Skip to main content
NCBI home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2023 Jun 1.
Published in final edited form as: Dev Biol. 2023 Mar 24;498:87–96. doi: 10.1016/j.ydbio.2023.03.008

Segmental expression of two ecdysone pathway genes during embryogenesis of hemimetabolous insects

Judith Wexler a,b,*, Leslie Pick b, Ariel Chipman a
PMCID: PMC10228571  NIHMSID: NIHMS1894900  PMID: 36967076

Abstract

Signaling networks are redeployed across different developmental times and places to generate phenotypic diversity from a limited genetic toolkit. Hormone signaling networks in particular have well-studied roles in multiple developmental processes. In insects, the ecdysone pathway controls critical events in late embryogenesis and throughout post-embryonic development. While this pathway has not been shown to function in the earliest stage of embryonic development in the model insect Drosophila melanogaster, one component of the network, the nuclear receptor E75A, is necessary for proper segment generation in the milkweed bug Oncopeltus fasciatus. Published expression data from several other species suggests possible conservation of this role across hundreds of millions of years of insect evolution. Previous work also demonstrates a second nuclear receptor in the ecdysone pathway, Ftz-F1, plays a role in segmentation in multiple insect species. Here we report tightly linked expression patterns of ftz-F1 and E75A in two hemimetabolous insect species, the German cockroach Blattella germanica and the two-spotted cricket Gryllus bimaculatus. In both species, the genes are expressed segmentally in adjacent cells, but they are never co-expressed. Using parental RNAi, we show the two genes have distinct roles in early embryogenesis. E75A appears necessary for abdominal segmentation in B. germanica, while ftz-F1 is essential for proper germband formation. Our results suggest that the ecdysone network is critical for early embryogenesis in hemimetabolous insects.

1. Introduction

Ecdysone was one of the first hormones discovered to control gene expression. Early work conducted on dipteran salivary glands from Chironomus tentans (Beermann, 1952) and Drosophila melanogaster (Becker, 1962) showed that application of ecdysone results in a stereotypic series of chromosomal puffs specific in both place and time. These puffs are regions of active gene transcription (Clever, 1964).

In holometabolous insects, two ecdysone pulses occur in the time leading to and then again several hours after pupariation (Riddiford, 1993). Puffs observed after the first ecdysone pulse are directly controlled by the hormone and are called “early puffs.” The much more numerous puffs occurring after the second pulse are under the control of both ecdysone and the gene products of the early puffs; they are known as “late puffs” (Ashburner et al., 1974). The transcripts resulting from the ecdysone-induced puffs encode gene products critical for successful metamorphosis.

E75 is a nuclear hormone receptor gene expressed in response to both the early and late ecdysone pulses. It is comprised of a DNA binding domain upstream of a ligand binding domain (King-Jones and Thummel, 2005; Erezyilmaz et al., 2009a,b). In D. melanogaster, there are three isoforms (E75A, E75B, and E75C) transcribed from the E75 locus (Bialecki et al., 2002). Mutants for each isoform have distinct physiological phenotypes. A fraction of E75A mutants die when failing to molt properly through larval instars; of those that survive to pupariation, a fraction fail to eclose to adults (Bialecki et al., 2002). The first pulse of E75A transcription is also necessary to set the stage for the organism’s response to the second ecdysone pulse. E75A is necessary for the transcription of ftz-F1, which encodes a nuclear hormone receptor expressed in between ecdysone pulses. Like with E75A, Ftz-F1 has a DNA binding domain upstream of a ligand binding domain (King-Jones and Thummel, 2005; Ohno and Petkovich, 1993). Ftz-F1 confers competence to cells so they can respond properly to the second ecdysone pulse (Woodard et al., 1994; Broadus et al., 1999), and in fact is necessary for transcription of E75A during the second ecdysone pulse (Broadus et al., 1999). Ectopic ftz-F1 expression leads to enhanced E75 transcription (Woodard et al., 1994). Thus, E75A and ftz-F1 both regulate each other – the first pulse of E75A turns on ftz-F1, which in turn amplifies E75A transcription in response to the second ecdysone pulse. ftz-F1 zygotic mutants cannot properly evert the head at the prepupal-pupal transition, and they also fail to elongate limbs properly (Broadus et al., 1999).

Unlike holometabolous insects, hemimetabolous insects lack a pupal stage. Nevertheless, the terminal molt from the last nymphal instar to the adult is qualitatively different from earlier molts in the life cycle. It is only during this last molt that wings appear in hemimetabolous insects and that they become sexually mature. The ecdysone network plays a critical role in coordinating the events of the terminal molt in hemimetabolous insects, just as it does in controlling the process of metamorphosis in holometabolous insects.

Much of our knowledge about the role of the ecdysone pathway in post-embryonic hemimetabolous insect development comes from the German cockroach Blattella germanica. Ecdysone spikes in B. germanica nymphs shortly before their terminal molt. This surge is responsible for the activation of ftz-F1, which in turn controls the programmed cell death of a nymphal specific gland in the transition to adulthood (Mané-Padrós et al., 2010). When E75A is knocked down in B. germanica, animals remain in their last nymphal stage and never progress to the terminal molt to adulthood (Mané-Padrós et al., 2008). Similarly, when ftz-F1 is knocked down in B. germanica last instar nymphs, the majority of animals either cannot molt to adulthood and die, or they molt without properly shedding their nymphal exuvia (Cruz et al., 2008).

In addition to the key roles played by ftz-F1 and E75A in post-embryonic development, each gene has a documented role during embryonic development in the segmentation of holo- and hemimetabolous insects, respectively. ftz-F1 acts maternally as a pair-rule gene in D. melanogaster, with mutants missing every other segment (Yu et al., 1997; Guichet et al., 1997). In D. melanogaster, Ftz-F1 is an obligate partner of the pair-rule protein Ftz, directly co-regulating the expression of the segment polarity gene engrailed (Florence et al., 1997) and other genes expressed in striped patterns in early embryos (Hou et al., 2009; Field et al., 2016). In Tribolium castaneum, ftz-F1 knockdown also produces embryos with a classic pair rule phenotype (Heffer et al., 2013). In the hide beetle Dermestes maculatus, ftz-F1 is expressed in stripes during early embryogenesis, but its role seems relegated to the processes of appendage formation, which occur after segmentation (Xiang et al., 2017). In the hemimetabolous milkweed bug Oncopeltus fasciatus, ftz-F1 is expressed in two stripes in the posterior part of the germband during appendage formation, although functional data for the gene is not available in this species (Reding et al., 2019). E75A was unexpectedly found to have pair-rule gene expression and function in O. fasciatus. Of-E75A is expressed in every other segment in the blastoderm, and depletion of the gene results in segmental fusion (Erezyilmaz et al., 2009a,b). In another hemipteran species, Murgantia histrionica, and in the hymenopteran Nasonia vitripennis, E75A is also expressed in a pair-rule pattern during early embryogenesis (Hernandez et al., 2020; Taylor and Dearden, 2022).

In B. germanica, ecdysone pulses occur both in early and in late embryogenesis (Maestro et al., 2005; Piulachs et al., 2010). Recent work by Cruz et al. demonstrated that ecdysone signaling is necessary for germband formation in B. germanica (Cruz et al., 2022; Fernandez-Nicolas et al., 2023). This research complements earlier work (Piulachs et al., 2010) showing that Broad-Complex, a gene turned on in the ecdysone response pathway, is necessary for germband development. In O. fasciatus, broad is expressed in two distinct pulses during early and late embryonic development respectively and is necessary for proper embryogenesis (Erezyilmaz et al., 2009a,b).

