Lineage-specific head development in the coffin-headed cricket Loxoblemmus equestris links concentrated insect metamorphosis with novel trait evolution

Lineage-specific head development in the coffin-headed cricket Loxoblemmus equestris links concentrated insect metamorphosis with novel trait evolution | Research Square window.SnipcartSettings = { analytics: { enabled: false } }; (function() { var accessVector = localStorage.getItem('access_vector') || ''; window.dataLayer = window.dataLayer || []; if (accessVector) { window.dataLayer.push({ user: { profile: { profileInfo: { snid: accessVector } } } }); } })(); (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0],j=d.createElement(s),dl=l!='dataLayer'?'&l='+l:'';j.async=true;j.src='https://www.googletagmanager.com/gtm.js?id='+i+dl;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-K279D39R'); Browse Preprints In Review Journals COVID-19 Preprints AJE Video Bytes Research Tools Research Promotion AJE Professional Editing AJE Rubriq About Preprint Platform In Review Editorial Policies Our Team Advisory Board Help Center Sign In Submit a Preprint Cite Share Download PDF Research Article Lineage-specific head development in the coffin-headed cricket Loxoblemmus equestris links concentrated insect metamorphosis with novel trait evolution Mizuho Yoneda, Shinichi Morita, Teruyuki Niimi, Takaaki Daimon, and 1 more This is a preprint; it has not been peer reviewed by a journal. https://doi.org/ 10.21203/rs.3.rs-6301946/v1 This work is licensed under a CC BY 4.0 License Status: Published Journal Publication published 16 Jul, 2025 Read the published version in Developmental Biology Advances → Version 1 posted 10 You are reading this latest preprint version Abstract Background Lineage-specific adult structures form through modifications of preexisting juvenile body parts during postembryonic development in insects. It remains unclear how these novel traits originate from ancestral structures within the constrained body plan. In the coffin-headed cricket Loxoblemmus equestris , an ancestral rounded head shape directly transforms into a flattened, derived form in a sex-specific manner. To understand the origin of novel traits, we investigated the development of the adult head in L . equestris as a model of lineage-specific novelty. Results Detailed two- and three-dimensional analyses of the developing head revealed that sexually dimorphic epithelial patterns formed in a specific region, the frons, during the preadult instar. The male-specific head shapes are formed following the final molt to adulthood even after timing shifts of the metamorphosis induced by RNA interference targeting the evolutionarily conserved metamorphic gene network. Conclusions These findings demonstrate that adult metamorphosis, led by E93, locally relaxes the body plan constraint to permit dramatic transformation of juvenile body parts into a novel head shape by modifying epithelial folding in L . equestris . This highlights concentrated metamorphosis through the final molt as a driver that creates lineage- and sex-specific adult forms in the hexapod lineage. Figures Figure 1 Figure 2 Figure 3 Figure 4 Figure 5 Figure 6 Figure 7 Background Lineage-specific diversification of homologous body parts creates evolutionary novelties from a constrained body plan shared within an animal phylum. Insecta is a taxon that shows prominent morphological novelties, such as pterygote thoracic wings, beetle head horns, and treehopper prothoracic helmets. It is suggested that these novelties originate from modifications of specific preexisting body parts, namely, the lateral tergum, the clypeolabrum, and the pronotum [1–5], and exhibit remarkable diversifications compared with the corresponding body parts in closely related lineages. Evo-devo studies have revealed that co-option of ancestral gene regulatory networks is one of the driving forces that diversifies those novel traits (e.g., [6–8]). However, how a novel trait originates from a preexisting structure in a constrained insect body plan remains unclear. To address this point, directly developing hemimetabolous insects are suitable models because dramatic remodeling of larval structures through the pupal stage obscures both tissue- and cellular-level relationships between juvenile and adult body parts in derived holometabolous insects (e.g., [9]). Orthopteran species, including grasshoppers and crickets, serve as well-established genetic models of hemimetabolous development with expanding genomic resources and functional genomics tools such as RNAi and genome editing [10]. In orthopteran families, agonistic behaviors between males have facilitated the evolution of novel sexual dimorphisms, such as exaggerated hindlegs in Rhaphidoridae [11] and enlarged mandibles in Anostostomatidae [12], in a lineage-specific manner (Fig. 1A) [11–17]. In Gryllidae, multiple genera exhibit sexual dimorphisms in adult heads, whereas other genera, including the popular laboratory model genera Gryllus and Acheta , form monomorphic head shapes in both sexes (Fig. 1B). In the genus Loxoblemmus , males specifically form a horn-like protrusion on the top of the head and a flattened frontal head region (Fig. 1C), whereas females retain the typical rounded head shape. In Gryllidae, the typical male fighting style involves fencing with antennae, flaring and grappling with mandibles, and a single headbutt or interlocking with mouthparts [15, 18–21]. In addition to these widespread fighting styles, Loxoblemmus species exhibit unique male‒male combat behavior by thrusting the male-specific flattened parts of their heads against each other (Movie S1). This fighting style is specifically described in Loxoblemmus among Gryllidae [15]. The coffin-headed cricket Loxoblemmus equestris , which is distributed on the southwest islands of Japan, Korea, China and Southeast Asia, is a suitable model species because of its ease of breeding and relatively short life cycle (3 months from egg to egg) under laboratory conditions. To understand the mechanisms underlying the origin of novel traits, we investigated regions modified to form the male-specific flattened and horned head in a lineage-specific manner and the temporal regulation behind this modification in L. equestris . Results A male-specific flattened head structure formed in the frons region during the last nymphal instar of L. equestris Hemimetabolous insects exhibit two distinct developmental patterns of novel adult three-dimensional (3D) structures during postembryonic development. The first involves gradual formation across multiple nymphal instars. For example, species-specific pronotal shapes emerge in early nymphal stages and progressively develop into novel adult helmet structures over multiple nymphal instars in membracid species of treehoppers [ 5 ]. The second is concentrated formation during late nymphal development. For example, long rod-like nymphal cerci transform into short forceps during a single molt to reach adulthood in the basal earwig Diplatys flavicollis [ 24 ]. To determine which developmental pattern applies to the novel adult head structure, we observed nymphal head morphologies in L. equestris . This species progresses through nine nymphal instars (N1 to N9) before reaching adulthood (Fig. 2 A). Females can be distinguished from males with developing ovipositors at the eighth abdominal segment from the fourth nymphal instar under a stereomicroscope (Fig. S1 ). Thus, we compared head morphology between males and females from the fourth nymphal instar to adulthood at each stage. We found that no remarkable sexual differentiations appeared in head structures during nymphal instars and that sexually dimorphic head structures abruptly formed in adults (Fig. 2 A). This finding reveals that major head dimorphism occurs during the last nymphal instar in L. equestris . To specify the flattened region in the adult male, we next annotated head landmarks according to Snodgrass (1935) [ 25 ] and compared head morphologies between males and females in L. equestris . We found that the lateral boundary of the flattened region strikingly corresponds to the subocular suture that “extends from the lower angle of each compound eye to the subgenal suture above the anterior mandibular articulation” in crickets [ 25 ] (Fig. 2 B). It thus revealed that the frons, which is defined as “the anterior region to the frontal suture, subocular suture and epistomal suture” [ 25 ], is dramatically modified in males (Fig. 2 C). In addition to the frons, we found notable sexual dimorphisms in the mandible and the clypeus (Fig. S2 A–H). The sexually differential forms of the proximal mandibles at N9 suggest that the sex-specific development begins before the last nymphal instar (Fig. S2 A, B). In adults, the male mandible exhibits a wider proximal region with a darker inner edge, thereby forming a more triangular shape than the female mandible does (Fig. S2 C, D). Another adult dimorphism is the wider clypeus in males than in females (Fig. S2 G, H). In contrast, we detected no obvious differences in the labrum, maxilla, or labium between males and females (Fig. S2 I–P). We focused on the frons dimorphism as an apparent lineage-specific novel trait for further analysis. The male-specific frons structures are prepatterned as epithelial folding from the ninth day of the last nymphal instar To further understand when and how the dimorphic heads formed, we next observed epidermal tissue during the last nymphal instar (N9). We first conducted precise staging to determine the duration of the instar. Since hemimetabolous insects maintain constant feeding even in the preadult stage, we supplied an equivalent amount of food and water to nymphs individually maintained in an incubator during the same time window every day to ensure that the environmental conditions were as identical as possible among the monitored individuals (Fig. 3 A). Under these conditions, the duration of the last nymphal instar was 299.86 ± 3.89 (mean ± standard deviation) hours (Fig. 3 B). We next compared sectioned epidermal tissues along the midline, including the ocellus in the late last instar nymphs of both sexes (Fig. 3 C). The nymphal cuticle and epidermis were attached 192 hours after ecdysis to the last instar nymph (HAE) (Fig. 3 D, E). From 192 to 204 HAE, the nymphal cuticle started detaching from the epidermis, which indicates the onset of apolysis (Fig. 3 F, G). By 216 HAE, the previously smooth epidermis had become corrugated (Fig. 3 H, I). In this stage, males start to form sex-specific epithelial folds around the dorsal side of the median ocellus (Fig. 3 H). By 240 HAE, deeper epithelial folds had been shaped, and the secretion of the adult cuticle had partially begun (Fig. 3 J, K). At this stage, we detected a sexually differential pattern in the epithelial folds around the ventral side of the median ocellus; males presented deeper folds than females did (Fig. 3 J, K). By 264 HAE, the secreted adult cuticle covered the entire epidermal surface, with the sexually differential folding pattern retained (Fig. 3 L, M). In the adult stage, the ventral side of the ocellus became flatter, and the dorsal side protruded in males, whereas the nymphal shapes were mostly retained in corresponding regions of females (Fig. 3 N, O). These observations revealed the formation of male-specific intensive epithelial folding after 204 HAE. To further investigate the sexually differential epithelial folds in three dimensions, we reconstructed the frons epidermis at 240 HAE by using micro-CT (Fig. 4 A). Males formed more complex two-dimensional (2D) epithelial patterns than females did at both the dorsal and ventral sides of the median ocellus and around subocular sutures at this stage (Fig. 4 B–D, F–H). In contrast, the posterior epidermis in the dorsal head was not obviously different between the sexes, suggesting that differential epithelial folding underlies the sexually dimorphic head shapes (Fig. 4 E, I). To quantify differences in epithelial folding patterns between males and females, we measured the density and furrow depth in right, medial, and left sagittal sections at the ventral frons region (Fig. S3A). Both the density and the furrow depth significantly differed between males and females in the right and left sections (Fig. 4 J, K). In the medial section, the furrow depth significantly differed, although we found no significant difference in the furrow density between the sexes (Fig. 4 J, K). These analyses collectively specified the frons as the modified head region that gives rise to both flat and protruding shapes in L. equestris . The metamorphic gene network controls the timing of dimorphic head development The metamorphic gene network (MGN), which