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Positive selection and comparative molecular evolution of reproductive proteins from New Zealand tree weta (Orthoptera, Hemideina)

  • Victoria G. Twort ,

    Roles Conceptualization, Formal analysis, Investigation, Writing – original draft, Writing – review & editing

    Current address: Department of Biology, Lund University, Lund, Sweden

    Affiliations School of Biological Sciences, The University of Auckland, Auckland, New Zealand, Landcare Research, Auckland, New Zealand


  • Alice B. Dennis,

    Roles Formal analysis, Investigation, Writing – review & editing

    Current address: Evolutionary Biology & Systematic Zoology, Institute for Biochemistry & Biology, University of Potsdam, Potsdam, Germany

    Affiliation Landcare Research, Auckland, New Zealand

  • Duckchul Park,

    Roles Investigation, Writing – review & editing

    Affiliation Landcare Research, Auckland, New Zealand

  • Kathryn F. Lomas,

    Roles Investigation, Writing – review & editing

    Affiliation CSIRO, Melbourne, Australia

  • Richard D. Newcomb,

    Roles Conceptualization, Supervision, Writing – review & editing

    Affiliations School of Biological Sciences, The University of Auckland, Auckland, New Zealand, The New Zealand Institute for Plant and Food Research Ltd, Auckland, New Zealand

  • Thomas R. Buckley

    Roles Conceptualization, Supervision, Writing – review & editing

    Affiliations School of Biological Sciences, The University of Auckland, Auckland, New Zealand, Landcare Research, Auckland, New Zealand

Positive selection and comparative molecular evolution of reproductive proteins from New Zealand tree weta (Orthoptera, Hemideina)

  • Victoria G. Twort, 
  • Alice B. Dennis, 
  • Duckchul Park, 
  • Kathryn F. Lomas, 
  • Richard D. Newcomb, 
  • Thomas R. Buckley


Animal reproductive proteins, especially those in the seminal fluid, have been shown to have higher levels of divergence than non-reproductive proteins and are often evolving adaptively. Seminal fluid proteins have been implicated in the formation of reproductive barriers between diverging lineages, and hence represent interesting candidates underlying speciation. RNA-seq was used to generate the first male reproductive transcriptome for the New Zealand tree weta species Hemideina thoracica and H. crassidens. We identified 865 putative reproductive associated proteins across both species, encompassing a diverse range of functional classes. Candidate gene sequencing of nine genes across three Hemideina, and two Deinacrida species suggests that H. thoracica has the highest levels of intraspecific genetic diversity. Non-monophyly was observed in the majority of sequenced genes indicating that either gene flow may be occurring between the species, or that reciprocal monophyly at these loci has yet to be attained. Evidence for positive selection was found for one lectin-related reproductive protein, with an overall omega of 7.65 and one site in particular being under strong positive selection. This candidate gene represents the first step in the identification of proteins underlying the evolutionary basis of weta reproduction and speciation.


Reproductive associated proteins have been shown to have increased evolutionary rates, and diverge rapidly between related taxa [16]. In particular, proteins present in the seminal fluid (SFPs) have been identified as often evolving under positive selection [4, 79]. In insects, the synthesis and secretion of SFPs occurs within male reproductive tract secretory tissues, such as accessory glands and testis [10, 11]. SFPs encompass a diverse range of functional classes and are involved in the modulation or induction of post mating responses in females [1218]. In addition, SFPs have been identified as playing a key role in reproductive isolation between diverging lineages [2, 1922].

Many Drosophila SFPs show increased evolutionary rates when compared to non-seminal proteins [1, 23, 24] and show evidence of positive selection [79, 25, 26]. However, not all SFPs exhibit rapid evolution; some show signs of evolutionary conservation [2730], while others exhibit both conservation and rapid evolution in different regions of a single protein [3133]. Drosophila SFPs have been shown to play a key role in reproductive isolation through species-specific gamete use, whereby SFPs need to have a specific structure and binding affinity for successful reproduction to occur [34]. Studies of orthopteran taxa have revealed similar patterns. In particular, studies on crickets have shown that SFPs have a higher level of divergence when compared with non-seminal proteins, with a significant proportion being under positive selection [7, 8, 25, 35]. Despite these studies in crickets, very little is known about SFP evolution in other orthopteran taxa.

Insects from the orthopteran family Anostostomatidae are collectively known in New Zealand by their Māori name, weta, and represent an important component of the native forest ecosystem [36]. All Tree (Hemideina) and Giant (Deinacrida) weta species are endemic to New Zealand, and include both relatively widespread and threatened species. The larger Deinacrida species have limited distributions and abundance, with 10 of the 11 species being under conservation management [3740]. There are seven species of Hemideina distributed throughout New Zealand, most of which are abundant [3741]. Hemideina crassidens (Blanchard) has the largest distribution of all weta species, with populations distributed in the south of the North Island, as well as in the north and west coast of the South Island in New Zealand [42]. Hemideina crassidens has two chromosomal races (15 and 19) that are morphologically identical and successfully produce offspring in laboratory crosses [43, 44]. Hemideina thoracica (White) is found in the upper three quarters of the North Island [39]. All populations are morphologically similar despite there being eight chromosomal races with diploid chromosome numbers ranging from 11 to 24 [45]. Individual populations have been shown to exhibit only a single karyotype, with interbreeding occurring in the narrow regions of contact [4547]. The presence of multiple chromosomal races within these species indicates that chromosomal differences are insufficient to lead to reproductive isolation [45, 48]. Hemideina trewicki has the smallest distribution of the three species, occurring only in southern and central Hawke’s Bay. In the northern parts of the range, H. trewicki is sympatric with H. thoracica [42, 49] and has one known chromosomal race [48]. Although much progress has been made in revealing patterns of speciation and hybridisation in tree weta, the molecular basis of mate recognition, fertilisation and other reproductive processes are not known. The present study describes the male reproductive transcriptomes for H. crassidens and H. thoracica. We identify putative male reproductive associated proteins, and investigate patterns of divergence of nine genes between three Hemideina species. We test the hypothesis that male reproductive associated proteins have elevated rates of positive selection compared to general metabolic, or housekeeping, genes.

Materials and methods

Sample collection

Hemideina thoracica, H. crassidens and Deinacrida mahoenui (Gibbs) specimens were collected across their known distributions between 2010 and 2012 (S1 Table), by day and night searching. All samples were collected under a permit issued by the Department of Conservation (CA-31615-OTH). Insects were transported live to Landcare Research, Auckland, and then snap frozen and stored at -80°C.

