Similar biological phenomena can result from different processes occurring in different organisms. For example, the early stages of how an insect develops from an egg can vary substantially between different species. Nonetheless, all insects have a body plan that develops in segments. The same outcome occurring as a result of different developmental steps is known as ‘system drift’, but the mechanisms underlying this phenomenon are largely unknown.

How the body segments of the fruit fly Drosophila develop has been extensively studied. First, a female fruit fly adds messenger RNA (or mRNA) molecules copied from a number of genes into her egg cells. These mRNA molecules are then used to produce proteins whose concentration varies along the length of each egg. These proteins in turn switch on so-called ‘gap genes’ in differing amounts in different locations throughout the fruit fly embryo. The activity of these genes goes on to define the position and extent of specific segments along the fruit fly's body.

Like the fruit fly, the scuttle fly Megaselia abdita has a segmented body. However, mothers of this species deposit somewhat different protein gradients into their eggs. How the regulation of development differs in the scuttle fly to compensate for this change is unknown. Now, Wotton et al. have studied, in detail, how gap genes are regulated in this less well-understood fly species to understand the mechanisms responsible for a specific example of system drift.

In the fruit fly, gap genes normally switch-off (or reduce the expression of) other gap genes within the same developing body segment, and Wotton et al. found that the same kind of interactions tended to occur in the scuttle fly. As such, the overall structure of the gap gene network was fairly similar between scuttle and fruit flies. There were, however, differences in the strength of these interactions in the two fly species. These quantitative differences result in a different way of making the same segmental pattern in the embryo. In this way, Wotton et al. show how tinkering with the strength of specific gene interactions can provide an explanation for system drift.

An important question for evolutionary biology is how developmental processes can compensate for variable environmental, signalling, or regulatory inputs to create a constant phenotypic outcome (Waddington, 1942). Segment determination during early insect embryogenesis is a well-studied example of this phenomenon. The segmentation gene network produces very robust and conserved output patterns despite fast-evolving upstream inputs through maternal gradients and vastly different modes of segmentation dynamics in different insect taxa (Sander, 1976; Davis and Patel, 2002). This type of neutral network evolution—producing the same output based on different regulatory principles—is called developmental system drift or phenogenetic drift (Weiss and Fullerton, 2000; True and Haag, 2001; Weiss, 2005; Haag, 2007; Pavlicev and Wagner, 2012). It is believed to be a very widespread phenomenon and can be interpreted as phenotypically neutral evolution along so-called genotype (meta-)networks. Genotype networks consist of different regulatory network structures—connected by simple mutational steps—that produce the same patterning or phenotypic output (Ciliberti et al., 2007a, 2007b; Wagner and Lynch, 2008; Draghi et al., 2010; Wagner, 2011).

In order to discover the mechanisms underlying developmental system drift, it is necessary to systematically and quantitatively investigate the structure and dynamics of evolving regulatory networks. In this study, we use the dipteran gap gene system—constituting the first regulatory layer of the segmentation gene network (Foe and Alberts, 1983; Akam, 1987; Ingham, 1988; Jaeger, 2011)—as a model system to study developmental system drift.

The gap gene system in Drosophila melanogaster (family: Drosophilidae) is one of the most thoroughly studied developmental gene regulatory networks today. The genetic and molecular mechanisms of gap gene regulation have been investigated extensively over the last few decades (reviewed in Jaeger (2011)), and a number of mathematical models exist that faithfully reproduce gap gene expression dynamics in this species (Reinitz et al., 1995; Jaeger et al., 2004a, 2004b; Perkins et al., 2006; Ashyraliyev et al., 2009; Manu et al., 2009a, 2009b; Crombach et al., 2012a, 2012b; Becker et al., 2013). In this study, we provide a brief summary of the most important regulatory principles that were revealed by this research.