comprises Krüppel-homolog 1 ( Kr-h1 ), broad ( br ), and Ecdysone induced protein 93F ( E93 ), is proposed as an evolutionarily conserved central regulator of insect metamorphosis [ 26 ]. In G. bimaculatus , Kr-h1 and br play major roles in suppressing adult metamorphosis, whereas E93 promotes the nymph-to-adult transition [ 27 ]. Since the male-specific head structure formed during the preadult stage, we hypothesized that the MGN is involved in the temporal regulation of sexually dimorphic head formation in L. equestris . To test this hypothesis, we first examined the temporal expression patterns of the three genes in the frons epithelial tissues of L . equestris from the penultimate nymphal stage to adulthood. The expression level of E93 was relatively low during the penultimate nymphal instar, gradually increased during the last nymphal instar, and reached its peak on the final day of the nymph (Fig. 5 A). Conversely, Kr-h1 was highly expressed during the penultimate instar, dramatically decreased in expression on the second day of the last nymphal instar, and maintained a low level thereafter (Fig. 5 B). The br transcript presented no explicit temporal pattern during the late nymphal stage (Fig. 5 C). These temporal gene expression patterns are similar to the reported patterns in the abdominal epidermis of G. bimaculatus [ 27 ], which suggests a conserved function of the MGN in regulating metamorphosis between the two cricket species. To further understand the roles of these genes in male-specific head formation in L. equestris , we next knocked down their transcript levels via nymphal RNAi. To implement the nymphal RNAi method in L. equestris , we injected double-stranded (ds) RNA targeting ebony , which synthesizes N-β-alanyl-dopamine for light body coloration in G . bimaculatus [ 28 ], into penultimate instar nymphs. We successfully obtained dark-colored adults after ebony dsRNA injection, which validated the effect of RNAi-mediated gene knockdown in this species (Fig. S4, Table S3). RNAi treatments targeting E93 , Kr-h1 , or br caused temporal shifts in adult metamorphosis, whereas all the control individuals became adults following N9 (Fig. 6 A–K, A’–G’, Table S4). All L. equestris nymphs injected with E93 dsRNA at N8 failed to metamorphose into adults after N9. Among the surviving crickets, 38.9% and 44.4% underwent supernumerary nymphal molting and became larger adults after N10 and N11, respectively (Fig. 6 C–E, C’–E’, I, Table S4). Notably, these supernumerary nymphs maintained the normal nymphal head proportion at N10 or N11, and the male-specific adult head shape first formed after metamorphic molts into adults (Fig. 6 C–E, C’–E’). Compared with the control adults, the resulting larger adults presented overgrown wings (Fig. 6 I). In contrast, RNAi treatments targeting Kr-h1 or br at N4 caused precocious adult metamorphosis after N7 or N8 (Fig. 6 F, G, F’, G’, J, K; Table S4), although the penetrance was relatively low (22.3% for Kr-h1 and 17.4% for br ). Despite being smaller in body size, the precocious adults presented male-specific adult head shapes, whereas their wings were disproportionate in shape (Fig. J, K). To analyze the effect of the shifted timing of adult metamorphosis on head proportion, we measured the ratio of head width to body length (Fig. S5A, B). Compared with those in the control group, we found no differences in the head-to-body ratio in adults that experienced supernumerary nymphal instars after E93 RNAi treatment, which indicates proportionate head formation (Fig. 6 L). We note that incomplete molting or failure to unfold wings in most E93 RNAi individuals prohibited similar size measurements of the wings (Fig. S5C). These RNAi phenotypes confirmed the conserved functions of the MGN in L. equestris and revealed that sex-specific novel head development is under the temporal control of the gene network. Discussion Male-specific novel head shapes are prepatterned as epithelial folding in L. equestris Recent studies have demonstrated that epithelial folding prepatterns sophisticated 3D structures in insects. For example, the protruding beetle horn is formed by the 3D expansion of patterned 2D epithelial furrows in the larval primordium [ 29 ]. Alterations in the furrow pattern, depth, and density affect the final shape of adult horns [ 30 ]. In L . equestris , males form denser, deeper furrows with more complex 2D patterns than females do in the frons epithelial cell sheets during the preadult stage (Figs. 3 , 4 ). These data support the idea that sex-specific modifications in epithelial folding patterns led to the evolution of the novel head from an ancestral shape in Gryllidae. Experimental and theoretical approaches have located the head region that undergoes differential growth to elaborate the protruding horn in the frons region of beetles [ 2 – 4 , 31 ]. Differential forms of modified frons structures between the cricket and horned beetles reveal variation in modifications, ranging from flat surfaces to large protrusions, of a homologous region in insects. In both cases, modifications in frons shape are associated with male‒male combat behavior [ 15 , 32 ]. The difference in fighting styles could have driven diversification of differential forms from the homologous region or vice versa. Sex-specific epithelial folding patterns develop through adult metamorphosis in L. equestris We specified the timing of sexually dimorphic development of epithelial folding from 204 to 240 hours after ecdysis to the preadult instar stage in L . equestris (Fig. 3 ). Critically, head sexual dimorphism occurs after adult metamorphosis despite the timing shifts caused by gene knockdown (Fig. 6 , Table S4). These data demonstrate that sex-specific patterns of epithelial folding are formed during adult metamorphosis, which is controlled by the MGN, and suggest that E93 is a major upstream regulator that determines the timing of dimorphic epithelial patterning. E93 and Kr-h1 presented contrasting expression patterns in the frons tissues of L . equestris last instar nymphs (Fig. 5 ). Targeted knockdown of E93 , Kr-h1 and br caused delayed and precocious formations of head dimorphism, respectively (Fig. 6 ). Antagonistic regulation between Kr-h1 and E93 is widely conserved in both hemi- and holometabolous insects [ 26 , 33 ]. Indeed, knockdown of Kr-h1 , and br , causes early upregulation of the E93 transcript level in another cricket species, G . bimaculatus [ 27 ]. In Drosophila , the temporal expression of E93 broadly provides competence for target enhancers to respond to spatial cues through controlling chromatin accessibility [ 34 ]. Taken together, it is conceivable that E93 could bind enhancers regulating gene expression in the male frons during adult metamorphosis, thereby locally relaxing the body plan constraint to permit dramatic transformation of the preexisting ancestral juvenile shape into the derived adult form, which is likely adaptive for agonistic behavior. The final molt as a driver of the evolution of novel traits in insects Whereas the male-specific head shape formed almost proportionately, the final size of the wings was highly disproportionate following timing shifts in metamorphosis caused by gene knockdowns, as reported in other hemimetabolous species (Fig. 6 ; e.g., [ 27 , 35 , 36 ]). The contrasting phenotypes between the head and wings underscore the robustness of the head proportion against timing shifts in development. In hemimetabolous insects, wing primordia externally appear as wing pads and undergo anisometric growth during nymphal development [ 1 , 37 ]. Paleozoic fossils and recent evo-devo studies suggest that wings originated as lateral tergal outgrowths, such as those formed in palaeodictyopteran nymphs, modified during postembryonic development to form functional adult wings and their serial homologs [ 38 ]. It has been proposed that those wing precursors play preadaptive roles, such as thermoregulation, surface-skimming behavior, aerial body control, and respiration [ 38 , 39 ]. Precursors of the novel treehopper helmets on the prothorax externally appear in early nymphs [ 5 ], following a developmental pattern similar to that of wings. The shared developmental pattern suggests that gradual modifications during juvenile development serve as a general path to generate evolutionary novelty from the constrained insect body plan. The concentrated development of adult head dimorphism in L. equestris demonstrates another path for the evolution of novelties from preexisting structures. The male-specific frons shape, which is used for male‒male combat behavior in adulthood, starts developing during a single preadult instar, thereby robustly forming a proportionate shape without specific early-forming precursors used in preadaptive roles. This highlights the evolution of the final molt and metamorphic life cycle in insects [ 40 ], which clearly splits the juvenile and adult stages by metamorphosis. The E93 expression level reflects the differential patterns of juvenile-to-adult transitions before and after the evolution of concentrated adult metamorphosis; the E93 level is mostly constant during the juvenile and adult stages in the ancestral ametabolous insect Thermobia domestica , whereas it exhibits a highly compressed peak during the late juvenile period in hemimetabolous insects [ 41 ]. Consistent with the expression pattern, it has been suggested that E93 acts on cuticle development throughout the nymphal stage in T. domestica [ 42 ]. In contrast, concentrated E93 expression enables centralized regulation of the expression of a broad range of genes for terminal differentiation of adult structures during late juvenile development in derived hemi- and holometabolous insects [ 43 – 45 ]. Our study reveals the direct link between the development of a lineage-specific 3D structure and the derived concentrated metamorphosis regulated by the MGN, suggesting the contribution of the evolution of a unique life cycle to the explosive morphological diversity of the hexapod lineage (Fig. 7 ). Conclusions In this study, using a cricket with a novel head, we found that the ancestral shape directly transforms into the novel shape through the final molt to adulthood, even after artificial timing shifts of metamorphosis, by creating sex-specific epithelial folding. This finding provides direct evidence that the unique metamorphic life cycle has facilitated the evolution of novelties in lineage- and sex-specific manners, thereby contributing to the morphological diversity of insects. Methods Cricket rearing and staging The coffin-headed cricket Loxoblemmus equestris (Saussure 1877) used in this study was derived from Iriomote Island, Okinawa, Japan. All crickets were maintained at 28–30°C and 30% relative humidity under a 12 L:12 D photoperiod in an air-conditioned room and fed artificial goldfish food (Spectrum Brands). For precise staging, cricket nymphs were individually maintained in a plastic cup placed in an incubator set at 28°C under 12 L:12 D. We supplied five pellets of artificial goldfish food and water daily during 5:00–8:00 PM. The water was supplied from a paper string to the maximum extent possible without it seeping out. Nymphs were isolated from the rearing group before the target stage. The timing of molting was monitored by taking time-lapse photographs every 3 minutes. The same staging method was used for cryosectioning and quantitative RT-PCR. Scanning electron microscopy Nonfixed head samples were mounted on sample stubs using carbon double-sided tape (NISSHIN EM, 7323). All images were captured at a voltage of 15 kV using a tabletop scanning electron microscope system (Miniscope® TM4000PlusII; HITACHI). Epithelial tissue histology Frozen sections were prepared according to the Kawamoto method [ 46 ]. Crickets were anesthetized on ice and dissected in phosphate-buffered saline (PBS). The whole-head samples were embedded in Super Cryoembedding Medium (SECTION-LAB, R060202). The embedded heads were sectioned at a thickness of 7 µm by using a cryostat (Leica CM3050 S; Leica Biosystems). Adhesive films (cryofilm type II C (9); SECTION-LAB, R050928) were used to capture cryosections. The sectioned tissues were fixed in 4% paraformaldehyde (PFA) for 5 minutes and then washed twice with PBS for 5 minutes each. The tissue samples were stained with Carrazzi’s hematoxylin solution (SECTION-LAB, R060213) for 5 minutes and rinsed with running tap water for 5 minutes to develop color. These samples were subsequently stained with 0.2% eosin Y solution (SECTION-LAB, R060214) for 30 seconds, followed by a 30 second rinse in running tap water. The stained sections were then washed with 100% ethanol to remove residual eosin and mounted in 