Transcriptome sequencing

Accessory gland and testis tissue from one adult male H. thoracica and H. crassidens (S1 Table) were dissected under a dissecting microscope in 100% ethanol. Total RNA was extracted from each tissue using TRIzol RNA extraction reagent (Life Technologies) according to the manufacturer’s protocol. A further RNA clean-up was performed using the RNeasy Mini Kit (Qiagen). RNA quality and quantity was determined using a Nanodrop (ThermoScientific) and an Agilent 2100 bioanalyzer (Agilent Technologies). High quality total RNA was used to synthesise cDNA using the SMARTerTM PCR cDNA synthesis kit (Clontech) with a modified oligo-dT primer (Cap-TRSA-CV) [50]. Double stranded cDNA was purified using AMPure beads (Agencourt). Library quality was assessed using an Agilent 2100 bioanalyzer (Agilent Technologies) and quantified with the Quanti-i TM Picogreen® assay (Life Technologies). Cleaned cDNA was fed into the Rapid Library Preparation Protocol (Roche, GS Junior Titanium Series, June 2010) at the fragment end repair step, with each tissue sample being MID barcoded. The resulting libraries were pooled by tissue and sequenced in two runs on a 454 GS Junior (Roche) at Landcare Research (Auckland).

Pre-processing, assembly and annotating RNA-seq data

Raw sequences were split by MID barcode using Geneious V5.4.6 [51]. Reads with ambiguous bases and low quality sequences were removed using SnoWhite 1.1.14 [52]. The primer and adaptor sequences were removed using CUTADAPT V1.1 [53]. Poly A/T tails longer than 15 bp from either end of the reads, and reads shorter than 50 bp were removed using PRINSEQ LITE V0.16 [54]. Cleaned reads were de novo assembled with Newbler GS de novo Assembler (V. 2.5.3), with default parameters, a minimum overlap of 25 bp and a minimum overlap identity of 95%. Redundancy in the alignment was removed using cd-hit-est V. 4.5.6 [55]. Poor de novo assembly of the H. thoracica dataset was observed, due to a lower sequencing quality of the testis library run. Therefore, in order to obtain a more representative dataset additional assembly steps were undertaken. The cleaned, trimmed reads from both H. thoracica libraries were reference assembled against the H. crassidens transcriptome using the Roche GS reference mapper (version 2.5.3, default parameters, except for a 25 bp overlap). The purpose of this reference assembly was to obtain contigs that were unassembled in the de novo assembly due to inadequate coverage. The reference and de novo assemblies were combined and subjected to a second round of redundancy removal with cd-hit-est. Preliminary tests on the combined assembly showed similar levels of blast homology, GO annotation and assembly statistics; therefore, this assembly replaced the de novo assembly for H. thoracica in downstream analyses. Singletons (unassembled reads) were excluded from downstream analysis. The assemblies were annotated using Blastx V2.6.0 [56] (e-value < 1e-5) against the GenBank non-redundant (nr) protein database (downloaded July 2017). Transcripts were searched for conserved protein domains with InterProScan [57] and GO terms were assigned using Blast2GO v2.8 [58]. Full-length transcripts were identified using Full-Lengther [59].

Identification of reproductive associated, orthologous, and candidate genes

Orthologous genes were identified using a bidirectional best hit method, which has been shown to outperform more complex algorithms for orthology predication [60]. A pair-wise reciprocal blastn approach was carried out in Geneious (e-value threshold 1e-3) with orthologues being called if the best blast hit was identical in both directions.

Putative reproductive associated genes were identified in a two-step process. First, transcripts were identified based on mapping counts of D. fallai muscle RNA-seq reads downloaded from SRA (SRA accession: SRR5965744) using RSEM [61]. Second, transcripts unique to the reproductive transcriptomes in H. thoracica and H. crassidens were identified by mapping the D. fallai short reads to each assembly using Bowtie2 [62] and identifying the transcripts with no counts. Within these candidates, signal peptides, cellular location and the presence of trans-membrane domains were identified with SignalP (v4.1) [63], ProtComp v9.0 ( and TMHMM v2.0 [64], respectively. Transcripts were retained as putative reproductive proteins if they had one of the following: (i) signal peptide, (ii) cellular localisation as extracellular and/or plasma membrane, (iii) transmembrane helix.

Candidate genes for downstream evolutionary analysis were chosen from among the contigs identified in the reproductive and orthologous gene screen. Candidates from the reproductive gene search were chosen based on their annotation, level of similarity (cut-off of 60%) and sequence length (minimum 400 bp). Orthologous candidates were based on a minimum transcript overlap of 200 bp between H. thoracica and H. crassidens transcripts, and the level of similarity at the amino acid and nucleotide level. Lastly, general metabolic control genes were chosen based on a minimum contig length of 300 bp and their involvement in general cellular processes, thereby ensuring tissue wide expression.

Sequencing of candidate genes

For Sanger sequencing samples, total RNA extractions from testis tissue followed the methods described above for the RNA-seq samples. Contaminating DNA was removed from RNA extractions prior to cDNA synthesis using TURBO DNase (Invitrogen). The first strand cDNA synthesis used the SuperScript III First Strand Kit (Invitrogen) following the manufacturer’s protocol. cDNA libraries were subsequently amplified using 5 μL first strand cDNA, 0.8 μM random hexamer primer, 2X PCR buffer (Roche), 2.5 mM MgCl2 (Roche), 0.2 mM dNTP (Roche), 1 U FastStart Taq DNA polymerase (Roche) in a total volume of 53 μL. Amplifications were performed on a GeneAmp PCR system 9700 thermal cycler (Applied Biosystems) using the following parameters: 5 min at 95°C, 3 min at 50°C, 40 sec 72°C; 40 cycles of 40 sec at 94°C, 40 sec at 65°C and 40 sec at 72°C; and 10 min at 72°C.

Primers for nine candidate genes (COI, Protease, Sflag, EFdelta, Unk2, Tkinase, Acp3, Acp4, Acp5) were designed using Primer 3 [65] implemented within Geneious. Primer pairs were designed to amplify products of 200–1500 bp in length, have TMs of 60°C (±3°C) and to have a GC continent of 40–60% (S2 Table). Target genes were PCR amplified with reactions consisting of approximately 5 ng DNA, 1X PCR buffer (Roche), 2 mM MgCl2 (Roche), 0.2 mM dNTP (Roche), 0.1–0.2 μM forward and reverse primers (Sigma-Aldrich), 1 U FastStart Taq DNA polymerase (Roche), in a total volume of 25 μl. Amplification were performed on a GeneAmp PCR system 9700 thermal cycler (Applied Biosystems) using the following parameters: 5 mins at 95°C; 40 cycles of 15 sec at 95°C, 30 sec at primer specific annealing temperature and 1 min 30 sec at 72°C; and 5 min at 72°C.

PCR products were sequenced using BigDye Terminator Cycle Sequencing Ready Reaction Mix v3.1 (Applied Biosystems). Cycle sequencing products were cleaned using the BigDye Xterminator Purification Kit (Applied Biosystems) and sequenced in both directions on the ABI Prism 3100 Genetic Analyzer (Applied Biosystems). Sequences were subsequently cleaned, trimmed and aligned using Geneious. In addition, the D. mahoenui and D. fallai transcriptomes (unpublished) were searched for COI and all candidate gene orthologues, respectively using a bidirectional tblastx approach, and included in downstream analysis (sequences given in S1 File).