Given the roles that ftz-F1 and E75A play in segmentation of insect embryos, and given the demonstrated roles of ecdysone in the germband formation of B. germanica, we studied these genes during embryogenesis in two hemimetabolous insect orders – Blattodea and Orthoptera. Within Blattodea, we described expression patterns and functions of ftz-F1 and E75A in B. germanica early embryogenesis. In Orthoptera, we examined expression patterns of the two genes in the two-spotted cricket Gryllus bimaculatus. Using hybridization chain reaction (HCR), we found similar patterns of ftz-F1 and E75A expression during abdominal segmentation of the two species. At this stage, ftz-F1 and E75A are expressed in stripes in mutually exclusive cell populations. Surprisingly, during gnathal and thoracic segmentation, we found expression in a pair-rule register for ftz-F1 in B. germanica, and for both ftz-F1 and E75A in G. bimaculatus. Using parental RNAi in B. germanica, we found that each gene plays distinct roles in the early development of this species’ embryos.

2. Results and discussion

2.1. Gene isolation

E75A and ftz-F1 have already been studied in B. germanica (Mané-Padrós et al., 2008, 2010; Borras-Castells et al., 2017). To identify the genes in G. bimaculatus, we used degenerate PCR and 5’ RACE to amplify transcripts of both genes from G. bimaculatus. We discovered three E75 isoforms which differed in their 5′ ends (Lu et al., 2020) (Supplemental Fig. 1), as do E75 isoforms from B. germanica and O. fasciatus (Mané-Padrós et al., 2008; Erezyilmaz et al., 2009a,b). Only one of these isoforms contains both a predicted DNA-binding domain and a ligand-binding domain. We refer to this isoform as Gb-E75A. The other two isoforms differ in their predicted translation start codons, both of which are downstream of the predicted start codon of Gb-E75A. 5’ RACE uncovered three separate isoforms of ftz-F1 expressed in G. bimaculatus embryos. Only one of these isoforms, which we called Isoform B, encodes both an intact DNA-binding domain and ligand-binding domain typical of Ftz-F1 protein (Supplemental Fig. 2).

2.2. Bg-ftz-F1 and Bg-E75A are never co-expressed in B. germanica embryos

The B. germanica germband condenses as two plates of cells migrate towards the ventral side of egg and then fuse from posterior to anterior (Supplemental Fig. 3). In this paper, we consider the germband stage to start immediately after fusion of the head lobes. Prior to this fusion event, we refer to germband condensation. In insects, the genes engrailed, wingless and hedgehog are expressed in every segment of the embryo, and their expression can be used as a marker for assessing correct segment formation (Marie and Bacon, 2000; Davis and Patel, 2002; Peel et al., 2005). engrailed expression during the condensation of the B. germanica germband shows gnathal and thoracic segments forming at this time (Supplemental Fig. 4). After the germband condenses, abdominal segmentation can be observed via appearance of an increasing number of engrailed stripes (Supplemental Fig. 4, panels C, D, E).

During germband condensation, Bg-ftz-F1 expression was detected in three broadly spaced stripes along the posterior two thirds of the condensing germband (Fig. 1A and B). Despite numerous colorimetric and fluorescent in-situs, a similar striped Bg-E75A expression pattern was never observed in early embryos. Instead, Bg-E75A was only detected in the posterior of the germ rudiment during this stage (Fig. 1C). Along with the two anterior most broad stripes (asterisks, Fig. 1D, D’) of Bg-ftz-F1, a single band of Bg-E75A expression (green) is present in the posterior region of the developing germband (Fig. 1D, D”). This stripe of Bg-E75A expression is flanked by two stripes of Bg-ftz-F1 (magenta) expression. Limited data show that Bg-ftz-F1 is expressed in proximity to Bg-hh stripes at some point during gnathal and thoracic segmentation, but more investigation is needed to understand the relationship between Bg-E75A, Bg-ftz-F1, and segment polarity genes in this species (Supplemental Fig. 5).

Fig. 1.

Fig. 1.

Open in a new tab

Expression of Bg-ftz-F1 and Bg-E75A in B. germanica embryos. (A, B, C) Colorimetric in-situ of indicated gene in embryos still attached to yolk, before germbands have completely formed. Embryos shown from lateral view. (A) Expression of Bg-ftz-F1 appears as three broadly spaced regions in the condensing germband. (B) Expression of Bg-ftz-F1 in a slightly older embryo, with stripes of expression which do not extend laterally as far as in panel (A). (C) Expression of Bg-E75A at the same stage appears only as a diffuse smear in the posterior region of the embryo. (D) Co-expression of Bg-E75A (green) and Bg-ftz-F1 (magenta) before germband formation is complete. DAPI stain is in grey. Embryos shown in ventral view. (D′) greyscale image of D, showing Bg-ftz-F1 expression only. (D″) grayscale image of D, showing Bg-E75A expression only. (D′″) grayscale image of D, showing DAPI only. Asterisk marks broad stripes of Bg-ftz-F1 expression in the middle of the embryo. (E–H) Co-expression of Bg-E75A and Bg-ftz-F1 as germband formation is completing (E) and after germband formation (F–H). SAZ denotes segment addition zone. (E′, F′, G′) show zoomed-in views of the segment addition zones from the embryos directly above in the top panel. The stripe of Bg-E75A is always posterior to that of Bg-ftz-F1. Cells do not appear to express both genes simultaneously.

Immediately before germband formation (Fig. 1E) and during germband extension (Fig. 1F–H), we observed one to two stripes each of Bg-E75-A and Bg-ftz-F1 expression, in and just anterior to the segment addition zone (SAZ). In early germband extension, this pattern was repeated twice, generating two pairs of bands, each with Bg-E75A posterior to Bg-ftz-F1. At this early stage, the domain of Bg-E75A expression in the more posterior of the two pairs of bands was sometimes more of a broad zone of expression than a stripe (Fig. 1G’). As the embryo matured, only one pair of Bg-E75A and Bg-ftz-F1 stripes remained, but Bg-E75A was still posterior to Bg-ftz-F1 (Fig. 1H).

The germband elongates as new segments are formed from the SAZ. This allows us to reconstruct temporal dynamics of gene expression from spatial patterns of gene expression. Gene expression observed in anterior cells of the SAZ occurs later, developmentally, than gene expression observed in posterior SAZ cells. In the case of Bg-E75A and Bg-ftz-F1, we saw that Bg-E75A is always expressed posterior to BgFtz-F1, suggesting two possibilities: either cells first express Bg-E75A and then express Bg-ftz-F1, or Bg-E75A and Bg-ftz-F1 expressing-cells move anteriorly as a block together, each set of cells specified by one of the two genes.

Gb-E75A and Gb-ftz-F1 reproduce expression patterns of BgE75-A and BgFtz-F1 during abdominal segmentation.

To compare expression in the cockroach to another hemimetabolous insect we used HCR to analyze E75A and ftz-1 expression in Gryllus bimaculatus (Gb). We staged embryos based on their morphology upon dissection, and used a double stain with Gb-E75A and wingless (Gb-wg) to establish embryonic stages as defined by (Mito and Noji, 2008). During embryonic stage (ES) 2.6–4.3 when the gnathal and thoracic segments are forming (Fig. 2C), Gb-E75A and Gb-ftz-F1 were expressed in three broadly spaced stripes (Fig. 2A). Gb-ftz-F1 stripes were always anterior to Gb-E75A stripes, and cells did not express both Gb-E75A and Gb-ftz-F1 at the same time. We observed Gb-E75A stripes expressed synchronously with Gb-wg in alternating segments of the developing germband at two distinct time points (Supplemental Fig. 6). The overlap of Gb-E75A with alternating Gb-wg segments suggests that Gb-E75A is expressed in a pair-rule register. During G. bimaculatus abdominal segmentation (ES 4.4–5), Gb-E75A and Gb-ftz-F1 were expressed very similarly as their orthologs in B. germanica embryos at the same stage. We observed pairs of Gb-E75A and Gb-ftz-F1 stripes in and anterior to the SAZ (Fig. 2B). As with all other times and places these genes are expressed, E75A was posterior to Ftz-F1, and the genes were never co-expressed in the same cell. We also observed an additional stripe of Gb-E75A expression slightly anterior to the aforementioned pairs of Gb-E75A and Gb-ftz-F1 stripes (Fig. 2B, B’).