50% glycerol. X-ray micro-computed tomography (CT) analysis Fixation, washing, and staining steps were performed on a rotary platform shaker at room temperature. Last-instar nymphs of L. equestris at 240 hours after molting were anesthetized on ice for head dissection. The head samples were fixed in 4% PFA overnight and then washed three times in PBS for 10 minutes each. Fixed samples were stained with 25% Lugol solution for 48 hours, followed by washing twice with PBS overnight. The stained samples were embedded in paraffin and scanned using an X-ray micro-CT device (SKYSCAN 1272 CMOS Edition; BRUKER) at a voltage peak of 60 kV and a current of 80 µA. The samples were rotated 360 degrees in steps of 0.08 degrees, generating 4500 projection images for each of the male and female samples. The micro-CT data were reconstructed at isotropic resolutions of 1.5 µm and 1.35 µm for the male and female samples, respectively, via NRecon software (v2.0.0.5, Micro Photonics Inc.). Three-dimensional (3D) tomographic images were obtained and processed using VGSTUDIO MAX software (Volume Graphics). Quantitative analysis of epidermal folding patterns The number of furrows and depth of epidermal folding were measured according to Adachi (2020) [ 30 ]. The cross-sectional images used to quantify furrow depths were captured as CT images using VGSTUDIO MAX software. Brightness adjustments and depth measurements of the furrows were performed using ImageJ. The “SegmentMeasure” plug-in of ImageJ was used to measure the length of the epidermis. De novo transcriptome assembly To obtain the transcript sequences of L. equestris , we assembled short RNA-seq reads. We pooled total RNA extracted from whole embryos at 3, 7, and 12 days after egg laying and whole heads of individual male and female last-instar nymphs at 2, 5, 10, and 11 days after molting. Total RNA extraction was performed using TRIzol reagent (Thermo Fischer Scientific, 15596018), followed by phenol‒chloroform extraction. Subsequent purification was conducted using RNeasy columns (RNeasy® Micro Kit (50); QIAGEN, 172045635). The purified total RNA was then submitted to Azenta Life Sciences for library preparation using NEBNext Ultra II Directional RNA Library Prep Kit for Illumina. Sequencing was carried out on an Illumina NovaSeq platform, generating 150 bp paired-end reads. Poor-quality reads and adapter sequences were discarded with Cutadapt (v4.4) with default parameter settings. Trinity (v2.13.2) was used for de novo transcriptome assembly. BLAST search for orthologous genes We searched for orthologs of Krüppel-homolog 1 ( Kr-h1 ), broad ( br ), and Ecdysone induced protein 93F ( E93 ) in the L. equestris transcriptome using public partial cDNA sequences from G. bimaculatus ( E93 , LC476893; Kr-h1 , LC476894; br , LC476892) as query sequences for the BLASTn program. We confirmed that the identified transcript sequences of L. equestris presented the best hits to the corresponding genes of D. melanogaster [ 47 ] via BLASTx searches. Quantitative RT-PCR (qRT-PCR) Total RNA was extracted from frons epidermal tissues, including the cuticles of penultimate and last-instar male nymphs. The tissues were dissected in PBS after the crickets were anesthetized on ice. All tissue samples were collected in TRIzol reagent and stored at -80°C until use. Total RNA was extracted according to the manufacturer’s instructions. The extracted RNA was reverse transcribed to cDNA using ReverTra Ace® qPCR RT Master Mix with gDNA Remover (TOYOBO, FSQ-301) according to the manufacturer’s instructions. qRT-PCR was performed using THUNDERBIRD SYBR qPCR mix (TOYOBO, QPS-201) on QuantStudio™ 3 Real-Time PCR System (Applied Biosystems). The primers used for qRT-PCR are listed in Table S1 . The qRT-PCR program was as follows: 95°C for 5 min, 95°C for 30 s, and 60°C for 45 s for 40 cycles with each primer at a concentration of 10 µM. The Ribosomal protein L32 ( RpL32 ) gene was used as a reference gene for qRT-PCR analysis. All qRT-PCRs were performed in triplicate as technical replicates. The relative quantity of transcripts was calculated by the 2-ΔΔCt method (Livak & Schmittggen, 2001). RNA interference (RNAi)-mediated gene knockdown DNA templates for double-stranded RNA (dsRNA) synthesis were amplified from late-stage embryo cDNA using primers with the T7 promoter sequence at the 5′ end listed in Table S2 . To obtain a sufficient amount of DNA template for in vitro transcription, we performed PCR twice; first, AmpliTaq Gold™ 360 DNA Polymerase (Applied Biosystems, 4398823) was used, and then ExTaq (Takara, RR001B) was used for all the targets except for egfp . The first PCR product is used as a template for the second PCR. For egfp , the DNA template was amplified from plasmid DNA using ExTaq. RNA was transcribed from a DNA template using the T7 RiboMAX Express Large Scale RNA System (Promega, P1320) and purified using phenol/chloroform/isoamyl alcohol (125:24:1) (Merck, 77619-100ML). The transcribed RNA was resuspended in annealing buffer [10 mM Tris-Cl (pH 7.5), 1 mM EDTA, and 5 mM NaCl/DEPC-treated H 2 O] and annealed by gradual cooling from 95°C to 25°C on a programmed thermal cycler to make dsRNA. Annealed dsRNA solutions were aliquoted and stored at -80°C. DsRNA solution was injected into the lateral ventral side of the intersegmental membrane between nymphal abdominal segments within 24 hours postecdysis. For E93 and Kr-h1 and br knockdown, 12 µg and 6 µg of dsRNA solution, respectively, were injected into penultimate and fourth instar nymphs of males. For all the RNAi experiments, an equivalent amount of egfp dsRNA was injected as a negative control. Declarations Acknowledgments We thank Mr. Hosei Wada for developing the “SegmentMeasure” plug-in of ImageJ, Ms. Junko Morita for her technical support, and Drs. Taro Nakamura, Toshiya Ando and Naoki Matsuda for their helpful discussion. Author contributions MY: all research performance, manuscript preparation. SM: X-ray micro-CT analysis, review the manuscript. TN: X-ray micro-CT analysis, review the manuscript. TD: review the manuscript. TO: research design, review the manuscript. Funding This work was supported by the Japan Society for the Promotion of Science KAKENHI [grant numbers 23K18026 (TO), 24H00511 (TD)] and the NIBB Collaborative Research Program [grant numbers 22NIBB336, 23NIBB343, 24NIBB326 (TO)]. Availability of data and materials All data are included in the main manuscript and Additional file 1. Ethics approval and consent to participate Not applicable. Consent for publication Not applicable. Competing interests The authors declare no competing interests. References Ohde T, Mito T, Niimi T. A hemimetabolous wing development suggests the wing origin from lateral tergum of a wingless ancestor. Nat Commun. 2022;13(1):979. Zattara EE, Busey HA, Linz DM, Tomoyasu Y, Moczek AP. 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Agonistic behaviour in male and female field crickets, Gryllus bimaculatus, and how behavioural context influences its expression. Anim Behav. 1995;49(6):1491-501. Tachon G, Murray AM, Gray DA, Cade WH. Journal of Insect Behavior. 1999;12(4):533-43. Hofmann HA, Schildberger K. Assessment of strength and willingness to fight during aggressive encounters in crickets. Anim Behav. 2001;62(2):337-48. Jang Y, Gerhardt HC, Choe JC. A comparative study of aggressiveness in eastern North American field cricket species (genus Gryllus). Behav Ecol Sociobiol. 2008;62(9):1397-407. Chintauan-Marquier IC, Legendre F, Hugel S, et al. Laying the foundations of evolutionary and systematic studies in crickets (Insecta, Orthoptera): a multilocus phylogenetic analysis. Cladistics. 2016;32(1):54-81. Song H, Béthoux O, Shin S, et al. Phylogenomic analysis sheds light on the evolutionary pathways towards acoustic communication in Orthoptera. Nat Commun. 2020;11(1):4939. Shimizu S, Machida R. Reproductive biology and postembryonic development in the basal earwig Diplatys flavicollis (Shiraki) (Insecta: Dermaptera: Diplatyidae). Arthropod Syst Phylogeny. 2011;69(2):83-97. Snodgrass RE. Head: Structure of the Definitive Insect Head. In: Snodgrass RE, editor. Principles of insect morphology. New York: McGraw-Hill; 1935. p. 109-110. Martín D, Chafino S, Franch-Marro X. How stage identity is established in insects: the role of the Metamorphic Gene Network. Curr Opin Insect Sci. 2021;43:29-38. Ishimaru Y, Tomonari S, Watanabe T, Noji S, Mito T. Regulatory mechanisms underlying the specification of the pupal-homologous stage in a hemimetabolous insect. Phil Trans R Soc B. 2019;374(1783):20190225. Inoue S, Watanabe T, Hamaguchi T, et al. Combinatorial expression of ebony and tan generates body color variation from nymph through adult stages in the cricket, Gryllus bimaculatus. PLoS ONE. 2023;18(5):e0285934. Matsuda K, Gotoh H, Tajika Y, et al. Complex furrows in a 2D epithelial sheet code the 3D structure of a beetle horn. Sci Rep. 2017;7(1):13939. Adachi H, Matsuda K, Niimi T, et al. Genetical control of 2D pattern and depth of the primordial furrow that prefigures 3D shape of the rhinoceros beetle horn. Sci Rep. 2020;10(1):18687. Morikawa K, Morita S, Sakura K, et al. Unveiling the role of differential growth in 3D morphogenesis: An inference method to analyze area expansion rate distribution in biological systems. J Theor Biol. 2023;575:111650. McCullough EL, Tobalske BW, Emlen DJ. Structural adaptations to diverse fighting styles in sexually selected weapons. Proc Natl Acad Sci U S A. 2014;111(40):14484-8. Ureña E, Chafino S, Manjón C, et al. The Occurrence of the Holometabolous Pupal Stage Requires the Interaction between E93, Krüppel-Homolog 1 and Broad-Complex. PLoS Genet. 2016;12(5):e1006020. Nystrom SL, Niederhuber MJ, McKay DJ. Expression of E93 provides an instructive cue to control dynamic enhancer activity and chromatin accessibility during development. Development. 2020;147(6):dev181909. Konopova B, Smykal V, Jindra M. Common and Distinct Roles of Juvenile Hormone Signaling Genes in Metamorphosis of Holometabolous and Hemimetabolous Insects. PLoS ONE. 2011;6(12):e28728. Lozano J, Belles X. Conserved repressive function of Krüppel homolog 1 on insect metamorphosis in hemimetabolous and holometabolous species. Sci Rep. 2011;1(1):163. Erezyilmaz DF, Riddiford LM, Truman JW. The pupal specifier broad directs progressive morphogenesis in a direct-developing insect. Proc Natl Acad Sci U S A. 2006;103(18):6925-30. Ohde T, Prokop J. The transition to flying insects: lessons from evo-devo and fossils. Curr Opin Insect Sci. 2025;68:101332. Treidel LA, Deem KD, Salcedo MK, et al. Insect Flight: State of the Field and Future Directions. Integr Comp Biol. 2024;64(2):533-55. Belles X. The innovation of the final moult and the origin of insect metamorphosis. Phil Trans R Soc B. 2019;374(1783):20180415. Fernandez-Nicolas A, Machaj G, Ventos-Alfonso A, et al. Reduction of embryonic E93 expression as a hypothetical driver of the evolution of insect metamorphosis. Proc Natl Acad Sci U S A. 2023;120(7):e2216640120. Bai Y, Lv YN, Zeng M, et al. E93 is indispensable for reproduction in ametabolous and hemimetabolous insects. Development. 2024;151(20):dev202518. Zeng M, Yan ZY, Lv YN, et al. Molecular basis of E93-dependent tissue morphogenesis and histolysis during insect metamorphosis. Insect Biochem Mol Biol. 2025;177:104249. Uyehara CM, Nystrom SL, Niederhuber MJ, et al. Hormone-dependent control of developmental timing through regulation of chromatin accessibility. Genes Dev. 2017;31(9):862-75. Cruz J, Ureña E, Iñiguez LP, et al. E93 controls adult differentiation by repressing broad in Drosophila. Proc Natl Acad Sci U S A. 2024;121(51):e2403162121. Kawamoto T, Kawamoto K. Preparation of Thin Frozen Sections from Nonfixed and Undecalcified Hard Tissues Using Kawamoto’s Film Method. In: Hilton MJ, editor. Skeletal Development and Repair. Totowa, NJ: Humana Press; 2014. p. 149-64. (Methods in Molecular Biology; vol. 1130). Available from: https://link.springer.com/10.1007/978-1-62703-989-5_11 Hoskins RA, Carlson JW, Wan KH, et al. The Release 6 reference sequence of the Drosophila melanogaster genome. Genome Res. 2015;25(3):445-58. Additional Declarations No competing interests reported. Supplementary Files MovieS1.mp4 yonedaevodevoSI.docx Cite Share Download PDF Status: Published Journal Publication published 16 Jul, 2025 Read the published version in Developmental Biology Advances → Version 1 posted Editorial decision: Revision requested 02 Jun, 2025 Reviews received at journal 25 May, 2025 Reviews received at journal 06 May, 2025 Reviewers agreed at journal 12 Apr, 2025 Reviewers agreed at journal 09 Apr, 2025 Reviewers agreed at journal 31 Mar, 2025 Reviewers invited by journal 26 Mar, 2025 Editor assigned by journal 26 Mar, 2025 Submission checks completed at journal 26 Mar, 2025 First submitted to journal 25 Mar, 2025 You are reading this latest preprint version Research Square lets you share your work early, gain feedback from the community, and start making changes to your manuscript prior to peer review in a journal. As a division of Research Square Company, we’re committed to making research communication faster, fairer, and more useful. 