Haplotype reconstruction for sequences that exhibited heterozygosity was performed using PHASE V. 2.1 [66, 67] prior to calculating descriptive statistics in DnaSP V. 5 [68]. Tajima’s D [69] and the McDonald-Kreitman test [70] were calculated in DNAsp. Haplotype networks were constructed using TCS V. 1.2.1 (Clement, Posada, Crandall 200), with gaps being considered as the 5th state and a 95% connection limit. The substitution model for the COI phylogeny was selected using the corrected Akaike information criterion [71] generated by jModel Test v.0.0.1 [72, 73]. A maximum likelihood phylogeny was constructed in Garli v2.0 [74] using 100 search and 1000 bootstrap replicates.

Inferring positive selection

Genes were identified for selection tests based on the number of non-synonymous changes. The three candidates with the most non-synonymous changes (Acp3, Protease, Unk2) were chosen for downstream analysis. For these three, a neighbour-joining phylogeny was generated for selection tests in Geneious. To screen for positive selection ω was estimated by maximum likelihood, using codon-based substitution models implemented in the CODEML package of PAML V. 4.5 [75]. The models implemented (M0, M1a, M2a, M3, M7, M8, M8a) are extensively described elsewhere [7678]. Complex models (M2a, M3, M8) allow more than one category of ω, thereby allowing individual codons to be identified as under positive selection when the average ω across the whole gene indicates purifying selection. Likelihood ration tests (LRTs) between nested models allows inference of positive selection acting on a sequence [75]. Codons under positive selection were identified using the Bayes empirical Bayes (BEB) method under the M8 model.

Results and discussion

Transcriptome assembly and characterisation

454 sequencing of H. thoracica and H. crassidens cDNA libraries resulted in a total of 254,628 reads, of which 73,012 and 59,465 reads were from H. thoracica, and 37,384 and 84,767 from H. crassidens testis and accessory gland tissue libraries, respectively. Raw sequences have been submitted to the GenBank Short Read Archive (BioProject: PRJNA353021). After trimming to remove bases with low quality scores, adapters and MID barcodes, 97% of the data remained. De novo assemblies were generated for each species as described above. The H. crassidens assembly generated 1,759 unigenes with an N50 of 608 bp and a maximum transcript size of 2,861 bp. In comparison, the H. thoracica assembly generated 2,691 unigenes with an N50 of 576 bp and a maximum transcript size of 1,860 bp. Both assemblies have been submitted to TSA under GFBX00000000 and GFBW00000000. A total of 890 and 1,537 transcripts were identified as being full length, respectively. Approximately 45% of the unigenes present in each assembly were functionally annotated using a tblastx search against the NCBI non-redundant database, with the species distribution of top matches overlapping among the two assemblies (S1 Fig). Functional annotation (GO) was similar across the two species, with the highest number of annotated transcripts related to cellular processes (GO:0009987) and metabolic process (GO:000812) (Fig 1). All unigenes were screened against the InterPro database, from which 2,690 and 1,753 H. thoracica and H. crassidens transcripts, respectively, were identified as containing conserved protein domains. The top 20 most frequent entries are shown in Table 1. These top entries show a diverse range of predicted functions, including proteins associated with general house-keeping roles and other that have been linked to reproductive functions. Domains identified include ubiquitin (IPR000626, IPR029071) and translation protein (IPR008991) domains. Many of the genes represented in these groups are probably highly conserved genes involved in the processes of transcription and protein degradation. Other domains identified, such as proteases (IPR001254, IPR018114) and protease inhibitors (IPR00215, IPR023796), have been associated with a number of reproductive functions, such as the modifying postmating changes in females [79], and are frequently identified in the study of insect SFPs [9, 80, 81].

Fig 1. Distribution of biological function annotation of two Hemideina transcriptomes.

Green bars: H. crassidens and blue bars: H. thoracica.

Table 1. The 20 most encountered InterPro accessions present in two Hemideina transcriptomes.

Identification of orthologous transcripts and reproductive proteins

The bidirectional best hit method identified 113 pairs of sequences that were putatively orthologous between the two species (S3 Table). For simplicity, these genes are hereafter referred to as orthologous. Some orthologues will have been missed using this approach, as in D. melanogaster the evolutionary rate of some SFPs has been shown to be so rapid that they lack any detectable similarity with their homologues from other Drosophila species [1, 24, 82, 83].

Putative reproductive associated transcripts were identified by mapping D. fallai muscle RNA-seq reads to each transcriptome. Transcripts unique to the reproductive transcriptomes (those lacking mapped reads) were further analysed to identify putative reproductive associated proteins. Of the transcripts unique to the reproductive transcriptome, 258 and 337 meet the criteria of having a signal peptide, transmembrane helix or localisation at the plasma membrane or extracellular for H. thoracica and H. crassidens, respectively (S4 Table). Roughly 19% of these had Blast hits, indicating that those lacking homology might be novel proteins or proteins highly diverged in weta. Among the genes with Blast hits, the most common molecular function GO terms were serine-type endopeptidase activity (GO:0004252), serine-type endopeptidase inhibitor activity (GO:0004867) and ATP binding (GO:0005524) (S2 Fig). Overall, the GO categories identified in the reproductive gene search are similar to categories commonly seen when studying insect SFPs [9, 16, 81, 84]. Various peptidase and peptidase regulators are among the reproductive proteins identified, and are believed to be essential for the regulation of reproduction through proteolytic cascades [28]. These types of proteins constitute a large proportion of the D. melanogaster [80], Anopheles gambiae [85], Aedes aegypti [86], Lutzomyia longipalphis [87] and Clitarchus hookeri [84] identified SFP and accessory gland proteins. The reproductive proteins identified in this screen are similar when compared with other insects, however a large proportion lack Blast hits. These unknown transcripts indicate the presence of novel or highly divergent proteins, and provide a large resource for the study of sexual reproduction and speciation in the New Zealand weta [80, 8590].

Candidate gene identification and sequencing

To study the patterns of molecular evolution of weta reproductive proteins, alignments of the candidates generated from the transcriptome sequencing were used to design PCR primers from nine genes (Table 2). Six putative reproductive proteins (Acp3, Acp4, Acp5, Sflag, Tkinase, and Protease) were identified as interesting candidates for downstream evolutionary analysis based on their blast homologies. The contig Unk2, despite lacking significant blast homology, was included for further analysis based on the interesting amino-acid pairwise identity observed during the orthologue gene screen. In addition, one nuclear (EFdelta) and one mitochondrial (COI) gene were included as general metabolic controls due to their tissue-wide expression. All nine genes were successfully amplified and sequenced from cDNA from, 19 H. thoracica, 11 H. crassidens, 5 H. trewicki and 1 D. mahoenui (outgroup) individuals. In addition, transcripts were identified within our unpublished RNA-seq data for D. fallai for all nine genes. All sequences have been submitted to NCBI GenBank (accession numbers: KY999988—KY999999, MF000001—MF000301).

Polymorphism, divergence and molecular evolution

Sequence data was obtained for COI from the majority of Hemideina individuals, resulting in a 672 bp alignment. The maximum likelihood phylogeny (Fig 2) supports each Hemideina species as monophyletic, with H. trewicki being sister to H. crassidens. The pairing of H. crassidens with H. trewicki is consistent with previous genetic and allozyme studies [41, 91].