Fig. 2.

Fig. 2.

Open in a new tab

Expression of E75A and ftz-F1 in Gryllus bimaculatus embryos. (A) Gb-E75A and Gb-ftz-F1 are both expressed in three broadly spaced stripes during the stage at which gnathal and thoracic segments are specified. Stripes of Gb-E75A are always posterior to stripes of Gb-ftz-F1, and cells do not express both genes at the same time. Green shows Gb-E75A; magenta is Gb-ftz-F1 and DAPI is shown in grey. (A′) shows grey scale for Gb-E75A; (A″) shows grey scale for Gb-ftz-F1. (B) During abdominal segmentation, Gb-E75A and Gb-ftz-F1 are expressed in repeated stripes in and just anterior to the segment addition zone. As with embryos from the earlier stage shown in A, stripes of Gb-E75A are always posterior to stripes of Gb-ftz-F1, and cells do not express both genes at the same time. Colors the same as in panel A. (B′) Zoomed in view of the segment addition zone of embryo shown in B. (B″) shows grey scale for Gb-E75A for the zoomed in view; (B′″) shows grey scale for Gb-ftz-F1 for the zoomed in view. (C, D) Expression of Gb-E75A and Gb wingless (Gb-wg) during gnathal and thoracic segmentation (C) and abdominal segmentation (D). Gb-E75A in green; Gb-wg in cyan. (C) The first two Gb-E75A stripes (from anterior to posterior) overlap with Gb-wg stripes of the labial and second thoracic segment, respectively. The third and most posterior of the Gb-E75A stripes appears posterior to Gb-wg stains in mature segments. (D) During abdominal segmentation, Gb-E75A stripes are posterior to all Gb-wg stains in mature segments. an = antennal segment, mn = mandibular segment, mx = maxillary segment, lb = labial segment, T1 =first thoracic segment, T2 = second thoracic segment, T3 = third thoracic segment, A1 =first abdominal segment.

2.3. Bg-ftz-F1 knockdown results in malformed germbands

We conducted several different RNAi experiments in B. germanica embryos to assess the function of Bg-E75A and Bg-ftz-F1 during development. ftz-F1 has a known role in the oogenesis of many insect species, including B. germanica (Borras-Castells et al., 2017). We repeated previous results showing that injection of Bg-ftz-F1 dsRNA into B. germanica females arrests oogenesis and causes degradation of oocytes (Supplemental Fig. 7). Going forward, we reasoned that there might be a time window in which we could inject Bg-ftz-F1 dsRNA after most of oogenesis had occurred but before egg lay. Female B. germanica lay their eggs into egg cases (oothecae) which remain attached to the female for the duration of embryogenesis. In our laboratory, this ootheca formation happens approximately 10 days after eclosion. We experimented with injecting Bg-ftz-F1 dsRNA at different time points post terminal molt (Supplemental Table 1). When injected with Bg-ftz-F1 dsRNA at 6–7 days post terminal molt, 4 out of 13 females successfully made oothecae (30.8%). When the same injection was given 7–8 days post terminal molt, 11 out of 19 females extruded oothecae (57.9%). Injections with control dsRNA targeting a region of the Promega pGEM vector yielded 100 percent egg formation (n = 9 females at 6–7 days; n = 2 females 7–8 days). Sometimes, if enough of the eggs in an ootheca are non-viable, a female B. germanica will drop the ootheca and all the embryos within will die. Of the females who made oothecae, there was no difference in the rate of those who dropped their oothecae shortly after formation between our experimental and control injectees (2/15 total oothecae dropped for the Bg-ftz-F1 dsRNA females, and 1/11 dropped for the pGEM dsRNA injectees). We found significant reduction in Bg-ftz-F1 transcripts in four-day-old oothecae from treated mothers when compared to expression of Bg-ftz-F1 in oothecae of the same stage from mothers injected with pGEM dsRNA (Supplemental Fig. 8).

We dissected a subset of the RNAi-treated oothecae 3–5 days after extrusion in order to observe the effect of Bg-ftz-F1 on germband formation. We left the rest of the oothecae to develop to hatchling stage, with the aim of observing cuticular defects. Within the oothecae dissected, we observed a range of phenotypes (Fig. 3). One of the oothecae dissected did not contain any wild type-like embryos, and four of those dissected contained a mix of wild type-like and affected embryos. We saw two types of abnormalities in Bg-ftz-F1 dsRNA oothecae. Some embryos failed to form germbands properly, with a patchy distribution of cells during germband condensation (Fig. 3A–D). For embryos that successfully formed germbands, we used HCR to examine gene expression in the knockdown embryos (Fig. 3). Embryos from mothers who had been injected with Bg-ftzf-F1 dsRNA showed barely detectable Bg-ftz-F1 expression (Fig. 3E’, G′), indicating successful knockdown of target gene expression. Variable Bg-E75A expression was detected in embryos from mothers injected with Bg-ftzf-F1 dsRNA (Fig. 3E’, G″). We investigated the effect of Bg-ftz-F1 dsRNA on segmentation by using HCR to examine expression of the segment polarity gene hedgehog (Bg-hh) in knockdown embryos. We observed wild type-like segment formation (i.e., no fusion of segments or change in segment width) despite failure of germbands to properly form in embryos from mothers injected with Bg-ftz-F1 dsRNA (Fig. I – K’). We distinguished between germbands that failed to form properly and general delayed development by counting the number of Bg-hh stripes in each embryo. While wild-type embryos have well-formed germbands and head lobes by the time abdominal segmentation is under way (two abdominal Bg-hh stripes), embryos from mothers injected with Bg-ftz-F1 dsRNA showed diffuse germbands, especially in the head region. It is worth noting that (Cruz et al., 2022) observed severe head defects when knocking down Bg-HR3. As Bg-HR3 is part of the ecdysone cascade in B. germanica, our results in conjunction with Cruz et al. suggest that ecdysone signaling may be required for head lobe formation in the German cockroach.

Fig. 3. Knockdown of BgFtz-F1 reduces expression of BgE75-A during abdominal segmentation and results in failed germband formation.

Fig. 3.