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Also discoverable on Platform About Our Team In Review Editorial Policies Advisory Board Help Center Resources Author Services Accessibility API Access RSS feed Manage Cookie Preferences © Research Square 2026 | ISSN 2693-5015 (online) Privacy Policy Terms of Service Do Not Sell My Personal Information {"props":{"pageProps":{"initialData":{"identity":"rs-6301946","acceptedTermsAndConditions":true,"allowDirectSubmit":false,"archivedVersions":[],"articleType":"Research Article","associatedPublications":[],"authors":[{"id":440691566,"identity":"f16b504b-c3db-48bc-a75e-4b092782579c","order_by":0,"name":"Mizuho Yoneda","email":"","orcid":"","institution":"Kyoto University","correspondingAuthor":false,"prefix":"","firstName":"Mizuho","middleName":"","lastName":"Yoneda","suffix":""},{"id":440691568,"identity":"ef28f89c-7b15-4b93-859c-5fc2c07859a7","order_by":1,"name":"Shinichi Morita","email":"","orcid":"","institution":"National Institute for Basic Biology","correspondingAuthor":false,"prefix":"","firstName":"Shinichi","middleName":"","lastName":"Morita","suffix":""},{"id":440691570,"identity":"3318b5ac-e4b4-4e3a-8f82-d3638e0de5bf","order_by":2,"name":"Teruyuki Niimi","email":"","orcid":"","institution":"National Institute for Basic Biology","correspondingAuthor":false,"prefix":"","firstName":"Teruyuki","middleName":"","lastName":"Niimi","suffix":""},{"id":440691571,"identity":"f4822a96-55a6-464b-8aa8-25c1ddc0dbec","order_by":3,"name":"Takaaki Daimon","email":"","orcid":"","institution":"Kyoto University","correspondingAuthor":false,"prefix":"","firstName":"Takaaki","middleName":"","lastName":"Daimon","suffix":""},{"id":440691572,"identity":"a4cb62df-812f-4c89-a81d-99af2a166094","order_by":4,"name":"Takahiro Ohde","email":"data:image/png;base64,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","orcid":"","institution":"Kyoto University","correspondingAuthor":true,"prefix":"","firstName":"Takahiro","middleName":"","lastName":"Ohde","suffix":""}],"badges":[],"createdAt":"2025-03-25 08:53:42","currentVersionCode":1,"declarations":"","doi":"10.21203/rs.3.rs-6301946/v1","doiUrl":"https://doi.org/10.21203/rs.3.rs-6301946/v1","draftVersion":[],"editorialEvents":[{"content":"https://doi.org/10.1186/s13227-025-00249-3","type":"published","date":"2025-07-16T16:05:34+00:00"}],"editorialNote":"","failedWorkflow":false,"files":[{"id":80346917,"identity":"2405ebd6-4d90-4604-b4a8-fcc6984d6159","added_by":"auto","created_at":"2025-04-10 20:18:17","extension":"png","order_by":1,"title":"Figure 1","display":"","copyAsset":false,"role":"figure","size":1051265,"visible":true,"origin":"","legend":"\u003cp\u003eNovel head morphologies in Gryllidae. (A) Simplified phylogenetic tree of Ensifera. Male agonistic behaviors have been reported in the families Anostostomatidae, Tettigoniidae, Rhaphidphoridae, Phalangopsidae and Gryllidae. In the family Gryllidae, species in three genera (indicated by stars on branches) exhibit novel morphologies specific to the male head. The phylogenetic tree was drawn according to Chintauan‐Marquier (2016) and Song (2020) [22, 23]. Frontal (B–E) and lateral (B’–E’) views of male heads of \u003cem\u003eGryllus bimaculatus\u003c/em\u003e (B, B’), \u003cem\u003eLoxoblemmus doenitzi\u003c/em\u003e (C, C’), \u003cem\u003eSciobia burmeister\u003c/em\u003e (D, D’) and \u003cem\u003eVelarifictorus longifrons\u003c/em\u003e (E, E’). Prominent protrusions on the top of the head (red arrowheads) and flattening of the frontal area (blue arrowheads) are formed in three species, whereas \u003cem\u003eG. bimaculatus\u003c/em\u003e shows typical ancestral rounded heads.\u003c/p\u003e\n\u003cp\u003ePhoto credits: (D) Goro García Moreno, Saber Animal (D’) Thijs Valkenburg, Associação Vita Nativa (E, E’) Kai Schütte, saltatoria.info\u003c/p\u003e","description":"","filename":"floatimage1.png","url":"https://assets-eu.researchsquare.com/files/rs-6301946/v1/d3df50b2fed4a74270c5155a.png"},{"id":80346918,"identity":"ac31f82f-626a-4608-bd32-d0270f5949ae","added_by":"auto","created_at":"2025-04-10 20:18:17","extension":"png","order_by":2,"title":"Figure 2","display":"","copyAsset":false,"role":"figure","size":1437110,"visible":true,"origin":"","legend":"\u003cp\u003eHead sexual dimorphism in \u003cem\u003eLoxoblemmus equestris. \u003c/em\u003e(A) Frontal and lateral views of heads from the fourth nymphal instar to adulthood. Adult males form a flat shape in the frontal area (blue arrowhead), whereas nymphs and adult females exhibit typical ancestral rounded heads. Scale bars indicate 1 mm. (B) The subocular sutures of the last instar nymphs and adults (white arrowheads). Adult males specifically form a flat shape at the anterior of the subocular suture. (C) Head morphological annotations in \u003cem\u003eL. equestris\u003c/em\u003e. Landmarks are according to the annotation of \u003cem\u003eGryllus assimilis\u003c/em\u003e (Snodgrass, 1935). e, compound eye; o, ocellus; md, mandible; clp, clypeus; lm, labrum; es, epistomal suture; fs, frontal suture; sos, subocular suture.\u003c/p\u003e","description":"","filename":"floatimage2.png","url":"https://assets-eu.researchsquare.com/files/rs-6301946/v1/27ca5c9b45693d989d8c6f37.png"},{"id":80346919,"identity":"9ad6a53c-a3af-4b9c-b1ce-89786f6920e8","added_by":"auto","created_at":"2025-04-10 20:18:17","extension":"png","order_by":3,"title":"Figure 3","display":"","copyAsset":false,"role":"figure","size":1244132,"visible":true,"origin":"","legend":"\u003cp\u003eHead epithelial tissue observation in the late last nymphal instar.\u003cstrong\u003e \u003c/strong\u003e(A) Individual rearing. (B) Staging of N8 and N9. Boxplots are presented to display the median, as well as the upper and lower quantiles, minimum values, and maximum values. (n=27). (C) Sectioned plane. Heads were sagittally sectioned along the midline. The bold black square indicates the region shown in D–O. (D–O) Temporal section images of the late last nymphal instars. Scale bars indicate 0.5 mm.\u003c/p\u003e","description":"","filename":"floatimage3.png","url":"https://assets-eu.researchsquare.com/files/rs-6301946/v1/660436da4c7c313d62a16189.png"},{"id":80346923,"identity":"19d6bc99-7539-4ff6-9add-94e392bde3aa","added_by":"auto","created_at":"2025-04-10 20:18:17","extension":"png","order_by":4,"title":"Figure 4","display":"","copyAsset":false,"role":"figure","size":1111333,"visible":true,"origin":"","legend":"\u003cp\u003eQuantification of epithelial folding patterns. (A) Regions where the epidermis at 240 HAE was reconstructed using micro-CT. In the reconstructed image, the head capsule was removed \u003cem\u003ein silico\u003c/em\u003e to obtain an epidermis image. (B–I) Reconstructed epidermis images of the ventral (B, F) and dorsal (C, G) sides of the median ocellus and regions around the subocular suture (D, H) and posterior side of the head (E, I). Arrows indicate M-shaped patterns that are specifically observed in males (B). (J) Comparison of furrow densities. The number of grooves was divided by the total length of the epidermis in a measured area to calculate the furrow density in a section. Means and standard deviations of values from three individuals are shown. (K) Comparison of furrow depths. The measured depth was divided by the head size and multiplied by 1,000 to calculate a normalized depth. A box and whisker plot summarizing the normalized values from three individuals is shown. Comparisons between males and females were made via Welch's t-test. * P \u0026lt; 0.05, n.s. = not significant.\u003c/p\u003e","description":"","filename":"floatimage4.png","url":"https://assets-eu.researchsquare.com/files/rs-6301946/v1/349ae0d7f0e74919d7d49ff6.png"},{"id":80346922,"identity":"a71e9fda-d497-4dcf-ad03-3e2cf910b119","added_by":"auto","created_at":"2025-04-10 20:18:17","extension":"png","order_by":5,"title":"Figure 5","display":"","copyAsset":false,"role":"figure","size":467510,"visible":true,"origin":"","legend":"\u003cp\u003eTemporal gene expression profile in late nymphal instars. Relative amounts of (A) \u003cem\u003eE93\u003c/em\u003e, (B)\u003cem\u003e Kr-h1\u003c/em\u003e, and (C)\u003cem\u003e br \u003c/em\u003emRNAs in N8, N9 and adult. The expression levels are relative to those on day 0 of N8 and are normalized to \u003cem\u003eRpL32\u003c/em\u003e. Three replicates were performed at each time point.\u003c/p\u003e","description":"","filename":"floatimage5.png","url":"https://assets-eu.researchsquare.com/files/rs-6301946/v1/2198029a851b5861440cb24b.png"},{"id":80346928,"identity":"753e0856-c87c-43f4-a6ca-d75312dda05d","added_by":"auto","created_at":"2025-04-10 20:18:17","extension":"png","order_by":6,"title":"Figure 6","display":"","copyAsset":false,"role":"figure","size":1175339,"visible":true,"origin":"","legend":"\u003cp\u003eFunctional analyses of \u003cem\u003eE93\u003c/em\u003e, \u003cem\u003eKr-h1\u003c/em\u003e and \u003cem\u003ebr\u003c/em\u003e in \u003cem\u003eL. equestris\u003c/em\u003e\u003cem\u003e\u003cstrong\u003e. \u003c/strong\u003e\u003c/em\u003e(A–G, A’–G’) Nymphal and adult heads. Scale bars indicate 1 mm. (H–K) Adult whole-body and wing phenotypes (filled with magenta). Scale bars indicate 5 mm. (L) The ratio of head width to body size. Each plot in \u003cem\u003eegfp\u003c/em\u003e, \u003cem\u003eE93\u003c/em\u003e (N10), and \u003cem\u003eE93\u003c/em\u003e (N11) represents the phenotypes of adult metamorphosis following N9, N10, and N11, respectively. Comparisons between the groups were evaluated by Welch's t-test with the Bonferroni adjustment. The means and standard deviations are shown. n.s. = not significant. (M) Summary of nymphal RNAi targeting \u003cem\u003eE93\u003c/em\u003e, \u003cem\u003eKr-h1\u003c/em\u003e and \u003cem\u003ebr\u003c/em\u003e.\u003c/p\u003e","description":"","filename":"floatimage6.png","url":"https://assets-eu.researchsquare.com/files/rs-6301946/v1/cd7587e277e47aa4df2f58d4.png"},{"id":80346931,"identity":"2cbc0f4c-4446-4ffd-8f05-179c3022a180","added_by":"auto","created_at":"2025-04-10 20:18:17","extension":"png","order_by":7,"title":"Figure 7","display":"","copyAsset":false,"role":"figure","size":579834,"visible":true,"origin":"","legend":"\u003cp\u003eNovel trait development through concentrated insect metamorphosis. (A) An ancestral ametabolous insect (\u003cem\u003eT\u003c/em\u003e. \u003cem\u003edomestica\u003c/em\u003e) presents a constant \u003cem\u003eE93\u003c/em\u003eexpression level and continues molting cycles, maintaining almost the same form throughout postembryonic development [41]. The gradually developing female ovipositors are in magenta. (B) In hemimetabolous insects, including \u003cem\u003eL. equestris\u003c/em\u003e,\u003cem\u003e E93\u003c/em\u003e dramatically increased at the late nymphal stage and undergo metamorphosis accompanied by terminal differentiations of adult structures through a final molt. The evolution of the unique life cycle with concentrated metamorphosis through a final molt could have facilitated novel trait development by locally modifying preexisting ancestral body parts. Arrowheads indicate the abruptly transformed male head. Gradually developing wings and female ovipositors are shown in magenta.