Fig 2. Maximum likelihood phylogeny constructed using mitochondrial cytochrome oxidase subunit I (COI) DNA sequences from individuals representing three Hemideina and one Deinacrida species.

Bootstrap support values greater than 0.5 are indicated. Scale bar represents the number of substitutions per site.

Haplotype networks were constructed for every gene, except COI, rather than phylogenetic trees. Very little sequence divergence was present within these genes, thereby reducing the statistical power of phylogenetic reconstruction [92, 93]. Two genes (Unk2, Protease, Fig 3A and 3D) showed monophyletic groupings of alleles, whereas the remaining six genes showed the presence of shared alleles between at least two of the species sequenced (Figs 3 and 4). Generally speaking, both the reproductive and general metabolic control genes showed similar patterns. Previous work has shown that at both a genetic [39, 94] and karyotypic [48] level H. crassidens and H. trewicki are more genetically similar to each other than either is to H. thoracica, and hence are more likely to produce fertile hybrids. Of the 8 genes sequenced, only Protease and Unk2 show a complete lack of allele sharing among the species. The genes Tkinase, Sflag, Acp4, and Acp5, show sharing of alleles between H. thoracica and H. crassidens. The geographically restricted H. trewicki shares alleles with H. crassidens (EFdelta, Acp3, Acp5) and H. thoracica (Acp5). The two Deinacrida species are well differentiated from the three sampled Hemideina species at all loci, in agreement with previous studies [39, 91]. Overall these results demonstrate genetic differentiation among the three tree weta species, in agreement with McKean et al. [48].

Fig 3.

Haplotype network of A) Unk2, B) Sflag, C) Tkinase, and D) Protease gene regions. Circles represent different haplotypes, with the circles area being proportional to the frequency of each haplotype. Lines between haplotypes represent mutational steps between sequences. The empty circles represent inferred unsampled haplotypes. Colours correspond to species: Red, H. crassidens; Blue, H. thoracica; Yellow, H. trewicki.

Fig 4.

Haplotype network of A) Acp4, B) Acp3, C) EFdelta, and D) Acp5 gene regions. Circles represent different haplotypes, with the circles area being proportional to the frequency of each haplotype. Lines between haplotypes represent mutational steps between sequences. The empty circles represent inferred unsampled haplotypes. Colours correspond to species: Red, H. crassidens; Blue, H. thoracica; Yellow, H. trewicki.

A summary of intraspecific sequence variation is shown in Table 3. Intraspecific variation within H. thoracica was greater than that observed in H. crassidens and H. trewicki for the majority of genes examined. This is consistent with allozyme and mitochondrial DNA studies that show H. thoracica has the highest levels of intraspecific diversity of all Hemideina species [39, 41, 91]. This signature is consistent with inferences that the range of H. thoracica has recently expanded southwards [46, 95],while in comparison, little to no diversity was observed within the H. trewicki samples. This is not an unexpected result as all samples originated from a single population.

Table 3. Summary statistics of intra-specific sequence variation within three Hemideina species.

The reproductive-associated candidate genes tended to display higher levels of within species diversity than the general metabolic controls. However, in some reproductive genes, especially Acp5 and Tkinase, the observed level of genetic diversity was at the same or similar levels as the general metabolic controls. The relatively lower levels of diversity in these two reproductive genes suggest they are functionally constrained. However, this requires further investigation, as only two metabolic controls were included in this study, and only partial transcripts were sequenced. Possible explanations for the lower diversity, include the sequenced region may be in a functionally constrained region of the protein, with relaxed selection occurring upstream or downstream of the sequenced region, or these genes may be located in regions of low recombination.

Within the Acp3 alignment an allelic variant containing a 24 bp indel was identified. All H. thoracica and two H. crassidens individuals have the insertion, while the remainder of H. crassidens and all H. trewicki samples lack the insertion. The 24 bp indel, appears to be a true allelic variant rather than the effect of preferential amplification of a paralogous gene as some individuals had only one copy of either the deletion or complete variant. If the deletion variant was in fact paralogous amplification then both copies would have been expected in all individuals of H. crassidens. InterProScan analysis revealed the presence of C-lectin type domains within the coding sequence. Lectins and lectin-related proteins have been shown to be involved in carbohydrate binding and the mediation of sperm-egg interactions [8890], suggesting that this is an interesting reproductive candidate gene family.

At the species level, Tajima’s D was significant for the serine protease gene (Protease) within H. crassidens, (D = 2.17, Table 4) which may indicate balancing selection or demographic influences. In addition, for Acp4, Acp3, Acp5, Unk2, and Tkinase Tajima’s D statistics were negative for at least one species, thereby indicating an excess of rare or recent mutations that may be due to purifying selection or a recent demographic expansion, the latter of which has been observed for H. crassidens and H. thoracica [95]. Under the McDonald-Kreitman test no departures from neutrality were detected for any of the genes (S5 Table).

To study the patterns of molecular evolution of weta reproductive proteins, the ratio of nonsynonymous to synonymous substitution rates of protein-coding sequences (ω) was calculated for two reproductive genes (Acp3, Protease) and the unknown gene, Unk2. The candidate Unk2 was included as part of the selection tests, as initial screening identified an ORF, which showed higher numbers of nonsynonymous than synonymous substitutions (Table 3). In the case of Acp3 the mean ω ratio was >1, indicating an excess of nonsynonymous changes across the protein coding region as a whole (Table 5). In contrast, Unk2 and Protease had an ω <1 (Table 5). Omega ratios averaged over an entire protein coding region are typically <1, due to positive selection commonly acting on specific domains or residues [96], with evidence suggesting genes with mean ω ratios above 0.5 are experiencing episodes of adaptive evolution [1, 7, 97, 98]. Individual amino acid residues likely to have been influenced by selection were identified using site-based ω calculations, with likelihood ratio tests and chi-squared distributions used to assess the goodness of fit for a given model [77, 99, 100]. For comparison, we used six codon-substitution models to assess the mode of selection acting on each amino acid residue in our candidate genes. We found that for Acp3 and Protease there is significant among-site variation in ω, with the M3 model permitting three ω values providing a significantly better fit to the data than the M0 model (p-value<0.05; df = 4; M0:M3; Table 5, S6 Table). The M1a:M2a comparison was insignificant for all three genes. A more conservative approach for testing for positive selection is the M8:M8a comparison, under this model Acp3 was identified was being under positive selection with a mean ω of 7.9 (Tables 5 and S6).

Table 5. Likelihood ratio tests of positive section using PAML site-specific models.

