with 370 bp predicted size (Fig. 6b), suggesting that isotig 01276 (corresponding to isoform B) had higher expression than isotig 01272 (corresponding to isoform A), and that the major expressed variants are isotig 01276 and/or 01277. We could not find any evidence of sex specific or organ specific expression of Cmtra2, regardless of the primer sets used for RT-PCR.

Fig. 6
figure 6

Expression patterns of CmSxl, Cmix and Cmtra2. Template cDNAs were derived from Stage 3 prepupae. H: head, L: leg, W: wing. a Expression patterns of Cmix and CmSxl. Only a single product was amplified regardless of sex or tissue type. b Expression patterns of Cmtra2 using different pairs of primers. See Fig. 5 for their position. Only a single product was amplified regardless of sex or tissue type. GAPDH was used as positive control for both analyses

Functional analyses of Sxl, ix and tra2

We performed gene knockdowns via RNAi during the prepupal period, which is the critical period of sex-specific mandibular tissue proliferation in the stag beetle [31, 39]. Knockdown of Cmtra2 resulted in death in both sexes (0/27 females and 0/5 males survived); all of the dsRNA injected larvae died during the prepupal period. Knockdown of CmSxl did not show any detectable abnormalities in morphology in either sex (Fig. 7). In contrast to these two early acting genes, knockdown of Cmix significantly affected sex-specific morphological characters such as mandible size and body color in females (Fig. 7), as predicted from its role in terminal differentiation in Drosophila. Specifically, mandible growth was induced in Cmix RNAi females compared with control GFP RNAi females (Fig. 7). Those Cmix RNAi female phenotypes exhibited phenotypes identical to the Cmdsx RNAi female phenotypes across all affected traits (Fig. 7). Also, dark non-metallic brown female body color was changed to be more male-like golden metallic color in Cmix RNAi females (Fig. 7). In contrast, Cmix RNAi males did not show any detectable effects (Fig. 7).

Fig. 7
figure 7

RNAi phenotypes of Cmdsx isoforms, Cmix and CmSxl. Each scatter plot panel indicates the scaling relationship between prothorax width (X-axis) and mandible length (Y-axis). GFP RNAi control individuals (closed circles, male: blue, female: pink) and Cmdsx-exon 1 RNAi (open diamonds, male: blue, female: pink) are indicated in all panels. Focal gene/exon RNAi individuals are indicated in closed triangles (male: red, female: light blue). The name of the focal genes or exons for each panel are indicated in the left row. N indicates the sample size for each gene/exon RNAi. Adult phenotypes of each gene knockdown are indicated on the right of each scatter plot panel. Scale bars indicate 20 mm for males and 10 mm for females

Identification of undescribed dsx variants by RACE

Although we were unable to locate a dsx homolog sequence in our database, we identified Cmdsx sequences by degenerate PCR and subsequent RACE-PCR in a previous study [31]. In Gotoh et al. (2014), we described the sex specific expression pattern and function of four Cmdsx splice variants [31]. However, other recent reports of dsx genes from other beetle species identified a larger number of dsx splice variants than we reported (i.e. - seven in Onthophagus taurus [28]; and 11 in Trypoxylus dichotomus [30]), thus we tried to clone potential unidentified Cmdsx splice variants by RACE-PCR. We identified 16 additional splice variants (Fig. 8). We discovered five new exons at the 3’ end of our previously described variants. These five exons have variation in their combination, which leads to variation in the 3’ UTR among the splice variants (Fig. 8). Our new sequence information also caused us to divide and rename exons 8 and 9 to exon 8a, 8b and exon 9a, 9b. In females, the specific variants of Cmdsx C6, C8, D6, and D8 lack exon 8b, 9a and 9b, and variants C7 and D7 lack exon 9b. All of the variants including newly identified exons appeared as faint minor bands during RACE-PCR, while previously identified variants appeared as strong, clear bands (data not shown). Although we found these variations in 3’ UTR length among our 16 splice variants, the number of putative protein products (or the number of CmDsx isoforms) is still four (Fig. 8). We performed semi-quantitative RT-PCR using exon-specific primers (primer positions are indicated in Fig. 8) to get a profile of expression patterns for these Cmdsx splice variants during development. RT-PCR expression analyses showed that all of exons 9b, 12 and 14 were used in both males and females, although each variant had sex-specific patterns of expression (Fig. 9). Some of variants were not detected by RT-PCR. For example DNA bands corresponding to C8 and D8 were not amplified in RT-PCR using F1-R2 primer pair (Fig. 8), probably due to low expression levels.