\u003c/p\u003e","description":"","filename":"floatimage7.png","url":"https://assets-eu.researchsquare.com/files/rs-6301946/v1/449a2b74a70eab748e8866b5.png"},{"id":87220842,"identity":"4e5ec24d-a5b0-4931-be4b-ba051609cf95","added_by":"auto","created_at":"2025-07-21 16:13:04","extension":"pdf","order_by":0,"title":"","display":"","copyAsset":false,"role":"manuscript-pdf","size":8038223,"visible":true,"origin":"","legend":"","description":"","filename":"manuscript.pdf","url":"https://assets-eu.researchsquare.com/files/rs-6301946/v1/5b4cee6c-e332-455f-bcd6-faa8113df14e.pdf"},{"id":80346947,"identity":"52ebaaa4-96b9-4573-ab57-d596bd0de4b9","added_by":"auto","created_at":"2025-04-10 20:18:22","extension":"mp4","order_by":0,"title":"","display":"","copyAsset":false,"role":"supplement","size":92109140,"visible":true,"origin":"","legend":"","description":"","filename":"MovieS1.mp4","url":"https://assets-eu.researchsquare.com/files/rs-6301946/v1/c92df62d2fedcdba59b2ddbe.mp4"},{"id":80347299,"identity":"3dc4b8aa-aa13-4f48-b2f1-5d3473348c30","added_by":"auto","created_at":"2025-04-10 20:26:17","extension":"docx","order_by":1,"title":"","display":"","copyAsset":false,"role":"supplement","size":6276100,"visible":true,"origin":"","legend":"","description":"","filename":"yonedaevodevoSI.docx","url":"https://assets-eu.researchsquare.com/files/rs-6301946/v1/67cbcad4fb2e2f6d5419406b.docx"}],"financialInterests":"No competing interests reported.","formattedTitle":"Lineage-specific head development in the coffin-headed cricket Loxoblemmus equestris links concentrated insect metamorphosis with novel trait evolution","fulltext":[{"header":"Background","content":"\u003cp\u003eLineage-specific diversification of homologous body parts creates evolutionary novelties from a constrained body plan shared within an animal phylum. Insecta is a taxon that shows prominent morphological novelties, such as pterygote thoracic wings, beetle head horns, and treehopper prothoracic helmets. It is suggested that these novelties originate from modifications of specific preexisting body parts, namely, the lateral tergum, the clypeolabrum, and the pronotum [1\u0026ndash;5], and exhibit remarkable diversifications compared with the corresponding body parts in closely related lineages. Evo-devo studies have revealed that co-option of ancestral gene regulatory networks is one of the driving forces that diversifies those novel traits (e.g., [6\u0026ndash;8]). However, how a novel trait originates from a preexisting structure in a constrained insect body plan remains unclear. To address this point, directly developing hemimetabolous insects are suitable models because dramatic remodeling of larval structures through the pupal stage obscures both tissue- and cellular-level relationships between juvenile and adult body parts in derived holometabolous insects (e.g., [9]).\u003c/p\u003e\n\u003cp\u003eOrthopteran species, including grasshoppers and crickets, serve as well-established genetic models of hemimetabolous development with expanding genomic resources and functional genomics tools such as RNAi and genome editing [10]. In orthopteran families, agonistic behaviors between males have facilitated the evolution of novel sexual dimorphisms, such as exaggerated hindlegs in Rhaphidoridae [11] and enlarged mandibles in Anostostomatidae [12], in a lineage-specific manner (Fig. 1A) [11\u0026ndash;17]. In Gryllidae, multiple genera exhibit sexual dimorphisms in adult heads, whereas other genera, including the popular laboratory model genera \u003cem\u003eGryllus\u003c/em\u003e and \u003cem\u003eAcheta\u003c/em\u003e, form monomorphic head shapes in both sexes (Fig. 1B). In the genus \u003cem\u003eLoxoblemmus\u003c/em\u003e, males specifically form a horn-like protrusion on the top of the head and a flattened frontal head region (Fig. 1C), whereas females retain the typical rounded head shape. In Gryllidae, the typical male fighting style involves fencing with antennae, flaring and grappling with mandibles, and a single headbutt or interlocking with mouthparts [15, 18\u0026ndash;21]. In addition to these widespread fighting styles, \u003cem\u003eLoxoblemmus\u003c/em\u003e species exhibit unique male‒male combat behavior by thrusting the male-specific flattened parts of their heads against each other (Movie S1). This fighting style is specifically described in \u003cem\u003eLoxoblemmus\u003c/em\u003e among Gryllidae [15]. The coffin-headed cricket \u003cem\u003eLoxoblemmus equestris\u003c/em\u003e, which is distributed on the southwest islands of Japan, Korea, China and Southeast Asia, is a suitable model species because of its ease of breeding and relatively short life cycle (3 months from egg to egg) under laboratory conditions.\u003c/p\u003e\n\u003cp\u003eTo understand the mechanisms underlying the origin of novel traits, we investigated regions modified to form the male-specific flattened and horned head in a lineage-specific manner and the temporal regulation behind this modification in \u003cem\u003eL. equestris\u003c/em\u003e.\u003c/p\u003e"},{"header":"Results","content":"\u003cp\u003e \u003cb\u003eA male-specific flattened head structure formed in the frons region during the last nymphal instar of\u003c/b\u003e \u003cb\u003eL. equestris\u003c/b\u003e\u003c/p\u003e \u003cp\u003eHemimetabolous insects exhibit two distinct developmental patterns of novel adult three-dimensional (3D) structures during postembryonic development. The first involves gradual formation across multiple nymphal instars. For example, species-specific pronotal shapes emerge in early nymphal stages and progressively develop into novel adult helmet structures over multiple nymphal instars in membracid species of treehoppers [\u003cspan citationid=\"CR5\" class=\"CitationRef\"\u003e5\u003c/span\u003e]. The second is concentrated formation during late nymphal development. For example, long rod-like nymphal cerci transform into short forceps during a single molt to reach adulthood in the basal earwig \u003cem\u003eDiplatys flavicollis\u003c/em\u003e [\u003cspan citationid=\"CR24\" class=\"CitationRef\"\u003e24\u003c/span\u003e].\u003c/p\u003e \u003cp\u003eTo determine which developmental pattern applies to the novel adult head structure, we observed nymphal head morphologies in \u003cem\u003eL. equestris\u003c/em\u003e. This species progresses through nine nymphal instars (N1 to N9) before reaching adulthood (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eA). Females can be distinguished from males with developing ovipositors at the eighth abdominal segment from the fourth nymphal instar under a stereomicroscope (Fig. \u003cspan refid=\"MOESM1\" class=\"InternalRef\"\u003eS1\u003c/span\u003e). Thus, we compared head morphology between males and females from the fourth nymphal instar to adulthood at each stage. We found that no remarkable sexual differentiations appeared in head structures during nymphal instars and that sexually dimorphic head structures abruptly formed in adults (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eA). This finding reveals that major head dimorphism occurs during the last nymphal instar in \u003cem\u003eL. equestris\u003c/em\u003e.\u003c/p\u003e \u003cp\u003eTo specify the flattened region in the adult male, we next annotated head landmarks according to Snodgrass (1935) [\u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e] and compared head morphologies between males and females in \u003cem\u003eL. equestris\u003c/em\u003e. We found that the lateral boundary of the flattened region strikingly corresponds to the subocular suture that \u0026ldquo;extends from the lower angle of each compound eye to the subgenal suture above the anterior mandibular articulation\u0026rdquo; in crickets [\u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e] (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eB). It thus revealed that the frons, which is defined as \u0026ldquo;the anterior region to the frontal suture, subocular suture and epistomal suture\u0026rdquo; [\u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e], is dramatically modified in males (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eC). In addition to the frons, we found notable sexual dimorphisms in the mandible and the clypeus (Fig. \u003cspan refid=\"MOESM2\" class=\"InternalRef\"\u003eS2\u003c/span\u003eA\u0026ndash;H). The sexually differential forms of the proximal mandibles at N9 suggest that the sex-specific development begins before the last nymphal instar (Fig. \u003cspan refid=\"MOESM2\" class=\"InternalRef\"\u003eS2\u003c/span\u003eA, B). In adults, the male mandible exhibits a wider proximal region with a darker inner edge, thereby forming a more triangular shape than the female mandible does (Fig. \u003cspan refid=\"MOESM2\" class=\"InternalRef\"\u003eS2\u003c/span\u003eC, D). Another adult dimorphism is the wider clypeus in males than in females (Fig. \u003cspan refid=\"MOESM2\" class=\"InternalRef\"\u003eS2\u003c/span\u003eG, H). In contrast, we detected no obvious differences in the labrum, maxilla, or labium between males and females (Fig. \u003cspan refid=\"MOESM2\" class=\"InternalRef\"\u003eS2\u003c/span\u003eI\u0026ndash;P). We focused on the frons dimorphism as an apparent lineage-specific novel trait for further analysis.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003e \u003cb\u003eThe male-specific frons structures are prepatterned as epithelial folding from the ninth day of the last nymphal instar\u003c/b\u003e \u003c/p\u003e \u003cp\u003eTo further understand when and how the dimorphic heads formed, we next observed epidermal tissue during the last nymphal instar (N9). We first conducted precise staging to determine the duration of the instar. Since hemimetabolous insects maintain constant feeding even in the preadult stage, we supplied an equivalent amount of food and water to nymphs individually maintained in an incubator during the same time window every day to ensure that the environmental conditions were as identical as possible among the monitored individuals (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eA). Under these conditions, the duration of the last nymphal instar was 299.86\u0026thinsp;\u0026plusmn;\u0026thinsp;3.89 (mean\u0026thinsp;\u0026plusmn;\u0026thinsp;standard deviation) hours (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eB). We next compared sectioned epidermal tissues along the midline, including the ocellus in the late last instar nymphs of both sexes (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eC). The nymphal cuticle and epidermis were attached 192 hours after ecdysis to the last instar nymph (HAE) (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eD, E). From 192 to 204 HAE, the nymphal cuticle started detaching from the epidermis, which indicates the onset of apolysis (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eF, G). By 216 HAE, the previously smooth epidermis had become corrugated (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eH, I). In this stage, males start to form sex-specific epithelial folds around the dorsal side of the median ocellus (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eH). By 240 HAE, deeper epithelial folds had been shaped, and the secretion of the adult cuticle had partially begun (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eJ, K). At this stage, we detected a sexually differential pattern in the epithelial folds around the ventral side of the median ocellus; males presented deeper folds than females did (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eJ, K). By 264 HAE, the secreted adult cuticle covered the entire epidermal surface, with the sexually differential folding pattern retained (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eL, M). In the adult stage, the ventral side of the ocellus became flatter, and the dorsal side protruded in males, whereas the nymphal shapes were mostly retained in corresponding regions of females (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eN, O). These observations revealed the formation of male-specific intensive epithelial folding after 204 HAE.\u003c/p\u003e \u003cp\u003eTo further investigate the sexually differential epithelial folds in three dimensions, we reconstructed the frons epidermis at 240 HAE by using micro-CT (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eA). Males formed more complex two-dimensional (2D) epithelial patterns than females did at both the dorsal and ventral sides of the median ocellus and around subocular sutures at this stage (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eB\u0026ndash;D, F\u0026ndash;H). In contrast, the posterior epidermis in the dorsal head was not obviously different between the sexes, suggesting that differential epithelial folding underlies the sexually dimorphic head shapes (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eE, I). To quantify differences in epithelial folding patterns between males and females, we measured the density and furrow depth in right, medial, and left sagittal sections at the ventral frons region (Fig. S3A). Both the density and the furrow depth significantly differed between males and females in the right and left sections (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eJ, K). In the medial section, the furrow depth significantly differed, although we found no significant difference in the furrow density between the sexes (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eJ, K). These analyses collectively specified the frons as the modified head region that gives rise to both flat and protruding shapes in \u003cem\u003eL. equestris\u003c/em\u003e.