A Bayes Empirical Bayes computation [101] implemented in PAML was used to assess the significance of the ω ratio at each codon position. Under the M8 model five sites were assigned to the positively selected class (ω >1), only one of which had a probability > 95% (ω = 6.366, Table 5, Fig 5). Observed changes at three of the five sites were non-conservative (S7 Table). It is acknowledged that likelihood-based methods can produce high levels of false positives [102, 103], however the alternative parsimony-based models tend to be very conservative and have low power detecting true positives, particularly in small datasets such as this one [104, 105]. The five sites presented here represent testable hypotheses for functionally important regions under selection within Acp3 that could be examined with a larger dataset and other analysis methods. However, it should be noted that the function of Acp3 is unknown, as is the exact position of these sites within the overall protein structure.

Fig 5. Positive selection within Acp3.

Red line represents the mean posterior omega, and the blue line represents the probability of each codon being under positive selection. The values were calculated using a Bayes Empirical Bayes analysis under the M8 site-specific model in Paml. Codon position based on full-length alignment. The annotations identified using InterProScan and the position of the 24 bp indel are shown in relation to codon position.


Here we present the first male reproductive transcriptomes for H. crassidens and H. thoracica, resulting in putative gene sets of 1,754 and 2,691 non-redundant gene sets, respectively. We identified 865 putative reproductive associated proteins, and 113 orthologs, from which nine candidates were used for downstream evolutionary analyses. Our results suggest that positive selection may be acting on some Hemideina SFPs; in contrast, we were unable to detect positive selection on the general metabolic control genes. The lectin-related Acp3 gene shows evidence for selection acting along the gene as a whole and on particular amino acids. This presents a testable hypothesis into what selection may be occurring on weta reproductive proteins. A better understanding can be achieved by incorporating functional and population genomics with candidate gene approaches to reveal the relationship between the evolution of these genes and mate recognition and speciation. In addition, the transcriptome data generated represents a first step in the identification of reproductive associated proteins in weta. These transcriptomic sequences will provide a valuable resource for further research into the evolution of reproductive proteins and speciation of New Zealand weta.

Supporting information

S1 Table. Sample collection details.

**Samples used for 454 sequencing.


S2 Table. Primer sequences for candidate genes.

Annotations from tblastx against non-redundant database.


S3 Table. Orthologous contigs pairs identified in the bidirectional tblastx search.


S4 Table. Contigs identified in the reproductive protein search.


S5 Table. McDonald Kreitman test results for each gene.

a Yates correction is applied to G tests.


S6 Table. Likelihood values and parameter estimates for site-specific models for each gene.

Parameters in brackets are not free, and therefore are not counted when considering difference in the number of parameters for nested model comparisons.


S7 Table. Sites identified as putatively under selection and their posterior probabilities under the M3 and M8 models.

a Amino acid site in the original alignment, including gaps; BEB, Bayes Empirical Bayes posterior probability; NEB, Naïve Empirical Bayes; **Probability > 99%.


S1 Fig.

Top-Hit species distribution for tblastx results for A) Hemideina thoracica and B) Hemideina crassidens.


S2 Fig.

Gene ontology categories identified in the reproductive gene screen for A) Hemideina thoracica and B) Hemideina crassidens.


S1 File. Deinacrida fallai and D. mahoenui orthologues identified from an unpublished de novo assembled illumina transcriptome data.



We would like to thank George Gibbs, Robert Hoare, George Twort, Owen Twort, Ian Twort, Genesta Twort, Dave Seldon, Peter Richie, Shelley Meyers, Chrissie Painting, Luke Dunning, Robin Howarth, Kathy Howarth, Paulina Giraldo Perez, Corrine Watts and Katy Hill for assistance with field work and providing samples, the New Zealand eScience Infrastructure (NeSI) high-performance computing facilities and the staff at the Centre for eResearch at the University of Auckland, for bioinformatic support and the Department of Conservation for access to field sites. We thank Joe Hull and three anonymous reviewers for comments that improved the manuscript.