Open in a new tab

(A–D) Embryos from mothers injected with dsBgFtz-f1 form disorganized germbands with patches in aggregating cells. (A, B): Sytox stain of representative embryos from an oothecae from a control female injected with dspGEM. (C, D): Sytox stain of representative embryos from an oothecae from a female injected with dsBgFtz-f1. (E-H″): Expression of BgFtz-f1 and BgE75-A in embryos from mothers injected with dsBgFtz-f1 (E-E″, G-G″) and comparably staged wild type embryos (F–F″, H–H″). All panels in E-H″ show the posterior end of the embryo. Embryos shown in E, F, G, and H are composites with BgFtz-f1 in magenta, BgE75-A in green and DAPI. Panels E′, E″, F′, F″, G′, G″, and H′, H″ are grey scale images of E, F, G, and H respectively. Top row shows embryos from the gnathal-thoracic stage of segmentation; bottom row shows embryos from abdominal segmentation stage. Embryos from mothers injected with dsBgFtz-f1 show reduced BgFtz-f1 expression (E′, G′ as compared with F′, H′). BgE75-A is still detected in an early embryo from a mother injected with dsBgFtz-f1 (E″) but it is almost absent in the abdominal segmentation stage (G″) of an embryo from a mother injected with dsBgFtz-f1. In contrast, the abdominal stage in a wildtype embryo shows a strong stripe of BgE75-A expression (H″). (I–K′) Embryos from oothecae from females treated with dsBgFtz-f1 show normal segment formation but disrupted germband condensation. HCR staining for BgFtz-f1 shown in grey in (I, J, K). Bghh (cyan) with DAPI (blue) shown in (G′, H′, I′). (I,I′): Wild type B. germanica embryo shown at stage with 2 abdominal segments (10 stripes total). (J, J′): dsftzf1 embryo shown at the same stage, with 2 abdominal segments. Despite being the same developmental age as the embryo in I, I′, the germband has not properly formed, evidenced most notably in the head region. (K, K″): dsftzf1 embryo shown at stage with 11 hh stripes.

Of the oothecae allowed to develop, we did not observe any defects in hatchlings, suggesting that any embryo affected by Bg-ftz-F1 dsRNA was not capable of successfully developing to the hatchling stage, or that some fraction of embryos with a Bg-ftz-F1 knockdown can recover.

In sum, knocking down Bg-ftz-F1 resulted in non-specific failures of germband formation rather than problems with segment formation specifically.

2.4. Bg-E75A knockdown results in abdominal truncation

Bg-E75A is one of five E75 transcripts expressed in early B. germanica development. To examine its role in segmentation, we injected 5- to 7-day-old females with dsRNA targeting just the region specific to the A isoform (the first 200 bp of the CDS, Supplemental Fig. 2). Of 11 females injected, 9 produced oothecae, all of which contained either wild type-like embryos when dissected at 4–5 days post ootheca emergence or embryos which developed into viable and wild type-like hatchlings when left undisturbed. To account for the fact that our lack of phenotype might be related to the short length of the dsRNA injected, or its location in the 5′UTR, we then injected a longer dsRNA fragment which contained all of the sequence unique to Bg-E75A but also some downstream sequence common to other E75 isoforms (Supplemental Fig. 2). Seventy-six percent (13 out of 17) of females injected with the longer Bg-E75A dsRNA fragment formed oothecae (Supplemental Table 1). We dissected 8 of these oothecae approximately 5 days after ootheca extrusion, the stage at which wild type embryos would already have appendages. In two of the oothecae from mother injected with Bg-E75A dsRNA, we found a small number of embryos with truncated abdomens (n = 3 affected embryos in one ootheca, and n = 1 affected embryo in a second ootheca) (Fig. 4, panels A–E). These embryos represented a small fraction of the ~30 embryos in each ootheca. The truncated abdomen phenotype resembles an RNAi phenotype observed in O. fasciatus when broad, an ecdysone response gene, was knocked down during embryogenesis (Erezyilmaz et al., 2009a,b). In four other dissected oothecae, development was generally delayed for all embryos compared to controls. In some of these embryos, the germband was not yet fully extended at a time when control embryos already had appendages forming. We used some of these developmentally delayed embryos for HCR to examine expression of Bg-E75A, Bg-ftz-F1, and Bg-hh. Relative to control embryos, expression of both Bg-E75A and Bg-ftz-F1 was less intense in those embryos whose mothers had been injected with dsE75A long (Fig. 4, panels F–H″), indicating that the Bg-E75A knockdown had reduced Bg-E75A RNA levels, which in turn had an impact on Bg-ftz-F1 expression. Because the dsRNA fragment we injected overlapped with all of the sequence unique to the A isoform, we were not able to perform reliable qPCR to assess the impact of our RNAi treatment on Bg-E75A specifically. Despite the fact that Bg-ftz-F1 expression appeared reduced in HCR images from Bg-E75A dsRNA embryos, we did not observe failure of germbands to properly form in ootheca from mothers injected with Bg-E75A dsRNA, as we had for Bg-ftz-F1 knockdowns. We note that our qPCR results did not show a definitive knockdown of Bg-ftz-F1 in Bg-E75A dsRNA embryos. The discrepancy between the HCR and qPCR could be attributed to variable rates of knockdown, as qPCR was performed on pooled embryos from a single ootheca.

Fig. 4. Knockdown of BgE75-A reduces expression of ftz-f1 during abdominal segmentation and occasionally results in embryos without abdomens.

Fig. 4.

Open in a new tab

(A–E) dsBgE75-A and dspGEM embryos at the appendage forming stage. Embryos A, B, C were all dissected from the same ootheca and show a range of phenotypes, with embryo C not seemingly affected by the dsRNA. Embryo E comes from a second ootheca treated with dsBgE75A, and embryo E shows a similarly aged embryo from a dspGEM treated ootheca. (F–H″) Zoomed in view of the posterior of embryos from mothers injected with dsBgE75A and wild-type reference showing HCR double in-situ (F, G, H) and single channels for BgE75-A (F′,G′,H′) and BgFtz-f1 (F″, G″, H″). Each row in panels F–H″ are images from the same embryo. In composite images (A, D, G) BgFtz-f1 is in magenta; BgE75-A is in green, and DAPI is shown in grey. Expression of both BgE75-A and BgFtz-f1 is reduced in BgE75-A knockdown embryos. (I–J) dsE75A knockdown does not directly impact Bghh expression. (I) embryo from a mother treated with dsBgE75A showing DAPI (blue) and hh (cyan). (I′) BgFtz-f1 expression in the same embryo. (J) WT embryo with the same number (n = 10) Bghh stripes (cyan) as dsBgFtz-f1 treated embryo in panel I. DAPI shown in blue. (J″) BgFtz-f1 expression in embryo from panel J.

We did not detect any impact of Bg-E75A knockdown on Bg-hh expression (Fig. 4, panels I–J″), indicating that segment formation still occurred despite the knockdown of Bg-E75A. As a general note, knockdown embryos appear to lack the constriction found at the anterior most part of the segment addition zone in wild type and control embryos (Fig. 4F–J’). It is possible that this lack of constriction and the abdominal truncation phenotypes were both related to patterns of Bg-E75A observed during abdominal segmentation in wild-type embryos. Lastly, we dissected three oothecae at later stages in development, and found general developmental defects (embryos lacking appendages, or dead embryos) in two out of these three oothecae (Supplemental Fig. 9).

To further test the interaction between Bg-E75A and Bg-ftz-F1, we performed a double knockdown by injecting five females with the short, specific Bg-E75A dsRNA fragment together with Bg-ftz-F1 dsRNA. Of these females, only one made an ootheca, which was deformed and dropped immediately after extrusion (Supplemental Table 1).

3. Conclusions

We have shown here that two genes from the ecdysone signaling cascade, ftz-F1 and E75A, are expressed in segmental patterns during abdominal segmentation of two hemimetabolous insects. In the cricket G. bimaculatus, both genes are also expressed in three broadly spaced stripes throughout the embryonic trunk during gnathal-thoracic segmentation. These stripes corresponded to every other band of Gb-wg expression, suggesting PR-like expression for each of these genes. In the cockroach B. germanica, only ftz-F1 is expressed in three broadly spaced stripes at this time. Bg-E75A appears expressed only in the posterior of the B. germanica embryo. In both G. bimaculatus and B. germanica, cells either expressed ftz-F1 or E75A. At no point in embryogenesis have we observed cells co-expressing these genes, in either species.