Fig. 8
figure 8

Predicted gene models for Cmdsx gene. In addition to the previously reported four splice variants (corresponding to isoforms A to D), we identified 16 new splice variants of dsx in Cyclommatus metallifer. We updated the exon number from those reported in Gotoh et al. (2014). Exon 8 and exon 9 were divided into 8a, 8b and 9a, 9b, respectively (green bars and numbers). Also, five new exons were named (exon 10 to exon 14; red bars and numbers). Coding sequences are indicated in gray, arrows indicate forward and reverse primer locations for RT-PCR

Fig. 9
figure 9

Expression patterns of Cmdsx using different pairs of primers. Template cDNAs were derived from Stage 3 prepupae. H: head, L: leg, W: wing. See Fig. 8 for primer positions. Clear differential usages of Cmdsx variants between sexes are recognized. GAPDH was used as positive control for analysis

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 [31]. 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 [41]) and other holometabolous insect genomes (approximately 10,000 – 19,000 genes [42]). 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 [45]. 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 [47]. 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 [34] 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 [50] and Tribolium castaneum [51]. 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 [51] and the honeybee Apis mellifera [50]. 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 [51]. 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 [37]. 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 [53]. Knockdown of the ix gene in the milkweed bug Oncopeltus fasciatus affected both male and female genital morphological structures [54]. 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; [31]). 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 [31]. 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 [30]. However, the T. dichotomus DsxF-Short specific knockdown did not cause any detectable deformation phenotype in either sex [30]. 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 [28]. 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 [55]. 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.


Beetle rearing

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 [39]. 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 [58] is not applicable to beetle species since, as in other insect species, the 28S ribosomal peak collapses into the 18S ribosomal peak [59].

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.

Bioinformatics analysis

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 [40] 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 ( [64]. 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 ( 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 [65] ( (Additional file 1: Figure S1-S14). We used the conserved 25 amino acid auto-regulation domain [32] 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 [31]. All dsRNA was diluted with PBS. One μg (which is sufficient to strongly knock down gene expression in C. metallifer [31]) 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.


  1. Darwin C. The Descent of Man. 1871. J. Murray.

    Google Scholar 

  2. Andersson M. Sexual Selection. Princeton Univ. Press: Princeton; 1994.

    Google Scholar 

  3. Emlen DJ, Nijhout HF. The development and evolution of exaggerated morphologies in insects. Annu Rev Entomol. 2000;45:661–708.

    Article  CAS  PubMed  Google Scholar 

  4. Panhuis TM, Butlin R, Zuk M, Tregenza T. Sexual selection and speciation. Trends Ecol Evol. 2001;16:364–71.

    Article  PubMed  Google Scholar 

  5. Ritchie MG. Sexual selection and speciation. Annu Rev Ecol Evol Syst. 2007;38:79–102.

    Article  Google Scholar 

  6. Emlen DJ. The evolution of animal weapons. Annu Rev Ecol Evol Syst. 2008;39:387–413.

    Article  Google Scholar 

  7. Kawano K. Sexual dimorphism and the making of oversized male characters in Beetles (Coleoptera). Ann Entomol Soc America. 2006;99:327–41.

    Article  Google Scholar 

  8. Kodric-Brown AK, Sibly RM, Brown JH. The allometry of ornaments and weapons. Proc Natl Acad Sci U S A. 2006;103:8733–8.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  9. Moczek AP, Nijhout HF. Developmental mechanisms of threshold evolution in a polyphenic beetle. Evol Dev. 2002;4:252–64.

    Article  PubMed  Google Scholar 

  10. 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.

    Article  CAS  PubMed  Google Scholar 

  11. 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.

    Article  Google Scholar 

  12. 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 

  13. Otronen M. Intra- and intersexual interactions at breeding burrows in the horned beetle, Coprophanaeus ensifer. Anim Behav. 1988;36:741–8.

    Article  Google Scholar 

  14. 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.

    Article  Google Scholar 

  15. 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.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  16. 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.

    Article  CAS  PubMed  Google Scholar 

  17. 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.

    Article  Google Scholar 

  18. Gempe T, Beye M. Function and evolution of sex determination mechanisms, genes and pathways in insects. Bioessays. 2011;33:52–60.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  19. 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.

    Article  CAS  PubMed  Google Scholar 

  20. 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.

    CAS  PubMed  Google Scholar 

  21. 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.

    Article  Google Scholar 

  22. 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.

    Article  CAS  PubMed  Google Scholar 

  23. Verhulst EC, Beukeboom LW, Van de Zande L. Maternal control of haplodiploid sex determination in the wasp Nasonia. Science. 2010;328:620–3.

    Article  CAS  PubMed  Google Scholar 

  24. 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.

    Article  CAS  PubMed  Google Scholar 

  25. Williams TM, Carroll SB. Genetic and molecular insights into the development and evolution of sexual dimorphism. Nat Rev Genet. 2009;10:797–804.

    Article  CAS  PubMed  Google Scholar 

  26. Shukla JN, Nagaraju J. Doublesex: a conserved downstream gene controlled by diverse upstream regulators. J Genet. 2010;89:341–56.

    Article  CAS  PubMed  Google Scholar 

  27. Kopp A. Dmrt genes in the development and evolution of sexual dimorphism. Trends Genet. 2012;28:175–84.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  28. 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.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  29. Shukla JN, Palli SR. Doublesex target genes in the red flour beetle, Tribolium castaneum. Sci Rep. 2012b;2:00948.