  1. 1. Swanson WJ, Clark AG, Waldrip-Dail HM, Wolfner MF, Aquadro CF. Evolutionary EST analysis identifies rapidly evolving male reproductive proteins in Drosophila. Proceedings of the National Academy of Sciences. 2001;98(13):7375–9. pmid:11404480
  2. 2. Swanson WJ, Vacquier VD. The rapid evolution of reproductive proteins. Nature Reviews Genetics. 2002;3(2):137–44. pmid:11836507
  3. 3. Clark NL, Aagaard JE, Swanson WJ. Evolution of reproductive proteins from animals and plants. Reproduction. 2006;131(1):11–22. pmid:16388004
  4. 4. Walters JR, Harrison RG. Combined EST and proteomic analysis identifies rapidly evolving seminal fluid proteins in heliconius butterflies. Molecular Biology and Evolution. 2010;27(9):2000–13. pmid:20375075
  5. 5. Walters JR, Harrison RG. Decoupling of rapid and adaptive evolution among seminal fluid proteins in Heliconius butterflies with divergent mating sytems. Evolution. 2011;65(10):2855–71. pmid:21967427
  6. 6. Wilburn DB, Swanson WJ. From molecules to mating: Rapid evolution and biochemical studies of reproductive proteins. Journal of Proteomics. 2016;135:12–25. pmid:26074353
  7. 7. Andrés JA, Maroja LS, Bogdanowicz SM, Swanson WJ, Harrison RG. Molecular evolution of seminal proteins in field crickets. Molecular Biology and Evolution. 2006;23(8):1574–84. pmid:16731569
  8. 8. Braswell WE, Andrés JA, Maroja LS, Harrison RG, Howard DJ, Swanson WJ. Identification and comparative analysis of accessory gland proteins in Orthoptera. Genome. 2006;49(9):1069–80. pmid:17110987
  9. 9. Ahmed-Braimah YH, Unckless RL, Clark AG. Evolutionary Dynamics of Male Reproductive Genes in the Drosophila virilis Subgroup. G3: Genes|Genomes|Genetics. 2017. pmid:28739599
  10. 10. Wolfner MF. Tokens of love: functions and regulation of Drosophila male accessory gland products. Insect Biochemistry and Molecular Biology. 1997;27(3):179–92. pmid:9090115
  11. 11. Gillott C. Male accessory gland secretions: modulators of female reproductive physiology and behavior. Annual Review of Entomology. 2003;48(1):163–84.
  12. 12. Neubaum DM, Wolfner MF. Mated Drosophila melanogaster Females Require a Seminal Fluid Protein, Acp36DE, to Store Sperm Efficiently. Genetics. 1999;153(2):845–57. pmid:10511562
  13. 13. Lange AB, Loughton BG. An oviposition-stimulating factor in the male accessory reproductive gland of the locust, Locusta migratoria. General and Comparative Endocrinology. 1985;57(2):208–15. pmid:3979802
  14. 14. Chen PS, Stumm-Zollinger E, Aigaki T, Balmer J, Bienz M, Böhlen P. A male accessory gland peptide that regulates reproductive behavior of female D. melanogaster. Cell. 1988;54(3):291–8. pmid:3135120
  15. 15. Marshall JL, Huestis DL, Hiromasa Y, Wheeler S, Oppert C, Marshall SA, et al. Identification, RNAi Knockdown, and Functional Analysis of an Ejaculate Protein that Mediates a Postmating, Prezygotic Phenotype in a Cricket. PLoS ONE. 2009;4(10):e7537. pmid:19851502
  16. 16. Bayram H, Sayadi A, Goenaga J, Immonen E, Arnqvist G. Novel seminal fluid proteins in the seed beetle Callosobruchus maculatus identified by a proteomic and transcriptomic approach. Insect Molecular Biology. 2017;26(1):58–73. pmid:27779332
  17. 17. Al-Wathiqui N, Dopman EB, Lewis SM. Postmating transcriptional changes in the female reproductive tract of the European corn borer moth. Insect Molecular Biology. 2016;25(5):629–45. pmid:27329655
  18. 18. Himuro C, Ikegawa Y, Honma A. Males Use Accessory Gland Substances to Inhibit Remating by Females in West Indian Sweetpotato Weevil (Coleoptera: Curculionidae). Annals of the Entomological Society of America. 2017;110(4):374–80.
  19. 19. Andrés JA, Arnqvist G. Genetic divergence of the seminal signal—receptor system in houseflies: the footprints of sexually antagonistic coevolution? Proceedings of the Royal Society of London Series B: Biological Sciences. 2001;268(1465):399–405. pmid:11270437
  20. 20. Rice WR. Sexually antagonistic male adaptation triggered by experimental arrest of female evolution. Nature. 1996;381(6579):232–4. pmid:8622764
  21. 21. Turner LM, Hoekstra HE. Causes and consequences of the evolution of reproductive proteins. International Journal of Developmental Biology. 2008;52(5):769.
  22. 22. Marshall JL, Huestis DL, Garcia C, Hiromasa Y, Wheeler S, Noh S, et al. Comparative Proteomics Uncovers the Signature of Natural Selection Acting on the Ejaculate Proteomes of Two Cricket Species Isolated by Postmating, Prezygotic Phenotypes. Molecular Biology and Evolution. 2011;28(1):423–35. pmid:20805188
  23. 23. Kern AD, Jones CD, Begun DJ. Molecular Population Genetics of Male Accessory Gland Proteins in the Drosophila simulans Complex. Genetics. 2004;167(2):725–35. pmid:15238524
  24. 24. Mueller JL, Ram KR, McGraw LA, Bloch Qazi MC, Siggia ED, Clark AG, et al. Cross-Species Comparison of Drosophila Male Accessory Gland Protein Genes. Genetics. 2005;171(1):131–43. pmid:15944345
  25. 25. Andrés JA, Maroja LS, Harrison RG. Searching for candidate speciation genes using a proteomic approach: seminal proteins in field crickets. Proceedings of the Royal Society B: Biological Sciences. 2008;275(1646):1975–83. pmid:18495616
  26. 26. Almeida FC, DeSalle R. Genetic differentiation and adaptive evolution at reproductive loci in incipient Drosophila species. Journal of Evolutionary Biology. 2017;30(3):524–37. pmid:27883252
  27. 27. Haerty W, Jagadeeshan S, Kulathinal RJ, Wong A, Ravi Ram K, Sirot LK, et al. Evolution in the Fast Lane: Rapidly Evolving Sex-Related Genes in Drosophila. Genetics. 2007;177(3):1321–35. pmid:18039869
  28. 28. LaFlamme BA, Ravi Ram K, Wolfner MF. The Drosophila melanogaster Seminal Fluid Protease “Seminase” Regulates Proteolytic and Post-Mating Reproductive Processes. PLoS Genet. 2012;8(1):e1002435. pmid:22253601
  29. 29. Dean MD, Clark NL, Findlay GD, Karn RC, Yi X, Swanson WJ, et al. Proteomics and Comparative Genomic Investigations Reveal Heterogeneity in Evolutionary Rate of Male Reproductive Proteins in Mice (Mus domesticus). Molecular Biology and Evolution. 2009;26(8):1733–43. pmid:19420050
  30. 30. Findlay GD, Yi X, MacCoss MJ, Swanson WJ. Proteomics Reveals Novel Drosophila Seminal Fluid Proteins Transferred at Mating. PLoS Biol. 2008;6(7):e178. pmid:18666829
  31. 31. Tsaur SC, Ting CT, Wu CI. Positive selection driving the evolution of a gene of male reproduction, Acp26Aa, of Drosophila: II. Divergence versus polymorphism. Molecular Biology and Evolution. 1998;15(8):1040–6. pmid:9718731
  32. 32. Tsaur SC, Wu CI. Positive selection and the molecular evolution of a gene of male reproduction, Acp26Aa of Drosophila. Molecular Biology and Evolution. 1997;14(5):544–9. pmid:9159932
  33. 33. Wong A, Albright SN, Wolfner MF. Evidence for structural constraint on ovulin, a rapidly evolving Drosophila melanogaster seminal protein. Proceedings of the National Academy of Sciences. 2006;103(49):18644–9. pmid:17130459