RNAi knockdown suggests that in B. germanica, Bg-ftz-F1 is necessary for the formation of the germband, specifically for fusion of lateral plates of cells in the head lobes and that Bg-E75A is necessary for growth of the abdomen. We observed a reduction in expression in Bg-ftz-F1 when Bg-E75A is knocked down, indicating either an indirect or direct regulation of Bg-ftz-F1 by Bg-E75A. The inability of females to produce viable offspring when injected with dsRNA targeting both genes, but not when injected with dsRNA targeting either one alone, suggests there could be some compensatory mechanism at play in the single knockdowns.

While the ecdysone pulse at the end of insect embryogenesis has been relatively well studied (Ruaud et al., 2010), less attention has been given to the role that ecdysone could play in early embryogenesis. This lack of attention is especially notable given that many papers which report spikes of ecdysone during embryogenesis measure either total ecdysone levels or ecdysone levels relative to the weight of input sample, which contains both embryo and yolk (Lagueux et al., 1979; Maróy et al., 1988 ; Mané-Padrós et al., 2008; Matsushima et al., 2019). Considering that the number of cells in an embryo shortly before hatching is orders of magnitude larger than the number of cells in a developing germband, ecdysone titer scaled to embryonic cell number would be a more appropriate measurement of hormone levels during embryogenesis. Recent work from (Cruz et al., 2022) demonstrates that ecdysone signaling is necessary for the early stages of B. germanica embryogenesis. In this paper, we have shown that two genes in the ecdysone signaling network, E75A and ftz-F1 are expressed segmentally in both B. germanica and G. bimaculatus segmentation. The potential involvement of the ecdysone signaling network in segmentation is intriguing, as previous work has shown E75A acts as a pair rule gene in the hemipteran Oncopeltus fasciatus. We also note that both E75A and ftz-F1 are expressed in a pattern resembling a pair-rule register (i.e., expressed in every other segment primordium) in G. bimaculatus. This expression data suggests that even though pair-rule patterning has not been functionally demonstrated in hemimetabolous insects outside of O. fasciatus, there could be some ecdysone-mediated patterning of every other segment that occurs during the formation of gnathal and thoracic segments in the two-spotted cricket.

4. Materials and methods

4.1. Insect husbandry

G. bimaculatus and B. germanica cultures were kept at 25 °C, with 40 percent humidity and were fed oats and dog food ad libitum. To collect B. germanica embryos, we selected females in the process of ootheca extrusion, and we set aside these females until embryos had aged to the desired stage. Under our laboratory conditions, germband condensation was completed approximately four days after extrusion started. We collected G. bimaculatus embryos from wet cotton into which females lay eggs.

4.2. Gbftz-F1 and GbE75-A gene isolation

All RNA was extracted with Trizol using the manufacturer’s protocol. For G. bimaculatus, RNA was extracted from 24 to 72 h eggs. For B. germanica, RNA was extracted from unstaged oothecae.

We used sequences deposited in NCBI to design in-situ probes and dsRNA templates for B. germanica E75A (Bg-E75A, AM238653.1) and ftz-F1 (Bg-ftz-F1, FM163377.1).

Fragments of Gb-ftz-F1 and Gb-E75A were first amplified by degenerate PCR using Phusion polymerase from New England Biolabs (CAT No. M0530S.) Reaction was prepared as follows: 10 μL GC buffer, 1 μL dNTP, 1 μL cDNA, 0.5 μL Phusion polymerase, 1.5 μL DMSO, water to 50 μL. The following cycling parameters were used − 98 °C for 3 min, followed by 35 cycles of: 98 °C for 10 s, Tm for 30 s, 72 °C for 30 s and then 72 °C for 10 min.

To obtain a longer Gb-ftz-F1 sequence, we used the ftz-F1 sequence from Oncopeltus fasciatus (Y. Lu, 2015) as bait in a tBLASTn search of the G. bimaculatus genome and we recovered an exon upstream of the degenerate PCR. RT-PCR using a forward primer in this exon and a reverse primer in the sequence amplified by degenerate PCR yielded a product of approximately 1 kb. We used SeqAmp polymerase from Takara for this PCR with the following cycling parameters: 95 °C for 1 min, then 35 cycles of 98 °C for 30 s, 61 °C for 30 s, 72 °C for 90 s, followed by a 3 min hold at 72 °C. A reverse primer for 5′ RACE was designed from within the sequence cloned out from this PCR.

We performed 5′ Rapid Amplification of cDNA Ends (RACE) for E75A and ftz-F1 in G. bimaculatus using the Takara SMARTer 5’/3′ RACE kit (Cat No. 634858). The reverse primer for the 5′ RACE reaction amplifying Gb-E75A was designed using sequence amplified by degenerate PCR as described above. RACE library construction was performed with 1 μg of input RNA and according to the manufacturer’s protocol. All RACE PCR was performed with SeqAmp polymerase from Takara. We obtained two bands when performing a 5′ RACE reaction for GbE75 with 30 cycles of the following parameters: 94 °C - 30 s, 62.5 °C - 30 s, 72 °C - 3 min. Both bands were cut out from a 1 percent agarose gel, and DNA was individually extracted from each using a gel extraction kit from Hylabs (Cat No. IMDF100/300). These were inserted into the pRACE vector (Takara), transformed, and sequenced. For Gb-ftz-F1, we performed PCR with 30 cycles of the following − 94 °C - 30 s, 64 °C - 30 s, 72 °C - 2 min 30 s. Bands were cut out and cloned into pRACE as described above for Gb-E75.

4.3. Colorimetric in-situ probe and dsRNA template preparation

cDNA was synthesized from RNA prepared as described above. One microgram of RNA was used in a reverse transcription reaction primed with a 1:1 mixture of oligodT:random hexamers. We used Bioline’s RTase (Cat No BiO-27036). Templates for B. germanica probes and dsRNA were amplified using Tiger polymerase from Hylabs (EZ-2031). Template for the Gb-wg probe was amplified using Takara’s SeqAmp Polymerase (Cat No 638504). To amplify template for in-situ probes, we added T7 sequence to the 5′ end of the reverse primer, and to amplify template for dsRNA fragments, we added T7 sequence to the 50 ends of both primers. To amplify templates for the Bg-E75A probe and dsRNA, we used the following cycling parameters: 95 °C for 3 min, followed by 32 cycles of 95 °C for 30 s, 55 °C for 30 s, and 72 °C for 30 s, with a 3 min extension at 72 °C at the end of the cycles. To amplify template for the Bg-ftz-F1 probe and dsRNA, we used the following cycling parameters: 95 °C for 3 min, followed by 32 cycles of 95 °C for 30 s, 59 °C for 30 s, and 72 °C for 1 min, with a 3 min extension at 72 °C at the end of the cycles. To amplify template for the Gb-wg probe, we used the following cycling parameters: 98 °C for 2 min, followed by 36 cycles of 98 °C for 10 s, 60 °C for 15 s, and 68 °C for 1 min, with a 3 min extension at 68 °C at the end of the cycles. Templates were purified using Macherey-Nagel’s NucleoSpin Gel and PCR Clean-up kit (740609.50). We synthesized DIG-labeled probes with T7 polymerase from Roche (10881767001). dsRNA was synthesized using ThermoFisher’s MEGAScript kit (AM1626M). Transcription reactions for probes incubated for 2.5–3 h before lithium chloride precipitation. dsRNA was synthesized overnight and followed the MEGAScript kit protocol. As a negative control for the dsRNA, sequence from Promega’s pGEM vector (A1360) between the M13F and M13R primers was amplified, and then SP6 and T7 polymerases were used to transcribe dsRNA.