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cdiv id=\"Sec3\" class=\"Section2\"\u003e \u003ch2\u003eThe metamorphic gene network controls the timing of dimorphic head development\u003c/h2\u003e \u003cp\u003eThe metamorphic gene network (MGN), which comprises \u003cem\u003eKr\u0026uuml;ppel-homolog 1\u003c/em\u003e (\u003cem\u003eKr-h1\u003c/em\u003e), \u003cem\u003ebroad\u003c/em\u003e (\u003cem\u003ebr\u003c/em\u003e), and \u003cem\u003eEcdysone induced protein 93F\u003c/em\u003e (\u003cem\u003eE93\u003c/em\u003e), is proposed as an evolutionarily conserved central regulator of insect metamorphosis [\u003cspan citationid=\"CR26\" class=\"CitationRef\"\u003e26\u003c/span\u003e]. In \u003cem\u003eG. bimaculatus\u003c/em\u003e, \u003cem\u003eKr-h1\u003c/em\u003e and \u003cem\u003ebr\u003c/em\u003e play major roles in suppressing adult metamorphosis, whereas \u003cem\u003eE93\u003c/em\u003e promotes the nymph-to-adult transition [\u003cspan citationid=\"CR27\" class=\"CitationRef\"\u003e27\u003c/span\u003e]. Since the male-specific head structure formed during the preadult stage, we hypothesized that the MGN is involved in the temporal regulation of sexually dimorphic head formation in \u003cem\u003eL. equestris\u003c/em\u003e. To test this hypothesis, we first examined the temporal expression patterns of the three genes in the frons epithelial tissues of \u003cem\u003eL\u003c/em\u003e. \u003cem\u003eequestris\u003c/em\u003e from the penultimate nymphal stage to adulthood. The expression level of \u003cem\u003eE93\u003c/em\u003e was relatively low during the penultimate nymphal instar, gradually increased during the last nymphal instar, and reached its peak on the final day of the nymph (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eA). Conversely, \u003cem\u003eKr-h1\u003c/em\u003e was highly expressed during the penultimate instar, dramatically decreased in expression on the second day of the last nymphal instar, and maintained a low level thereafter (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eB). The \u003cem\u003ebr\u003c/em\u003e transcript presented no explicit temporal pattern during the late nymphal stage (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eC). These temporal gene expression patterns are similar to the reported patterns in the abdominal epidermis of \u003cem\u003eG. bimaculatus\u003c/em\u003e [\u003cspan citationid=\"CR27\" class=\"CitationRef\"\u003e27\u003c/span\u003e], which suggests a conserved function of the MGN in regulating metamorphosis between the two cricket species.\u003c/p\u003e \u003cp\u003eTo further understand the roles of these genes in male-specific head formation in \u003cem\u003eL. equestris\u003c/em\u003e, we next knocked down their transcript levels via nymphal RNAi. To implement the nymphal RNAi method in \u003cem\u003eL. equestris\u003c/em\u003e, we injected double-stranded (ds) RNA targeting \u003cem\u003eebony\u003c/em\u003e, which synthesizes N-β-alanyl-dopamine for light body coloration in \u003cem\u003eG\u003c/em\u003e. \u003cem\u003ebimaculatus\u003c/em\u003e [\u003cspan citationid=\"CR28\" class=\"CitationRef\"\u003e28\u003c/span\u003e], into penultimate instar nymphs. We successfully obtained dark-colored adults after \u003cem\u003eebony\u003c/em\u003e dsRNA injection, which validated the effect of RNAi-mediated gene knockdown in this species (Fig. S4, Table S3).\u003c/p\u003e \u003cp\u003eRNAi treatments targeting \u003cem\u003eE93\u003c/em\u003e, \u003cem\u003eKr-h1\u003c/em\u003e, or \u003cem\u003ebr\u003c/em\u003e caused temporal shifts in adult metamorphosis, whereas all the control individuals became adults following N9 (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eA\u0026ndash;K, A\u0026rsquo;\u0026ndash;G\u0026rsquo;, Table S4). All \u003cem\u003eL. equestris\u003c/em\u003e nymphs injected with \u003cem\u003eE93\u003c/em\u003e dsRNA at N8 failed to metamorphose into adults after N9. Among the surviving crickets, 38.9% and 44.4% underwent supernumerary nymphal molting and became larger adults after N10 and N11, respectively (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eC\u0026ndash;E, C\u0026rsquo;\u0026ndash;E\u0026rsquo;, I, Table S4). Notably, these supernumerary nymphs maintained the normal nymphal head proportion at N10 or N11, and the male-specific adult head shape first formed after metamorphic molts into adults (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eC\u0026ndash;E, C\u0026rsquo;\u0026ndash;E\u0026rsquo;). Compared with the control adults, the resulting larger adults presented overgrown wings (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eI). In contrast, RNAi treatments targeting \u003cem\u003eKr-h1\u003c/em\u003e or \u003cem\u003ebr\u003c/em\u003e at N4 caused precocious adult metamorphosis after N7 or N8 (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eF, G, F\u0026rsquo;, G\u0026rsquo;, J, K; Table S4), although the penetrance was relatively low (22.3% for \u003cem\u003eKr-h1\u003c/em\u003e and 17.4% for \u003cem\u003ebr\u003c/em\u003e). Despite being smaller in body size, the precocious adults presented male-specific adult head shapes, whereas their wings were disproportionate in shape (Fig. J, K). To analyze the effect of the shifted timing of adult metamorphosis on head proportion, we measured the ratio of head width to body length (Fig. S5A, B). Compared with those in the control group, we found no differences in the head-to-body ratio in adults that experienced supernumerary nymphal instars after \u003cem\u003eE93\u003c/em\u003e RNAi treatment, which indicates proportionate head formation (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eL). We note that incomplete molting or failure to unfold wings in most \u003cem\u003eE93\u003c/em\u003e RNAi individuals prohibited similar size measurements of the wings (Fig. S5C). These RNAi phenotypes confirmed the conserved functions of the MGN in \u003cem\u003eL. equestris\u003c/em\u003e and revealed that sex-specific novel head development is under the temporal control of the gene network.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003c/div\u003e"},{"header":"Discussion","content":"\u003cp\u003e \u003cb\u003eMale-specific novel head shapes are prepatterned as epithelial folding in\u003c/b\u003e \u003cb\u003eL. equestris\u003c/b\u003e\u003c/p\u003e \u003cp\u003eRecent studies have demonstrated that epithelial folding prepatterns sophisticated 3D structures in insects. For example, the protruding beetle horn is formed by the 3D expansion of patterned 2D epithelial furrows in the larval primordium [\u003cspan citationid=\"CR29\" class=\"CitationRef\"\u003e29\u003c/span\u003e]. Alterations in the furrow pattern, depth, and density affect the final shape of adult horns [\u003cspan citationid=\"CR30\" class=\"CitationRef\"\u003e30\u003c/span\u003e]. In \u003cem\u003eL\u003c/em\u003e. \u003cem\u003eequestris\u003c/em\u003e, males form denser, deeper furrows with more complex 2D patterns than females do in the frons epithelial cell sheets during the preadult stage (Figs.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003e, \u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003e). These data support the idea that sex-specific modifications in epithelial folding patterns led to the evolution of the novel head from an ancestral shape in Gryllidae.\u003c/p\u003e \u003cp\u003eExperimental and theoretical approaches have located the head region that undergoes differential growth to elaborate the protruding horn in the frons region of beetles [\u003cspan additionalcitationids=\"CR3\" citationid=\"CR2\" class=\"CitationRef\"\u003e2\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR4\" class=\"CitationRef\"\u003e4\u003c/span\u003e, \u003cspan citationid=\"CR31\" class=\"CitationRef\"\u003e31\u003c/span\u003e]. Differential forms of modified frons structures between the cricket and horned beetles reveal variation in modifications, ranging from flat surfaces to large protrusions, of a homologous region in insects. In both cases, modifications in frons shape are associated with male‒male combat behavior [\u003cspan citationid=\"CR15\" class=\"CitationRef\"\u003e15\u003c/span\u003e, \u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e32\u003c/span\u003e]. The difference in fighting styles could have driven diversification of differential forms from the homologous region or vice versa.\u003c/p\u003e \u003cp\u003e \u003cb\u003eSex-specific epithelial folding patterns develop through adult metamorphosis\u003c/b\u003e \u003cb\u003ein L. equestris\u003c/b\u003e\u003c/p\u003e \u003cp\u003eWe specified the timing of sexually dimorphic development of epithelial folding from 204 to 240 hours after ecdysis to the preadult instar stage in \u003cem\u003eL\u003c/em\u003e. \u003cem\u003eequestris\u003c/em\u003e (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003e). Critically, head sexual dimorphism occurs after adult metamorphosis despite the timing shifts caused by gene knockdown (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003e, Table S4). These data demonstrate that sex-specific patterns of epithelial folding are formed during adult metamorphosis, which is controlled by the MGN, and suggest that E93 is a major upstream regulator that determines the timing of dimorphic epithelial patterning. \u003cem\u003eE93\u003c/em\u003e and \u003cem\u003eKr-h1\u003c/em\u003e presented contrasting expression patterns in the frons tissues of \u003cem\u003eL\u003c/em\u003e. \u003cem\u003eequestris\u003c/em\u003e last instar nymphs (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003e). Targeted knockdown of \u003cem\u003eE93\u003c/em\u003e, \u003cem\u003eKr-h1\u003c/em\u003e and \u003cem\u003ebr\u003c/em\u003e caused delayed and precocious formations of head dimorphism, respectively (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003e). Antagonistic regulation between \u003cem\u003eKr-h1\u003c/em\u003e and \u003cem\u003eE93\u003c/em\u003e is widely conserved in both hemi- and holometabolous insects [\u003cspan citationid=\"CR26\" class=\"CitationRef\"\u003e26\u003c/span\u003e, \u003cspan citationid=\"CR33\" class=\"CitationRef\"\u003e33\u003c/span\u003e]. Indeed, knockdown of \u003cem\u003eKr-h1\u003c/em\u003e, and \u003cem\u003ebr\u003c/em\u003e, causes early upregulation of the \u003cem\u003eE93\u003c/em\u003e transcript level in another cricket species, \u003cem\u003eG\u003c/em\u003e. \u003cem\u003ebimaculatus\u003c/em\u003e [\u003cspan citationid=\"CR27\" class=\"CitationRef\"\u003e27\u003c/span\u003e]. In \u003cem\u003eDrosophila\u003c/em\u003e, the temporal expression of \u003cem\u003eE93\u003c/em\u003e broadly provides competence for target enhancers to respond to spatial cues through controlling chromatin accessibility [\u003cspan citationid=\"CR34\" class=\"CitationRef\"\u003e34\u003c/span\u003e]. Taken together, it is conceivable that E93 could bind enhancers regulating gene expression in the male frons during adult metamorphosis, thereby locally relaxing the body plan constraint to permit dramatic transformation of the preexisting ancestral juvenile shape into the derived adult form, which is likely adaptive for agonistic behavior.\u003c/p\u003e\n\u003ch3\u003eThe final molt as a driver of the evolution of novel traits in insects\u003c/h3\u003e\n\u003cp\u003eWhereas the male-specific head shape formed almost proportionately, the final size of the wings was highly disproportionate following timing shifts in metamorphosis caused by gene knockdowns, as reported in other hemimetabolous species (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003e; e.g., [\u003cspan citationid=\"CR27\" class=\"CitationRef\"\u003e27\u003c/span\u003e, \u003cspan citationid=\"CR35\" class=\"CitationRef\"\u003e35\u003c/span\u003e, \u003cspan citationid=\"CR36\" class=\"CitationRef\"\u003e36\u003c/span\u003e]). The contrasting phenotypes between the head and wings underscore the robustness of the head proportion against timing shifts in development. In hemimetabolous insects, wing primordia externally appear as wing pads and undergo anisometric growth during nymphal development [\u003cspan citationid=\"CR1\" class=\"CitationRef\"\u003e1\u003c/span\u003e, \u003cspan citationid=\"CR37\" class=\"CitationRef\"\u003e37\u003c/span\u003e]. Paleozoic fossils and recent evo-devo studies suggest that wings originated as lateral tergal outgrowths, such as those formed in palaeodictyopteran nymphs, modified during postembryonic development to form functional adult wings and their serial homologs [\u003cspan citationid=\"CR38\" class=\"CitationRef\"\u003e38\u003c/span\u003e]. It has been proposed that those wing precursors play preadaptive roles, such as thermoregulation, surface-skimming behavior, aerial body control, and respiration [\u003cspan citationid=\"CR38\" class=\"CitationRef\"\u003e38\u003c/span\u003e, \u003cspan citationid=\"CR39\" class=\"CitationRef\"\u003e39\u003c/span\u003e]. Precursors of the novel treehopper helmets on the prothorax externally appear in early nymphs [\u003cspan citationid=\"CR5\" class=\"CitationRef\"\u003e5\u003c/span\u003e], following a developmental pattern similar to that of wings. The shared developmental pattern suggests that gradual modifications during juvenile development serve as a general path to generate evolutionary novelty from the constrained insect body plan.