  34. 34. Fuyama Y. Species-specificity of paragonial substances as an isolating mechanism in Drosophila. Cellular and Molecular Life Sciences. 1983;39(2):190–2.
  35. 35. Andrés JA, Larson EL, Bogdanowicz SM, Harrison RG. Patterns of Transcriptome Divergence in the Male Accessory Gland of Two Closely Related Species of Field Crickets. Genetics. 2012. pmid:23172857
  36. 36. Griffin MJ, Trewick SA, Wehi PM, Morgan-Richards M. Exploring the concept of niche convergence in a land without rodents: The case of weta as small mammals. New Zealand Journal of Ecology. 2011;35(3).
  37. 37. Gibbs G. New Zealand Weta: Kyodo Printing Co Ltd, Singapore; 1998.
  38. 38. McGuinness CA. The conservation requirements of New Zealand’s nationally threatened invertebrates. In: Department of Conservation, editor. Department of Conservation, Wellington, New Zealand.2001.
  39. 39. Trewick SA, Morgan-Richards M. After the deluge: Mitochondrial DNA indicates Miocene radiation and Pliocene adaptation of tree and giant weta (Orthoptera: Anostostomatidae). Journal of Biogeography. 2005;32(2):295–309.
  40. 40. White DJ, Watts C, Allwood J, Prada D, Stringer I, Thornburrow D, et al. Population history and genetic bottlenecks in translocated Cook Strait giant weta, Deinacrida rugosa: recommendations for future conservation management. Conservation Genetics. 2016:1–12.
  41. 41. Trewick SA, Morgan-Richards M. Phylogenetics of New Zealand's tree, giant and tusked weta (Orthoptera: Anostostomatidae): evidence from mitochondrial DNA. Journal of Orthoptera Research. 2004;13(2):185–96.
  42. 42. Trewick SA, Morgan-Richards M. On the distribution of tree weta in the North Island, New Zealand. Journal—Royal Society of New Zealand. 1995;25(4):485–93.
  43. 43. Morgan-Richards M. Robertsonian translocations and B chromosomes in the Wellington tree weta, Hemideina crassidens (Orthoptera: Anostostomatidae). Hereditas. 2000;132(1):49–54. pmid:10857259
  44. 44. Morgan-Richards M, King T, Trewick S. The evolutionary history of tree weta; A genetic approach. In: Field LH, editor. The biology of wetas, king crickets and their allies: Wallingford, Oxon., UK; New York, N.Y., USA: CABI Pub., c2001; 2001.
  45. 45. Morgan-Richards M. Intraspecific karyotype variation is not concordant with allozyme variation in the Auckland tree weta of New Zealand, Hemideina thoracica (Orthoptera: Stenopelmatidae). Biological Journal of the Linnean Society. 1997;60(4):423–42.
  46. 46. Morgan-Richards M, Trewick SA, Wallis GP. Chromosome races with Pliocene origins: Evidence from mtDNA. Heredity. 2001;86(3):303–12.
  47. 47. Morgan-Richards M, Wallis GP. A comparison of five hybrid zones of the weta Hemideina thoracica (Orthoptera: Anostostomatidae): Degree of cytogenetic differentiation fails to predict zone width. Evolution. 2003;57(4):849–61. pmid:12778554
  48. 48. McKean NE, Trewick SA, Morgan-Richards M. Comparative cytogenetics of North Island tree wētā in sympatry. New Zealand Journal of Zoology. 2015:1–12.
  49. 49. Morgan-Richards M. A new species of tree weta from the North Island of New Zealand (Hemideina Stenopelmatidae: Orthoptera). New Zealand Entomologist. 1995;18(1):15–23.
  50. 50. Meyer E, Aglyamova GV, Wang S, Buchanan-Carter J, Abrego D, Colbourne JK, et al. Sequencing and de novo analysis of a coral larval transcriptome using 454 GSFlx. BMC Genomics. 2009;10.
  51. 51. Drummond A, Ashton B, Buxton S, Cheung M, Heled J, Kearse M, et al. Geneious. 5.4.6 ed: Available from; 2011.
  52. 52. Dlugosch KM, Rieseberg LH. SnoWhite: A pipeline for aggressive cleaning of next-generation sequence reads. In prep.
  53. 53. Martin M. Cutadapt removes adapter sequences from high-throughput sequencing reads. EMBnetJournal. 2011;17(1):10–2.
  54. 54. Schmieder R, Edwards R. Quality control and preprocessing of metagenomic datasets. Bioinformatics. 2011. pmid:21278185
  55. 55. Li W, Godzik A. Cd-hit: a fast program for clustering and comparing large sets of protein or nucleotide sequences. Bioinformatics. 2006;22(13):1658–9. pmid:16731699
  56. 56. Altschul SF, Madden TL, Schäffer AA, Zhang J, Zhang Z, Miller W, et al. Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Research. 1997;25(17):3389–402. pmid:9254694
  57. 57. Quevillon E, Silventoinen V, Pillai S, Harte N, Mulder N, Apweiler R, et al. InterProScan: protein domains identifier. Nucleic Acids Research. 2005;33(suppl_2):W116–W20. pmid:15980438
  58. 58. Conesa A, Götz S, García-Gómez JM, Terol J, Talón M, Robles M. Blast2GO: A universal tool for annotation, visualization and analysis in functional genomics research. Bioinformatics. 2005;21(18):3674–6. pmid:16081474
  59. 59. Lara A, Pérez-Trabado G, Villalobos D, Díaz-Moreno S, Cantón F, Claros M. A Web Tool to Discover Full-Length Sequences—Full-Lengther Innovations in Hybrid Intelligent Systems. In: Corchado E, Corchado J, Abraham A, editors. Advances in Soft Computing. 44: Springer Berlin / Heidelberg; 2007. p. 361–8.
  60. 60. Altenhoff AM, Dessimoz C. Phylogenetic and Functional Assessment of Orthologs Inference Projects and Methods. PLoS Comput Biol. 2009;5(1):e1000262. pmid:19148271
  61. 61. Li B, Dewey CN. RSEM: accurate transcript quantification from RNA-Seq data with or without a reference genome. BMC Bioinformatics. 2011;12(1):323. pmid:21816040
  62. 62. Langmead B, Salzberg SL. Fast gapped-read alignment with Bowtie 2. Nat Meth. 2012;9(4):357–9. pmid:22388286
  63. 63. Petersen TN, Brunak S, von Heijne G, Nielsen H. SignalP 4.0: discriminating signal peptides from transmembrane regions. Nat Meth. 2011;8(10):785–6. pmid:21959131
  64. 64. Krogh A, Larsson B, von Heijne G, Sonnhammer EL. Predicting transmembrane protein topology with a hidden Markov model: application to complete genomes. J Mol Biol. 2001;305(3):567–80. Epub 2001/01/12. pmid:11152613.
  65. 65. Rozen S, Skaletsky H. Primer3 on the WWW for General Users and for Biologist Programmers. 1321999. p. 365–86.
  66. 66. Stephens M, Donnelly P. A Comparison of Bayesian Methods for Haplotype Reconstruction from Population Genotype Data. American journal of human genetics. 2003;73(5):1162–9. pmid:14574645
  67. 67. Stephens M, Smith NJ, Donnelly P. A New Statistical Method for Haplotype Reconstruction from Population Data. American journal of human genetics. 2001;68(4):978–89. pmid:11254454
  68. 68. Librado P, Rozas J. DnaSP v5: a software for comprehensive analysis of DNA polymorphism data. Bioinformatics. 2009;25(11):1451–2. pmid:19346325
  69. 69. Tajima F. Statistical Method for Testing the Neutral Mutation Hypothesis by DNA Polymorphism. Genetics. 1989;123(3):585–95. pmid:2513255
  70. 70. McDonald JH, Kreitman M. Adaptive protein evolution at the Adh locus in Drosophila. Nature. 1991;351(6328):652–4. pmid:1904993
  71. 71. Hurvich CM, Tsai C-L. Regression and time series model selection in small samples. Biometrika. 1989;76(2):297–307.