4.4. dsRNA injection

Newly eclosed females were collected and put into a small container with adult males. Female:male ratio was approximately 1:2, depending on availability of adult males. After 5–8 days, females were injected (see results section for discussion of different ages used) with 1.5–3 μg of dsRNA in 1.2 μL of water between sternites three and four.

4.5. qPCR

Embryos from a single egg case from a mother injected with either BgFtz-F1 dsRNA, BgE75-A dsRNA, or pGEM dsRNA were dissected and pooled into one sample. RNA was extracted as described in the Gene Isolation section, and cDNA was synthesized as described in the Colorimetric in-situ probe and dsRNA template preparation section. qPCR was performed in 10 μL reactions (with primers at X concentration) using Applied Biosystems Fast SYBR Green Master Mix (cat no 4385616) and run on a Quant Studio 3 system with an extension temperature of 59 °C. Before experimental qPCR was run, all primers used were validated and found to have efficiencies between 92 and 104 percent. Relative expression of target genes (BgFtz-F1 and BgE75-A) in treated samples was calculated relative the geometric mean of expression of two housekeeping genes, actin and cyclophilin, using the Pfaffl method (Pfaffl, 2001).

4.6. Embryo dissection and fixation

The dissection and fixation protocols described below were used for both colorimetric in-situs and HCR. B. germanica oothecae were removed manually, placed in 1 mL of water, and heated at 85 °C for 10 min followed by 2 min on ice. Embryos were then dissected out in phosphate buffered saline with one percent tween (PBST) and placed directly into four percent formaldehyde. Embryos were fixed in four percent formaldehyde for 1 h and then gradually dehydrated to 100 percent methanol. Embryos were stored in methanol at −20 °C for at least overnight; up to several months. G. bimaculatus eggs were collected from wet cotton and rinsed several times in distilled water. The eggs were then dechorionated in 30 percent bleach for 3 min, and then rinsed three times in phosphate buffered saline. We manually peeled the vitellin membrane off the eggs after they were dechorionated, and embryos were then dissected away from the yolk. Both steps were done in PBST. Freshly dissected embryos were fixed in four percent formaldehyde for 1 h and then gradually dehydrated to 100 percent methanol. Embryos were stored in methanol at −20 °C for at least overnight; up to several months.

4.7. Colorimetric in-situ

Embryos were gradually rehydrated from 100 percent methanol to 75 percent methanol in phosphate buffered saline (PBST) to 50 percent methanol in PBST to 25 percent methanol in PBST to 100 percent PBST with 4 min in each intermediate step. After 3 × 1 minute PBST washes, a post fixation of 20 min in 4 percent formaldehyde in PBST and another round of 3 × 1 minute PBST washes were carried out. Embryos were incubated in hybridization buffer at 65 °C for 5 min, then hybridization buffer was refreshed and then embryos were incubated again in hybridization buffer at 65 °C for 2–4 h. DIG-labeled probes were applied at a concentration of 1 ng/uL in hybridization buffer and left overnight. The following day, the following washes were performed: 20 min in preheated hyb buffer (65 °C); 3 × 20 min in 2x SSC/0.1% Tween-20 (65 °C); 3 × 20 min in 0.2x SSC/0.1% Tween-20 (65 °C); 2 × 10 min in PBST at room temp; 1 h in blocking solution (5 percent sheep or goat serum, 2 mg/mL BSA, 1 percent DMSO in PBST); 4 h in 1:1500 solution of anti-digoxigenin antibody (Roche, Cat No. 11333089001):blocking solution; 3 × 1 minute PBST washes, and left overnight in PBST at 4 °C. The following day, embryos were washed in the following series: 4 × 20 min wash in PBST; 2 × 10 min of washing in freshly prepared staining solution (0.1 M 9.5 TrisHCl; 0.05M MgCl2; 0.1M NaCl; 0.1M Tween-20); 1 × 10 min of staining solution + PVA (0.1 M 9.5 TrisHCl; 0.05M MgCl2; 0.1M NaCl; 0.1M Tween-20; 0.025 percent polyvinyl alcohol). 20 μL of NBT/BCIP from Roche in 980 μL of staining solution + PVA was used for color development, which continued up to 4 h. Staining reactions were stopped when we observed background stain developing. Staining reactions were stopped with PBST washes and fixed with 50 percent methanol:PBST. Embryos were imaged in 70 percent glycerol.

4.8. Hybridization chain reaction

We used probes designed by Molecular Instruments for Bg-E75A (lot number PRK282), Bg-ftz-F1 (lot number PRJ351), Bg-hh (lot number PRK420), Gb-E75A (lot number PRO035), and Gb-ftz-F1 (lot number PRO034). For probe design, we provided Molecular Instruments with the entirety of each gene’s available coding sequence. Probes designed by Molecular Instruments target several dozen nucleotides each and span the entirety of the coding sequence in a non-contiguous fashion. Probes for Bg-E75A, Bg-hh, and Gb-E75A were all designed to anchor an amplification reaction with a 488 fluorophore. Probes for Bg-ftz-F1, and Gb-ftz-F1 were designed to anchor an amplification reaction with a 594 fluorophore.

We attempted HCR for Bg-E75A with probes targeting both the CDS region specific to the A isoform (~200 bp) and with probes targeting this specific region and downstream sequence common to other isoforms (Supplemental Fig. 2). A much better signal was obtained with probes targeting a larger region, and we confirmed that the expression patterns detected via HCR with the more expansive probe set is identical to the expression pattern seen with a colorimetric in-situ done with a probe specific to the A isoform (Supplemental Fig. 10).

For HCR-RNA FISH, a modified version of the protocol described by Bruce et al., 2021 (dx.doi.org/10.17504/protocols.io.bunznvf6) was used. Hybridization buffer, wash buffer, and amplification buffer were provided by Molecular Instruments, and we made the detergent solution (1 percent SDS, 0.5 percent Tween, 50 mM Tris-HCl (pH 7.5), 1 mM EDTA (pH 8.0), 150 mM NaCl) as described by Bruce et al., 2021. Embryos were rehydrated from methanol to PBST as described above in the colorimetric in-situ section and then washed 1 × 10 min, 2 × 5 min in PBST. Embryos were incubated at RT for 30 min in the detergent solution, then 30 min at 37 °C 37 °C in 200 μL of hybridization buffer. Probes were prepared at a concentration of approximately 10 nM, made by diluting 1.6 μL of the 1 μM stock from Molecular Instruments into 150 μL of hybridization buffer. Embryos were incubated overnight at 37 °C in the probe solution. The following day, embryos were washed in 1 mL of wash buffer at 37 °C (4 × 15 min), followed by 2 × 5 minute washes at RT with five percent SSCT (5x sodium chloride sodium citrate with 1 percent Tween). Embryos were incubated in amplification buffer (Molecular Instruments) for 30 min at RT while hairpins were snap cooled by heating to 95 °C for 30 s followed by 30 min at RT in the dark. 4 μL of each hairpin were added to 100 μL of amplification buffer. This solution was applied to embryos after removing the 1 mL of amplification buffer used for the pre-amplification step. Embryos were then incubated overnight at RT in the dark. Embryos were washed the next morning with five percent SSCT washes at RT (volume 1 mL) twice for 5 min and once for 15 min. We then did a 15 min wash with 1 μL DAPI into 1 mL of five percent SSCT, followed by two 15 min washes with five percent SSCT. Embryos were placed in 50% glycerol/PBS for 30 min before being transferred to 70% glycerol in PBS and mounted on slides.