  30. 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.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  31. 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.

    Article  PubMed  PubMed Central  Google Scholar 

  32. 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.

    Article  PubMed  PubMed Central  Google Scholar 

  33. Salz HK. Sex determination in insects: a binary decision based on alternative splicing. Curr Opin Genet Dev. 2011;21:395–400.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  34. Traut W, Niimi T, Ikeo K, Sahara K. Phylogeny of the sex-determining gene Sex-lethal in insects. Genome. 2006;49:254–62.

    Article  CAS  PubMed  Google Scholar 

  35. 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.

    CAS  PubMed  PubMed Central  Google Scholar 

  36. 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.

    CAS  PubMed  Google Scholar 

  37. Verhulst EC, van de Zande L, Beukeboom LW. Insect sex determination: it all evolves around transformer. Curr Opin Genet Dev. 2010;20:376–83.

    Article  CAS  PubMed  Google Scholar 

  38. 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.

    Article  CAS  PubMed  Google Scholar 

  39. 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.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  40. 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.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  41. 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.

    Article  CAS  PubMed  Google Scholar 

  42. Gilbert LI. Insect Molecular Biology and Biochemistry. Cambridge: Academic; 2012.

    Google Scholar 

  43. Clough E, Oliver B. Genomics of sex determination in Drosophila. Brief Funct Genomics. 2012;11:387–94.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  44. Mullon C, Pomiankowski A, Reuter M. Molecular evolution of Drosophila Sex-lethal and related sex determining genes. BMC Evol Biol. 2012;12:5.

    Article  PubMed  PubMed Central  Google Scholar 

  45. 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.

    Article  CAS  PubMed  Google Scholar 

  46. Cline TW. Two closely linked mutations in Drosophila melanogaster that are lethal to opposite sexes and interact with daughterless. Genetics. 1978;90:683–97.

    CAS  PubMed  PubMed Central  Google Scholar 

  47. 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.

    Article  CAS  PubMed  Google Scholar 

  48. 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.

    Article  CAS  PubMed  Google Scholar 

  49. Tian M, Maniatis T. A splicing enhancer complex controls alternative splicing of doublesex pre-mRNA. Cell. 1993;74:105–14.

    Article  CAS  PubMed  Google Scholar 

  50. 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.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  51. 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.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  52. Shukla JN, Palli SR. Sex determination in beetles: production of all male progeny by parental RNAi knockdown of transformer. Sci Rep. 2012;2.

  53. Chase BA, Baker BS. A genetic analysis of intersex, a gene regulating sexual differentiation in Drosophila melanogaster females. Genetics. 1995;139:1649–61.

    CAS  PubMed  PubMed Central  Google Scholar 

  54. Aspiras AC, Smith FW, Angelini DR. Sex-specific gene interactions in the patterning of insect genitalia. Dev Biol. 2011;360:369–80.

    Article  CAS  PubMed  Google Scholar 

  55. 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.

    Article  CAS  PubMed  Google Scholar 

  56. 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.

    Article  CAS  PubMed  Google Scholar 

  57. 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.

    Article  Google Scholar 

  58. 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.

    Article  PubMed  PubMed Central  Google Scholar 

  59. Winnebeck EC, Millar CD, Warman GR. Why does insect RNA look degraded? J Insect Sci. 2010;10:159.

    Article  PubMed  PubMed Central  Google Scholar 

  60. 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.

    Article  CAS  PubMed  Google Scholar 

  61. 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.

    Article  PubMed  PubMed Central  Google Scholar 

  62. Kanehisa M, Goto S. KEGG: kyoto encyclopedia of genes and genomes. Nucleic Acids Res. 2000;28:27–30.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  63. 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.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  64. 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.

  65. 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.

    Article  CAS  PubMed  Google Scholar 

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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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Correspondence to Laura C. Lavine.

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Authors’ contributions

Conceived and designed the experiments: HG, RAZ, IW, ID, DJE, LCL. Performed the experiments: HG, RAZ, IW. Analyzed the data: HG, RAZ, IW, MD, TN, ID, DJE, TM, LCL. Contributed reagents/materials/analysis tools: HG, RAZ, IW, MD, TN, ID, DJE, TM, LCL. Wrote the manuscript: HG, RAZ, IW, MD, TN, ID, DJE, TM, LCL. All authors have read and approved the manuscript.

Additional files

Additional file 1: Figure S1-S14.

Constructed unrooted phylogenetic trees of candidate sex-determination genes using ClustalX program. (DOCX 3072 kb)

Additional file 2: Table S1.

List of primer sequences for RT-PCR. (XLS 18 kb)

Additional file 3: Table S2.

List of primer sequences for dsRNA synthesis. (XLS 18 kb)

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Gotoh, H., Zinna, R.A., Warren, I. et al. Identification and functional analyses of sex determination genes in the sexually dimorphic stag beetle Cyclommatus metallifer . BMC Genomics 17, 250 (2016).

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