  72. 72. Posada D. jModelTest: Phylogenetic Model Averaging. Molecular Biology and Evolution. 2008;25(7):1253–6. pmid:18397919
  73. 73. Guindon S, Gascuel O. A Simple, Fast, and Accurate Algorithm to Estimate Large Phylogenies by Maximum Likelihood. Systematic biology. 2003;52(5):696–704. pmid:14530136
  74. 74. Zwickl . Derrick J. Genetic algorithm approaches for the phylogenetic analysis of large biological sequence datasets under the maximum likelihood criterion: University of Texas as Austin; 2006.
  75. 75. Yang Z. PAML 4: Phylogenetic analysis by maximum likelihood. Molecular Biology and Evolution. 2007;24(8):1586–91. pmid:17483113
  76. 76. Yang Z, Nielsen R. Estimating synonymous and nonsynonymous substitution rates under realistic evolutionary models. Molecular Biology and Evolution. 2000;17(1):32–43. pmid:10666704
  77. 77. Yang Z, Nielsen R, Goldman N, Pedersen A-MK. Codon-Substitution Models for Heterogeneous Selection Pressure at Amino Acid Sites. Genetics. 2000;155(1):431–49. pmid:10790415
  78. 78. Swanson WJ, Nielsen R, Yang Q. Pervasive Adaptive Evolution in Mammalian Fertilization Proteins. Molecular Biology and Evolution. 2003;20(1):18–20. pmid:12519901
  79. 79. Ravi Ram K, Ji S, Wolfner MF. Fates and targets of male accessory gland proteins in mated female Drosophila melanogaster. Insect Biochemistry and Molecular Biology. 2005;35(9):1059–71. pmid:15979005
  80. 80. Ravi Ram K, Wolfner MF. Seminal influences: Drosophila Acps and the molecular interplay between males and females during reproduction. Integrative and Comparative Biology. 2007;47(3):427–45. pmid:21672851
  81. 81. Sirot LK, Hardstone MC, Helinski MEH, Ribeiro JMC, Kimura M, Deewatthanawong P, et al. Towards a Semen Proteome of the Dengue Vector Mosquito: Protein Identification and Potential Functions. PLOS Neglected Tropical Diseases. 2011;5(3):e989. pmid:21423647
  82. 82. Schully S, Hellberg M. Positive Selection on Nucleotide Substitutions and Indels in Accessory Gland Proteins of the Drosophila pseudoobscura Subgroup. Journal of Molecular Evolution. 2006;62(6):793–802. pmid:16752217
  83. 83. Begun DJ, Lindfors HA, Thompson ME, Holloway AK. Recently evolved genes identified from Drosophila yakuba and Drosophila erecta accessory gland expressed sequence tags. GENETICS; 2006.
  84. 84. Wu C, Crowhurst RN, Dennis AB, Twort VG, Liu S, Newcomb RD, et al. De Novo Transcriptome Analysis of the Common New Zealand Stick Insect Clitarchus hookeri (Phasmatodea) Reveals Genes Involved in Olfaction, Digestion and Sexual Reproduction. PLOS ONE. 2016;11(6):e0157783. pmid:27336743
  85. 85. Rogers DW, Baldini F, Battaglia F, Panico M, Dell A, Morris HR, et al. Transglutaminase-Mediated Semen Coagulation Controls Sperm Storage in the Malaria Mosquito. PLoS Biol. 2009;7(12):e1000272. pmid:20027206
  86. 86. Sirot LK, Poulson RL, Caitlin McKenna M, Girnary H, Wolfner MF, Harrington LC. Identity and transfer of male reproductive gland proteins of the dengue vector mosquito, Aedes aegypti: Potential tools for control of female feeding and reproduction. Insect Biochemistry and Molecular Biology. 2008;38(2):176–89. pmid:18207079
  87. 87. Azevedo RVDM, Dias DBS, Bretãs JAC, Mazzoni CJ, Souza NA, Albano RM, et al. The Transcriptome of Lutzomyia longipalpis (Diptera: Psychodidae) Male Reproductive Organs. PLoS ONE. 2012;7(4):e34495. pmid:22496818
  88. 88. Romero A, Romao MJ, Varela PF, Kolln I, Dias JM, Carvalho AL, et al. The crystal structures of two spermadhesins reveal the CUB domain fold. Nat Struct Biol. 1997;4(10):783–8. pmid:9334740
  89. 89. Mueller JL, Ripoll DR, Aquadro CF, Wolfner MF. Comparative structural modeling and inference of conserved protein classes in Drosophila seminal fluid. Proceedings of the National Academy of Sciences of the United States of America. 2004;101(37):13542–7. pmid:15345744
  90. 90. Takemori N, Yamamoto M-T. Proteome mapping of the Drosophila melanogaster male reproductive system. PROTEOMICS. 2009;9(9):2484–93. pmid:19343724
  91. 91. Morgan-Richards M, Gibbs G. A phylogenetic analysis of New Zealand giant and tree weta (Orthoptera: Anostostomatidae: Deinacrida and Hemideina) using morphological and genetic characters. Invertebrate Taxonomy. 2001;15(1):1–12.
  92. 92. Schierup MH, Hein J. Consequences of Recombination on Traditional Phylogenetic Analysis. Genetics. 2000;156(2):879–91. pmid:11014833
  93. 93. Posada D, Crandall KA. Intraspecific gene genealogies: trees grafting into networks. Trends in Ecology & Evolution. 2001;16(1):37–45.
  94. 94. Bulgarella M, Trewick SA, Godfrey AJR, Sinclair BJ, Morgan-Richards M. Elevational variation in adult body size and growth rate but not in metabolic rate in the tree weta Hemideina crassidens. Journal of Insect Physiology. 2015;75:30–8. pmid:25753546
  95. 95. Minards NA. Physiological Ecology of Two Tree Weta Species [Master's dissertation]. Palmerston North—New Zealand: Massey University; 2011.
  96. 96. Nielsen R, Yang Z. Likelihood Models for Detecting Positively Selected Amino Acid Sites and Applications to the HIV-1 Envelope Gene. Genetics. 1998;148(3):929–36. pmid:9539414
  97. 97. Clark NL, Swanson WJ. Pervasive Adaptive Evolution in Primate Seminal Proteins. PLoS Genet. 2005;1(3):e35. pmid:16170411
  98. 98. Swanson WJ, Wong A, Wolfner MF, Aquadro CF. Evolutionary Expressed Sequence Tag Analysis of Drosophila Female Reproductive Tracts Identifies Genes Subjected to Positive Selection. Genetics. 2004;168(3):1457–65. pmid:15579698
  99. 99. Yang Z, Nielsen R. Codon-Substitution Models for Detecting Molecular Adaptation at Individual Sites Along Specific Lineages. Molecular Biology and Evolution. 2002;19(6):908–17. pmid:12032247
  100. 100. Yang Z. PAML: a program package for phylogenetic analysis by maximum likelihood. Computer applications in the biosciences: CABIOS. 1997;13(5):555–6. pmid:9367129
  101. 101. Yang Z, Wong WSW, Nielsen R. Bayes Empirical Bayes Inference of Amino Acid Sites Under Positive Selection. Molecular Biology and Evolution. 2005;22(4):1107–18. pmid:15689528
  102. 102. Suzuki Y, Nei M. Simulation Study of the Reliability and Robustness of the Statistical Methods for Detecting Positive Selection at Single Amino Acid Sites. Molecular Biology and Evolution. 2002;19(11):1865–9. pmid:12411595
  103. 103. Suzuki Y, Nei M. Reliabilities of Parsimony-based and Likelihood-based Methods for Detecting Positive Selection at Single Amino Acid Sites. Molecular Biology and Evolution. 2001;18(12):2179–85. pmid:11719567
  104. 104. Suzuki Y, Gojobori T. A method for detecting positive selection at single amino acid sites. Molecular Biology and Evolution. 1999;16(10):1315–28. pmid:10563013
  105. 105. Wong W, Yang Z, Goldman N, Nielsen R. Accuracy and Power of Statistical Methods for Detecting Adaptive Evolution in Protein Coding Sequences and for Identifying Positively Selected Sites. Genetics. 2004;168(2):1041–51. pmid:15514074