4.9. Imaging

Fluorescent images were acquired with an Olympus FV1200 confocal based on an IX-83 inverted microscope (Olympus, Japan). Images were processed in Fiji (Fiji is just ImageJ) (Schindelin et al., 2012). For each fluorescent channel, we selected and projected the Z-stacks which contained signal, as determined subjectively by eye. After projecting stacks, we adjusted the contrast and brightness of each channel (using the window, level, contrast, and brightness scales in Fiji) to maximize signal from the HCR. We then false colored and merged channels.

Supplementary Material

MMC2
MMC1

Acknowledgements

We are grateful to Dr. Naomi Melamed-Book from The Bioimaging Facility at The Hebrew University in Jerusalem for help with confocal imaging. We thank Dr. Alys Jarvela for performing two degenerate PCRs for ftz-F1 and E75 in Gryllus bimaculatus. Thanks very much to Dr. Jarvela and to Katie Reding for thoughtful comments on the manuscript. This work was supported by a Zuckerman Fellowship (J.R.W); the National Institutes of Health (R01GM113230 to L.P.); and the Israeli Science Foundation (120/16 to A.C.). We thank two anonymous reviewers for helpful feedback on the manuscript.

Footnotes

Appendix A. Supplementary data

Supplementary data to this article can be found online at https://doi.org/10.1016/j.ydbio.2023.03.008.

Data availability

Data will be made available on request.