\u003c/p\u003e \u003cp\u003eThe concentrated development of adult head dimorphism in \u003cem\u003eL. equestris\u003c/em\u003e demonstrates another path for the evolution of novelties from preexisting structures. The male-specific frons shape, which is used for male‒male combat behavior in adulthood, starts developing during a single preadult instar, thereby robustly forming a proportionate shape without specific early-forming precursors used in preadaptive roles. This highlights the evolution of the final molt and metamorphic life cycle in insects [\u003cspan citationid=\"CR40\" class=\"CitationRef\"\u003e40\u003c/span\u003e], which clearly splits the juvenile and adult stages by metamorphosis. The \u003cem\u003eE93\u003c/em\u003e expression level reflects the differential patterns of juvenile-to-adult transitions before and after the evolution of concentrated adult metamorphosis; the \u003cem\u003eE93\u003c/em\u003e level is mostly constant during the juvenile and adult stages in the ancestral ametabolous insect \u003cem\u003eThermobia domestica\u003c/em\u003e, whereas it exhibits a highly compressed peak during the late juvenile period in hemimetabolous insects [\u003cspan citationid=\"CR41\" class=\"CitationRef\"\u003e41\u003c/span\u003e]. Consistent with the expression pattern, it has been suggested that E93 acts on cuticle development throughout the nymphal stage in \u003cem\u003eT. domestica\u003c/em\u003e [\u003cspan citationid=\"CR42\" class=\"CitationRef\"\u003e42\u003c/span\u003e]. In contrast, concentrated \u003cem\u003eE93\u003c/em\u003e expression enables centralized regulation of the expression of a broad range of genes for terminal differentiation of adult structures during late juvenile development in derived hemi- and holometabolous insects [\u003cspan additionalcitationids=\"CR44\" citationid=\"CR43\" class=\"CitationRef\"\u003e43\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR45\" class=\"CitationRef\"\u003e45\u003c/span\u003e]. Our study reveals the direct link between the development of a lineage-specific 3D structure and the derived concentrated metamorphosis regulated by the MGN, suggesting the contribution of the evolution of a unique life cycle to the explosive morphological diversity of the hexapod lineage (Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003e).\u003c/p\u003e \u003cp\u003e \u003c/p\u003e"},{"header":"Conclusions","content":"\u003cp\u003eIn this study, using a cricket with a novel head, we found that the ancestral shape directly transforms into the novel shape through the final molt to adulthood, even after artificial timing shifts of metamorphosis, by creating sex-specific epithelial folding. This finding provides direct evidence that the unique metamorphic life cycle has facilitated the evolution of novelties in lineage- and sex-specific manners, thereby contributing to the morphological diversity of insects.\u003c/p\u003e"},{"header":"Methods","content":"\u003cdiv id=\"Sec8\" class=\"Section2\"\u003e \u003ch2\u003eCricket rearing and staging\u003c/h2\u003e \u003cp\u003eThe coffin-headed cricket \u003cem\u003eLoxoblemmus equestris\u003c/em\u003e (Saussure 1877) used in this study was derived from Iriomote Island, Okinawa, Japan. All crickets were maintained at 28\u0026ndash;30\u0026deg;C and 30% relative humidity under a 12 L:12 D photoperiod in an air-conditioned room and fed artificial goldfish food (Spectrum Brands). For precise staging, cricket nymphs were individually maintained in a plastic cup placed in an incubator set at 28\u0026deg;C under 12 L:12 D. We supplied five pellets of artificial goldfish food and water daily during 5:00\u0026ndash;8:00 PM. The water was supplied from a paper string to the maximum extent possible without it seeping out. Nymphs were isolated from the rearing group before the target stage. The timing of molting was monitored by taking time-lapse photographs every 3 minutes. The same staging method was used for cryosectioning and quantitative RT-PCR.\u003c/p\u003e \u003c/div\u003e\n\u003ch3\u003eScanning electron microscopy\u003c/h3\u003e\n\u003cp\u003eNonfixed head samples were mounted on sample stubs using carbon double-sided tape (NISSHIN EM, 7323). All images were captured at a voltage of 15 kV using a tabletop scanning electron microscope system (Miniscope\u0026reg; TM4000PlusII; HITACHI).\u003c/p\u003e\n\u003ch3\u003eEpithelial tissue histology\u003c/h3\u003e\n\u003cp\u003eFrozen sections were prepared according to the Kawamoto method [\u003cspan citationid=\"CR46\" class=\"CitationRef\"\u003e46\u003c/span\u003e]. Crickets were anesthetized on ice and dissected in phosphate-buffered saline (PBS). The whole-head samples were embedded in Super Cryoembedding Medium (SECTION-LAB, R060202). The embedded heads were sectioned at a thickness of 7 \u0026micro;m by using a cryostat (Leica CM3050 S; Leica Biosystems). Adhesive films (cryofilm type II C (9); SECTION-LAB, R050928) were used to capture cryosections. The sectioned tissues were fixed in 4% paraformaldehyde (PFA) for 5 minutes and then washed twice with PBS for 5 minutes each. The tissue samples were stained with Carrazzi\u0026rsquo;s hematoxylin solution (SECTION-LAB, R060213) for 5 minutes and rinsed with running tap water for 5 minutes to develop color. These samples were subsequently stained with 0.2% eosin Y solution (SECTION-LAB, R060214) for 30 seconds, followed by a 30 second rinse in running tap water. The stained sections were then washed with 100% ethanol to remove residual eosin and mounted in 50% glycerol.\u003c/p\u003e \u003cdiv id=\"Sec11\" class=\"Section2\"\u003e \u003ch2\u003eX-ray micro-computed tomography (CT) analysis\u003c/h2\u003e \u003cp\u003eFixation, washing, and staining steps were performed on a rotary platform shaker at room temperature. Last-instar nymphs of \u003cem\u003eL. equestris\u003c/em\u003e at 240 hours after molting were anesthetized on ice for head dissection. The head samples were fixed in 4% PFA overnight and then washed three times in PBS for 10 minutes each. Fixed samples were stained with 25% Lugol solution for 48 hours, followed by washing twice with PBS overnight. The stained samples were embedded in paraffin and scanned using an X-ray micro-CT device (SKYSCAN 1272 CMOS Edition; BRUKER) at a voltage peak of 60 kV and a current of 80 \u0026micro;A. The samples were rotated 360 degrees in steps of 0.08 degrees, generating 4500 projection images for each of the male and female samples. The micro-CT data were reconstructed at isotropic resolutions of 1.5 \u0026micro;m and 1.35 \u0026micro;m for the male and female samples, respectively, via NRecon software (v2.0.0.5, Micro Photonics Inc.). Three-dimensional (3D) tomographic images were obtained and processed using VGSTUDIO MAX software (Volume Graphics).\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec12\" class=\"Section2\"\u003e \u003ch2\u003eQuantitative analysis of epidermal folding patterns\u003c/h2\u003e \u003cp\u003eThe number of furrows and depth of epidermal folding were measured according to Adachi (2020) [\u003cspan citationid=\"CR30\" class=\"CitationRef\"\u003e30\u003c/span\u003e]. The cross-sectional images used to quantify furrow depths were captured as CT images using VGSTUDIO MAX software. Brightness adjustments and depth measurements of the furrows were performed using ImageJ. The \u0026ldquo;SegmentMeasure\u0026rdquo; plug-in of ImageJ was used to measure the length of the epidermis.\u003c/p\u003e \u003cp\u003e \u003cb\u003eDe novo\u003c/b\u003e \u003cb\u003etranscriptome assembly\u003c/b\u003e\u003c/p\u003e \u003cp\u003eTo obtain the transcript sequences of \u003cem\u003eL. equestris\u003c/em\u003e, we assembled short RNA-seq reads. We pooled total RNA extracted from whole embryos at 3, 7, and 12 days after egg laying and whole heads of individual male and female last-instar nymphs at 2, 5, 10, and 11 days after molting. Total RNA extraction was performed using TRIzol reagent (Thermo Fischer Scientific, 15596018), followed by phenol‒chloroform extraction. Subsequent purification was conducted using RNeasy columns (RNeasy\u0026reg; Micro Kit (50); QIAGEN, 172045635). The purified total RNA was then submitted to Azenta Life Sciences for library preparation using NEBNext Ultra II Directional RNA Library Prep Kit for Illumina. Sequencing was carried out on an Illumina NovaSeq platform, generating 150 bp paired-end reads. Poor-quality reads and adapter sequences were discarded with Cutadapt (v4.4) with default parameter settings. Trinity (v2.13.2) was used for de novo transcriptome assembly.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec13\" class=\"Section2\"\u003e \u003ch2\u003eBLAST search for orthologous genes\u003c/h2\u003e \u003cp\u003eWe searched for orthologs of \u003cem\u003eKr\u0026uuml;ppel-homolog 1\u003c/em\u003e (\u003cem\u003eKr-h1\u003c/em\u003e), \u003cem\u003ebroad\u003c/em\u003e (\u003cem\u003ebr\u003c/em\u003e), and \u003cem\u003eEcdysone induced protein 93F\u003c/em\u003e (\u003cem\u003eE93\u003c/em\u003e) in the \u003cem\u003eL. equestris\u003c/em\u003e transcriptome using public partial cDNA sequences from \u003cem\u003eG. bimaculatus\u003c/em\u003e (\u003cem\u003eE93\u003c/em\u003e, LC476893; \u003cem\u003eKr-h1\u003c/em\u003e, LC476894; \u003cem\u003ebr\u003c/em\u003e, LC476892) as query sequences for the BLASTn program. We confirmed that the identified transcript sequences of \u003cem\u003eL. equestris\u003c/em\u003e presented the best hits to the corresponding genes of \u003cem\u003eD. melanogaster\u003c/em\u003e [\u003cspan citationid=\"CR47\" class=\"CitationRef\"\u003e47\u003c/span\u003e] via BLASTx searches.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec14\" class=\"Section2\"\u003e \u003ch2\u003eQuantitative RT-PCR (qRT-PCR)\u003c/h2\u003e \u003cp\u003eTotal RNA was extracted from frons epidermal tissues, including the cuticles of penultimate and last-instar male nymphs. The tissues were dissected in PBS after the crickets were anesthetized on ice. All tissue samples were collected in TRIzol reagent and stored at -80\u0026deg;C until use. Total RNA was extracted according to the manufacturer\u0026rsquo;s instructions. The extracted RNA was reverse transcribed to cDNA using ReverTra Ace\u0026reg; qPCR RT Master Mix with gDNA Remover (TOYOBO, FSQ-301) according to the manufacturer\u0026rsquo;s instructions. qRT-PCR was performed using THUNDERBIRD SYBR qPCR mix (TOYOBO, QPS-201) on QuantStudio\u0026trade; 3 Real-Time PCR System (Applied Biosystems). The primers used for qRT-PCR are listed in Table \u003cspan refid=\"MOESM1\" class=\"InternalRef\"\u003eS1\u003c/span\u003e. The qRT-PCR program was as follows: 95\u0026deg;C for 5 min, 95\u0026deg;C for 30 s, and 60\u0026deg;C for 45 s for 40 cycles with each primer at a concentration of 10 \u0026micro;M. The \u003cem\u003eRibosomal protein L32\u003c/em\u003e (\u003cem\u003eRpL32\u003c/em\u003e) gene was used as a reference gene for qRT-PCR analysis. All qRT-PCRs were performed in triplicate as technical replicates. The relative quantity of transcripts was calculated by the 2-ΔΔCt method (Livak \u0026amp; Schmittggen, 2001).\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec15\" class=\"Section2\"\u003e \u003ch2\u003eRNA interference (RNAi)-mediated gene knockdown\u003c/h2\u003e \u003cp\u003eDNA templates for double-stranded RNA (dsRNA) synthesis were amplified from late-stage embryo cDNA using primers with the T7 promoter sequence at the 5\u0026prime; end listed in Table \u003cspan refid=\"MOESM2\" class=\"InternalRef\"\u003eS2\u003c/span\u003e. To obtain a sufficient amount of DNA template for \u003cem\u003ein vitro\u003c/em\u003e transcription, we performed PCR twice; first, AmpliTaq Gold\u0026trade; 360 DNA Polymerase (Applied Biosystems, 4398823) was used, and then ExTaq (Takara, RR001B) was used for all the targets except for \u003cem\u003eegfp\u003c/em\u003e. The first PCR product is used as a template for the second PCR. For \u003cem\u003eegfp\u003c/em\u003e, the DNA template was amplified from plasmid DNA using ExTaq. RNA was transcribed from a DNA template using the T7 RiboMAX Express Large Scale RNA System (Promega, P1320) and purified using phenol/chloroform/isoamyl alcohol (125:24:1) (Merck, 77619-100ML). The transcribed RNA was resuspended in annealing buffer [10 mM Tris-Cl (pH 7.5), 1 mM EDTA, and 5 mM NaCl/DEPC-treated H\u003csub\u003e2\u003c/sub\u003eO] and annealed by gradual cooling from 95\u0026deg;C to 25\u0026deg;C on a programmed thermal cycler to make dsRNA. Annealed dsRNA solutions were aliquoted and stored at -80\u0026deg;C. DsRNA solution was injected into the lateral ventral side of the intersegmental membrane between nymphal abdominal segments within 24 hours postecdysis. For \u003cem\u003eE93\u003c/em\u003e and \u003cem\u003eKr-h1\u003c/em\u003e and \u003cem\u003ebr\u003c/em\u003e knockdown, 12 \u0026micro;g and 6 \u0026micro;g of dsRNA solution, respectively, were injected into penultimate and fourth instar nymphs of males. For all the RNAi experiments, an equivalent amount of \u003cem\u003eegfp\u003c/em\u003e dsRNA was injected as a negative control.