References

  1. Ashburner Michael, Chihara Carol, Meltzer Paul, Richards Geoff, 1974. Temporal control of puffing activity in polytene chromosomes. Cold Spring Harbor Symp. Quant. Biol. 38 (January), 655–662. 10.1101/SQB.1974.038.01.070. [DOI] [PubMed] [Google Scholar]
  2. Becker Hans Joachim, 1962. Die Puffs der Speicheldrüsenchromosomen von Drosophila melanogaster. Chromosoma 13 (4), 341–384. 10.1007/BF00327339. [DOI] [Google Scholar]
  3. Beermann W, 1952. [Chromomore constancy and specific modifications of the chromosome structure in development and organ differentiation of Chironomus tentans]. Chromosoma 5 (2). Unknown. [PubMed] [Google Scholar]
  4. Bialecki Michael, Shilton Alycia, Fichtenberg Caroline, Segraves William A., Thummel Carl S., 2002. Loss of the ecdysteroid-inducible E75A orphan nuclear receptor uncouples molting from metamorphosis in Drosophila. Dev. Cell 3 (2), 209–220. 10.1016/S1534-5807(02)00204-6. [DOI] [PubMed] [Google Scholar]
  5. Borras-Castells Ferran, Nieva Claudia, Maestro Jose L., Maestro Oscar, Belles Xavier, Martín David, 2017. Juvenile hormone biosynthesis in adult Blattella germanica requires nuclear receptors seven-up and FTZ-F1. Sci. Rep. 7 (1), 40234. 10.1038/srep40234. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Broadus Julie, McCabe Jennifer R., Endrizzi Bart, Thummel Carl S., Woodard Craig T., 1999. The Drosophila ВFTZ-F1 orphan nuclear receptor provides competence for stage-specific responses to the steroid hormone ecdysone. Mol. Cell 3 (2), 143–149. 10.1016/S1097-2765(00)80305-6. [DOI] [PubMed] [Google Scholar]
  7. Bruce Heather S., Jerz Gabby, Kelly Sophia, McCarthy Jenny, Pomerantz Aaron, Senevirathne Gayani, Sherrard Alice, Sun Dennis A., Wolff Carsten, Patel Nipam H., 2021. Hybridization Chain Reaction (HCR) In Situ Protocol. July. https://www.protocols.io/view/hybridization-chain-reaction-hcr-in-situ-protocol-bunznvf6. [Google Scholar]
  8. Clever U, 1964. Actinomycin and puromycin: effects on sequential gene activation by ecdysone. Science (New York, N.Y.) 146 (3645), 794–795. 10.1126/science.146.3645.794. [DOI] [PubMed] [Google Scholar]
  9. Cruz Josefa, Maestro Oscar, Franch-Marro Xavier, Martín David, 2022. Ecdysone Signaling Controls Early Embryogenesis in the Short-Germ Hemimetabolous Insect Blattella Germanica.” bioRxiv. 10.1101/2022.03.10.483750. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Cruz Josefa, Nieva Claudia, Mané-Padrós Daniel, Martín David, Bellés Xavier, 2008. Nuclear receptor BgFtz-F1 regulates molting and the timing of ecdysteroid production during nymphal development in the hemimetabolous insect Blattella germanica. Dev. Dynam.: Off. Publ. Am. Assoc. Anat 237 (11), 3179–3191. 10.1002/dvdy.21728. [DOI] [PubMed] [Google Scholar]
  11. Davis Gregory K., Patel Nipam H., 2002. SHORT, long, and beyond: molecular and embryological approaches to insect segmentation. Annu. Rev. Entomol. 47 (1), 669–699. 10.1146/annurev.ento.47.091201.145251. [DOI] [PubMed] [Google Scholar]
  12. Erezyilmaz Deniz F., Kelstrup Hans C., Riddiford Lynn M., 2009a. The nuclear receptor E75A has a novel pair-rule-like function in patterning the milkweed bug, Oncopeltus fasciatus. Dev. Biol. 334 (1), 300–310. 10.1016/j.ydbio.2009.06.038. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Erezyilmaz Deniz F., Rynerson Melody R., Truman James W., Riddiford Lynn M., 2009b. The role of the pupal determinant broad during embryonic development of a direct-developing insect. Dev. Gene. Evol. 219 (11–12), 535–544. 10.1007/s00427-009-0315-7. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Fernandez-Nicolas Ana, Machaj Gabriela, Ventos-Alfonso Alba, Pagone Viviana, Minemura Toshinori, Ohde Takahiro, Daimon Takaaki, Ylla Guillem, Belles Xavier, 2023. Reduction of embryonic E93 expression as a hypothetical driver of the evolution of insect metamorphosis. Proc. Natl. Acad. Sci. USA 120 (7), e2216640120. 10.1073/pnas.2216640120. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Field Amanda, Xiang Jie, Anderson Ray, Graham W, Patricia Pick, Leslie, 2016. Activation of ftz-F1-responsive genes through ftz/ftz-F1 dependent enhancers. PLoS One 11 (10), e0163128. 10.1371/journal.pone.0163128. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Florence B, Guichet A, Ephrussi A, Laughon A, 1997. Ftz-F1 is a cofactor in ftz activation of the Drosophila engrailed gene. Development 124 (4), 839–847. 10.1242/dev.124.4.839. [DOI] [PubMed] [Google Scholar]
  17. Guichet Antoine, Copeland John W.R., Erdelyi Miklos, Hlousek Daniela, Závorszky Péter, Ho Jacqueline, Brown Susan, Percival-Smith Anthony, Krause Henry M., Ephrussi Anne, 1997. The nuclear receptor homologue ftz-F1 and the homeodomain protein ftz are mutually dependent cofactors. Nature 385 (6616), 548–552. 10.1038/385548a0. [DOI] [PubMed] [Google Scholar]
  18. Heffer Alison, Grubbs Nathaniel, James Mahaffey, Pick Leslie, 2013. The evolving role of the orphan nuclear receptor ftz-F1, a pair-rule segmentation gene. Evol. Dev 15 (6), 406–417. 10.1111/ede.12050. [DOI] [PubMed] [Google Scholar]
  19. Hernandez Jessica, Pick Leslie, Katie Reding, 2020. Oncopeltus-like gene expression patterns in Murgantia histrionica, a new Hemipteran model system, suggest ancient regulatory network divergence. EvoDevo 11 (1), 9. 10.1186/s13227-020-00154-x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Hou Hui Ying, Heffer Alison, Anderson Ray, Liu W, Jingnan, Bowler Timothy, Pick Leslie, 2009. Stripy ftz target genes are coordinately regulated by ftz-F1. Dev. Biol. 335 (2), 442–453. 10.1016/j.ydbio.2009.08.004. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. King-Jones Kirst, Thummel Carl S., 2005. Nuclear receptors — a perspective from Drosophila. Nat. Rev. Genet. 6 (4), 311–323. 10.1038/nrg1581. [DOI] [PubMed] [Google Scholar]
  22. Lagueux M, Hetru C, Goltzene F, Kappler C, Hoffmann JA, 1979. Ecdysone titre and metabolism in relation to cuticulogenesis in embryos of Locusta migratoria. J. Insect Physiol. 25 (9), 709–723. 10.1016/0022-1910(79)90123-9. [DOI] [Google Scholar]
  23. Lu Shennan, Wang Jiyao, Chitsaz Farideh, Derbyshire Myra K., Geer Renata C., Gonzales Noreen R., Gwadz Marc, et al. , 2020. CDD/SPARCLE: the conserved domain database in 2020. Nucleic Acids Res. 48 (D1), D265–D268. 10.1093/nar/gkz991. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Lu Yong, 2015. Evolution of Pair-Rule Genes. Doctoral Dissertation University of Maryland. [Google Scholar]
  25. Maestro Oscar, Cruz Josefa, Pascual Núria, Martín David, Bellés Xavier, 2005. Differential expression of two RXR/ultraspiracle isoforms during the life cycle of the hemimetabolous insect Blattella germanica (dictyoptera, blattellidae). Mol. Cell. Endocrinol. 238 (1), 27–37. 10.1016/j.mce.2005.04.004. [DOI] [PubMed] [Google Scholar]
  26. Mané-Padrós Daniel, Cruz Josefa, Vilaplana Lluisa, Nieva Claudia, Ureña Enric, Bellés Xavier, Martín David, 2010. The hormonal pathway controlling cell death during metamorphosis in a hemimetabolous insect. Dev. Biol. 346 (1), 150–160. 10.1016/j.ydbio.2010.07.012. [DOI] [PubMed] [Google Scholar]
  27. Mane-Padros Daniel, Cruz Josefa, Vilaplana Lluïsa, Pascual Nuria, Bellés Xavier, Martín David, 2008. The nuclear hormone receptor BgE75 links molting and developmental progression in the direct-developing insect Blattella germanica. Dev. Biol. 315 (1), 147–160. 10.1016/j.ydbio.2007.12.015. [DOI] [PubMed] [Google Scholar]
  28. Marie Bruno, Bacon JP, 2000. Two engrailed-related genes in the cockroach: cloning, phylogenetic analysis, expression and isolation of splice variants. Dev. Gene. Evol. 210 (8), 436–448. 10.1007/s004270000082. [DOI] [PubMed] [Google Scholar]
  29. Maroy Peter, Kaufmann Gabrielle, Dübendorfer Andreas, 1988. Embryonic ecdysteroids of Drosophila melanogaster. J. Insect Physiol. 34 (7), 633–637. 10.1016/0022-1910(88)90071-6. Ecdysone: From Biosynthesis to Mode of Action. [DOI] [Google Scholar]
  30. Matsushima Daijiro, Kasahara Ryota, Matsuno Kumiko, Aoki Fugaku, Suzuki Masataka G., 2019. Involvement of ecdysone signaling in the expression of the doublesex gene during embryonic development in the silkworm, Bombyx mori. Sex. Dev 13 (3), 151–163. 10.1159/000502361. [DOI] [PubMed] [Google Scholar]
  31. Mito Taro, Noji Sumihare, 2008. The two-spotted cricket Gryllus bimaculatus: an emerging model for developmental and regeneration studies. Cold Spring Harb. Protoc. 2008 (12), emo110. 10.1101/pdb.emo110. [DOI] [PubMed] [Google Scholar]
  32. Ohno Carolyn K., Martin Petkovich, 1993. FTZ-F1β, a novel member of the Drosophila nuclear receptor family. Mech. Dev. 40 (1), 13–24. 10.1016/0925-4773(93)90084-B. [DOI] [PubMed] [Google Scholar]
  33. Peel Andrew D., Chipman Ariel D., Akam Michael, 2005. Arthropod segmentation: beyond the Drosophila paradigm. Nat. Rev. Genet. 6 (12), 905–916. 10.1038/nrg1724. [DOI] [PubMed] [Google Scholar]
  34. Pfaffl Michael W., 2001. A new mathematical model for relative quantification in real-time RT–PCR. Nucleic Acids Res. 29 (9), e45. 10.1093/nar/29.9.e45. [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Maria-Dolors Piulachs, Pagone Viviana, Belles Xavier, 2010. Key roles of the broad-complex gene in insect embryogenesis. Insect Biochem. Mol. Biol. 40 (6), 468–475. 10.1016/j.ibmb.2010.04.006. [DOI] [PubMed] [Google Scholar]
  36. Reding Katie, Chen Mengyao, Lu Yong, Alys M, Jarvela Cheatle, Pick Leslie, 2019. Shifting roles of Drosophila pair-rule gene orthologs: segmental expression and function in the milkweed bug Oncopeltus fasciatus. Development 146 (17), dev181453. 10.1242/dev.181453. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Riddiford Lynn, 1993. Hormones and Drosophila Development. January. [Google Scholar]
  38. Anne-Françoise Ruaud, Lam Geanette, Thummel Carl S., 2010. The Drosophila nuclear receptors DHR3 and ВFTZ-F1 control overlapping developmental responses in late embryos. Development (Cambridge, England) 137 (1), 123–131. 10.1242/dev.042036. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Schindelin Johannes, Arganda-Carreras Ignacio, Frise Erwin, Kaynig Verena, Longair Mark, Tobias Pietzsch, Preibisch Stephan, et al. , 2012. Fiji: an open-source platform for biological-image analysis. Nat. Methods 9 (7), 676–682. 10.1038/nmeth.2019. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Taylor Shannon E., Dearden Peter K., 2022. The Nasonia pair-rule gene regulatory network retains its function over 300 million years of evolution. Development 149 (5), dev199632. 10.1242/dev.199632. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Woodard Craig T., Baehrecke Eric H., Thummel Carl S., 1994. A molecular mechanism for the stage specificity of the Drosophila prepupal genetic response to ecdysone. Cell 79 (4), 607–615. 10.1016/0092-8674(94)90546-0. [DOI] [PubMed] [Google Scholar]
  42. Xiang Jie, Katie Reding, Heffer Alison, Pick Leslie, 2017. Conservation and variation in pair-rule gene expression and function in the intermediate-germ beetle Dermestes maculatus. Development 144 (24), 4625–4636. 10.1242/dev.154039. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Yu Yan, Li Willis, Su Kai, Yussa Miyuki, Han Wei, Perrimon Norbert, Pick Leslie, 1997. The nuclear hormone receptor ftz-F1 is a cofactor for the Drosophila homeodomain protein ftz. Nature 385 (6616), 552–555. 10.1038/385552a0. [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

MMC2
NIHMS1894900-supplement-MMC2.docx (9.8MB, docx)
MMC1
NIHMS1894900-supplement-MMC1.xlsx (7.2KB, xlsx)

Data Availability Statement

Data will be made available on request.

RESOURCES