\u003c/p\u003e \u003c/div\u003e"},{"header":"Declarations","content":"\u003cp\u003e\u003cstrong\u003eAcknowledgments\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eWe thank Mr. Hosei Wada for developing the \u0026ldquo;SegmentMeasure\u0026rdquo; plug-in of ImageJ, Ms. Junko Morita for her technical support, and Drs. Taro Nakamura, Toshiya Ando and Naoki Matsuda for their helpful discussion.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAuthor contributions\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eMY: all research performance, manuscript preparation. SM: X-ray micro-CT analysis, review the manuscript. TN: X-ray micro-CT analysis, review the manuscript. TD: review the manuscript. TO: research design, review the manuscript.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eFunding\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThis work was supported by the Japan Society for the Promotion of Science KAKENHI [grant numbers 23K18026 (TO), 24H00511 (TD)] and the NIBB Collaborative Research Program [grant numbers 22NIBB336, 23NIBB343, 24NIBB326 (TO)].\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAvailability of data and materials\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eAll data are included in the main manuscript and Additional file 1.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eEthics approval and consent to participate\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eNot applicable.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eConsent for publication\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eNot applicable.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eCompeting interests\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe authors declare no competing interests.\u003c/p\u003e"},{"header":"References","content":"\u003col\u003e\n\u003cli\u003eOhde T, Mito T, Niimi T. 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Complex furrows in a 2D epithelial sheet code the 3D structure of a beetle horn. Sci Rep. 2017;7(1):13939.\u003c/li\u003e\n\u003cli\u003eAdachi H, Matsuda K, Niimi T, et al. Genetical control of 2D pattern and depth of the primordial furrow that prefigures 3D shape of the rhinoceros beetle horn. Sci Rep. 2020;10(1):18687.\u003c/li\u003e\n\u003cli\u003eMorikawa K, Morita S, Sakura K, et al. Unveiling the role of differential growth in 3D morphogenesis: An inference method to analyze area expansion rate distribution in biological systems. J Theor Biol. 2023;575:111650.\u003c/li\u003e\n\u003cli\u003eMcCullough EL, Tobalske BW, Emlen DJ. Structural adaptations to diverse fighting styles in sexually selected weapons. Proc Natl Acad Sci U S A. 2014;111(40):14484-8.\u003c/li\u003e\n\u003cli\u003eUre\u0026ntilde;a E, Chafino S, Manj\u0026oacute;n C, et al. The Occurrence of the Holometabolous Pupal Stage Requires the Interaction between E93, Kr\u0026uuml;ppel-Homolog 1 and Broad-Complex. PLoS Genet. 2016;12(5):e1006020.\u003c/li\u003e\n\u003cli\u003eNystrom SL, Niederhuber MJ, McKay DJ. Expression of E93 provides an instructive cue to control dynamic enhancer activity and chromatin accessibility during development. Development. 2020;147(6):dev181909.\u003c/li\u003e\n\u003cli\u003eKonopova B, Smykal V, Jindra M. Common and Distinct Roles of Juvenile Hormone Signaling Genes in Metamorphosis of Holometabolous and Hemimetabolous Insects. PLoS ONE. 2011;6(12):e28728.\u003c/li\u003e\n\u003cli\u003eLozano J, Belles X. Conserved repressive function of Kr\u0026uuml;ppel homolog 1 on insect metamorphosis in hemimetabolous and holometabolous species. Sci Rep. 2011;1(1):163.\u003c/li\u003e\n\u003cli\u003eErezyilmaz DF, Riddiford LM, Truman JW. The pupal specifier broad directs progressive morphogenesis in a direct-developing insect. Proc Natl Acad Sci U S A. 2006;103(18):6925-30.\u003c/li\u003e\n\u003cli\u003eOhde T, Prokop J. The transition to flying insects: lessons from evo-devo and fossils. Curr Opin Insect Sci. 2025;68:101332.\u003c/li\u003e\n\u003cli\u003eTreidel LA, Deem KD, Salcedo MK, et al. Insect Flight: State of the Field and Future Directions. Integr Comp Biol. 2024;64(2):533-55.\u003c/li\u003e\n\u003cli\u003eBelles X. The innovation of the final moult and the origin of insect metamorphosis. Phil Trans R Soc B. 2019;374(1783):20180415.\u003c/li\u003e\n\u003cli\u003eFernandez-Nicolas A, Machaj G, Ventos-Alfonso A, et al. Reduction of embryonic E93 expression as a hypothetical driver of the evolution of insect metamorphosis. Proc Natl Acad Sci U S A. 2023;120(7):e2216640120.\u003c/li\u003e\n\u003cli\u003eBai Y, Lv YN, Zeng M, et al. E93 is indispensable for reproduction in ametabolous and hemimetabolous insects. Development. 2024;151(20):dev202518.\u003c/li\u003e\n\u003cli\u003eZeng M, Yan ZY, Lv YN, et al. Molecular basis of E93-dependent tissue morphogenesis and histolysis during insect metamorphosis. Insect Biochem Mol Biol. 2025;177:104249.\u003c/li\u003e\n\u003cli\u003eUyehara CM, Nystrom SL, Niederhuber MJ, et al. Hormone-dependent control of developmental timing through regulation of chromatin accessibility. Genes Dev. 2017;31(9):862-75.\u003c/li\u003e\n\u003cli\u003eCruz J, Ure\u0026ntilde;a E, I\u0026ntilde;iguez LP, et al. E93 controls adult differentiation by repressing broad in Drosophila. Proc Natl Acad Sci U S A. 2024;121(51):e2403162121.\u003c/li\u003e\n\u003cli\u003eKawamoto T, Kawamoto K. Preparation of Thin Frozen Sections from Nonfixed and Undecalcified Hard Tissues Using Kawamoto\u0026rsquo;s Film Method. In: Hilton MJ, editor. Skeletal Development and Repair. Totowa, NJ: Humana Press; 2014. p. 149-64. (Methods in Molecular Biology; vol. 1130). Available from: https://link.springer.com/10.1007/978-1-62703-989-5_11\u003c/li\u003e\n\u003cli\u003eHoskins RA, Carlson JW, Wan KH, et al. The Release 6 reference sequence of the Drosophila melanogaster genome. Genome Res. 2015;25(3):445-58.\u003c/li\u003e\n\u003c/ol\u003e"}],"fulltextSource":"","fullText":"","funders":[],"hasAdminPriorityOnWorkflow":false,"hasManuscriptDocX":true,"hasOptedInToPreprint":true,"hasPassedJournalQc":"","hasAnyPriority":false,"hideJournal":false,"highlight":"","institution":"","isAcceptedByJournal":true,"isAuthorSuppliedPdf":false,"isDeskRejected":"","isHiddenFromSearch":false,"isInQc":false,"isInWorkflow":false,"isPdf":false,"isPdfUpToDate":true,"isWithdrawnOrRetracted":false,"journal":{"display":true,"email":"[email protected]","identity":"developmental-biology-advances","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":false,"externalIdentity":"evod","sideBox":"Learn more about [EvoDevo](http://evodevojournal.biomedcentral.com/)","snPcode":"13227","submissionUrl":"https://submission.nature.com/new-submission/13227/3","title":"Developmental Biology Advances","twitterHandle":"@BioMedCentral","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"stoa","reportingPortfolio":"BMC/SO AJ","inReviewEnabled":true,"inReviewRevisionsEnabled":true},"keywords":"","lastPublishedDoi":"10.21203/rs.3.rs-6301946/v1","lastPublishedDoiUrl":"https://doi.org/10.21203/rs.3.rs-6301946/v1","license":{"name":"CC BY 4.0","url":"https://creativecommons.org/licenses/by/4.0/"},"manuscriptAbstract":"\u003ch2\u003eBackground\u003c/h2\u003e \u003cp\u003eLineage-specific adult structures form through modifications of preexisting juvenile body parts during postembryonic development in insects. It remains unclear how these novel traits originate from ancestral structures within the constrained body plan. In the coffin-headed cricket \u003cem\u003eLoxoblemmus equestris\u003c/em\u003e, an ancestral rounded head shape directly transforms into a flattened, derived form in a sex-specific manner. To understand the origin of novel traits, we investigated the development of the adult head in \u003cem\u003eL\u003c/em\u003e. \u003cem\u003eequestris\u003c/em\u003e as a model of lineage-specific novelty.\u003c/p\u003e\u003ch2\u003eResults\u003c/h2\u003e \u003cp\u003eDetailed two- and three-dimensional analyses of the developing head revealed that sexually dimorphic epithelial patterns formed in a specific region, the frons, during the preadult instar. The male-specific head shapes are formed following the final molt to adulthood even after timing shifts of the metamorphosis induced by RNA interference targeting the evolutionarily conserved metamorphic gene network.\u003c/p\u003e\u003ch2\u003eConclusions\u003c/h2\u003e \u003cp\u003eThese findings demonstrate that adult metamorphosis, led by E93, locally relaxes the body plan constraint to permit dramatic transformation of juvenile body parts into a novel head shape by modifying epithelial folding in \u003cem\u003eL\u003c/em\u003e. \u003cem\u003eequestris\u003c/em\u003e. This highlights concentrated metamorphosis through the final molt as a driver that creates lineage- and sex-specific adult forms in the hexapod lineage.\u003c/p\u003e","manuscriptTitle":"Lineage-specific head development in the coffin-headed cricket Loxoblemmus equestris links concentrated insect metamorphosis with novel trait evolution","msid":"","msnumber":"","nonDraftVersions":[{"code":1,"date":"2025-04-10 20:18:12","doi":"10.21203/rs.3.rs-6301946/v1","editorialEvents":[{"type":"communityComments","content":1},{"type":"decision","content":"Revision requested","date":"2025-06-02T07:09:12+00:00","index":"","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2025-05-25T20:52:13+00:00","index":"hide","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2025-05-06T14:09:25+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"229359230083512715796309285204610082099","date":"2025-04-12T18:40:49+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"272924609984989365543506471533194921102","date":"2025-04-09T14:20:59+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"96368126644547898068225036113727874994","date":"2025-03-31T07:10:48+00:00","index":"hide","fulltext":""},{"type":"reviewersInvited","content":"","date":"2025-03-26T12:32:20+00:00","index":"","fulltext":""},{"type":"editorAssigned","content":"","date":"2025-03-26T12:13:08+00:00","index":"","fulltext":""},{"type":"checksComplete","content":"","date":"2025-03-26T12:12:05+00:00","index":"","fulltext":""},{"type":"submitted","content":"EvoDevo","date":"2025-03-25T08:41:12+00:00","index":"","fulltext":""}],"status":"published","journal":{"display":true,"email":"[email protected]","identity":"developmental-biology-advances","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":false,"externalIdentity":"evod","sideBox":"Learn more about [EvoDevo](http://evodevojournal.biomedcentral.com/)","snPcode":"13227","submissionUrl":"https://submission.nature.com/new-submission/13227/3","title":"Developmental Biology Advances","twitterHandle":"@BioMedCentral","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"stoa","reportingPortfolio":"BMC/SO AJ","inReviewEnabled":true,"inReviewRevisionsEnabled":true}}],"origin":"","ownerIdentity":"6c8f92f1-68ea-4a67-bdd4-8a742b7c1ca8","owner":[],"postedDate":"April 10th, 2025","published":true,"recentEditorialEvents":[],"rejectedJournal":[],"revision":"","amendment":"","status":"published-in-journal","subjectAreas":[],"tags":[],"updatedAt":"2025-07-21T16:11:40+00:00","versionOfRecord":{"articleIdentity":"rs-6301946","link":"https://doi.org/10.1186/s13227-025-00249-3","journal":{"identity":"developmental-biology-advances","isVorOnly":false,"title":"Developmental Biology Advances"},"publishedOn":"2025-07-16 16:05:34","publishedOnDateReadable":"July 16th, 2025"},"versionCreatedAt":"2025-04-10 20:18:12","video":"","vorDoi":"10.1186/s13227-025-00249-3","vorDoiUrl":"https://doi.org/10.1186/s13227-025-00249-3","workflowStages":[]},"version":"v1","identity":"rs-6301946","journalConfig":"researchsquare"},"__N_SSP":true},"page":"/article/[identity]/[[...version]]","query":{"redirect":"/article/rs-6301946","identity":"rs-6301946","version":["v1"]},"buildId":"8U1c8b4HqxoKbykW_rLl7","isFallback":false,"isExperimentalCompile":false,"dynamicIds":[84888],"gssp":true,"scriptLoader":[]}