An overview of the structure of the gap gene network is given in Figure 1 (grey inset). Gap genes receive regulatory inputs from maternal protein gradients formed by the gene products of bicoid (bcd), hunchback (hb), and caudal (cad) (Figure 1, top row of graphs) (reviewed in St Johnston and Nüsslein-Volhard (1992)). These gradients set up an initial asymmetry along the major or antero-posterior (A–P) axis of the embryo. Bcd and Cad activate the four trunk gap genes hb, Krüppel (Kr), knirps (kni), and giant (gt), which become expressed in broad, overlapping domains (Figure 1, bottom row of graphs). As development proceeds, gap gene cross-repression refines the pattern (Figure 1, grey inset) (see Jaeger, 2011 for review). Domains of hb and kni as well as Kr and gt have mutually exclusive expression patterns and show strong mutual repression. This ‘alternating cushions’ mechanism maintains and sharpens the basic staggered arrangement of gap domains. The expression patterns of Kr/kni, kni/gt, and gt/hb overlap and show weaker repression with a posterior-to-anterior bias, which results in anterior shifts of each of these domains over time. Finally, trunk gap gene expression is repressed in the posterior pole region of the embryo by the terminal gap genes tailless (tll) and huckebein (hkb).

Figure 1

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Our understanding of the dipteran gap gene system outside the drosophilid family is much less well developed. This limitation applies in particular to nematoceran midges and mosquitoes (Figure 1, black phylogenetic branches on the right) (Jiménez-Guri et al., 2013). Qualitative (Rohr et al., 1999; García-Solache et al., 2010; Jiménez-Guri et al., 2014) and quantitative (Crombach et al., 2014; Janssens et al., 2014) studies of segmentation gene expression in the moth midge Clogmia albipunctata (family: Psychodidae) indicate that posterior patterning—especially the onset of posterior hb expression—is delayed and that repression between Kr and gt may not be conserved in this species, which lacks a posterior gt domain. A similar posterior delay is observed in the malaria mosquito Anopheles gambiae (Culicidae), where both posterior domains of hb and gt are present, but appear only after gastrulation in an A–P order which is reversed compared to that observed in D. melanogaster (Goltsev et al., 2004). Unfortunately, several factors involved in gap gene regulation are still unknown in these nematoceran species, and the lack of functional evidence based on genetic perturbations limits the interpretation of experimental results and verification of network models.

The situation is slightly better within the Cyclorrhapha, a sub-taxon of brachyceran or ‘higher’ flies (Figure 1, red phylogenetic branches on the left) (Wiegmann et al., 2011; Jiménez-Guri et al., 2013). Apart from D. melanogaster, two other cyclorrhaphan species are emerging as experimentally tractable models. One of these is the marmalade hoverfly Episyrphus balteatus (Syrphidae). In this species, the pattern and order of gap gene activation along the A–P axis is largely conserved compared to D. melanogaster, even though there is no maternal contribution to hb expression (Figure 1) (Lemke and Schmidt-Ott, 2009; Lemke et al., 2010). However, the precise dynamics of gap gene expression and the nature of gap gene cross-regulation have not yet been investigated in this species.

The second emerging non-drosophilid model system is Megaselia abdita, a member of the early-branching cyclorrhaphan family Phoridae (Figure 1) (Wiegmann et al., 2011; Rafiqi et al., 2011a; Jiménez-Guri et al., 2013). In what follows, we will focus on this species. Gap gene expression in M. abdita is less well studied than in E. balteatus (Figure 1) and little is known about gap gene regulation. Maternal transcripts of bcd and hb can be detected (Stauber et al., 1999, 2000; Lemke et al., 2008), but no maternal contribution to cad expression is present in this species (Stauber et al., 2008). Localised maternal bcd transcripts at the anterior pole extend further posterior, and bcd RNAi cuticle phenotypes affect more posterior segments than in D. melanogaster, indicating a broader role of Bcd in M. abdita (Stauber et al., 1999, 2000). In contrast, expression of maternal and zygotic hb appears very similar in both species (Stauber et al., 2000; Lemke et al., 2008). The zygotic anterior hb domain is severely depleted in embryos treated with bcd RNAi, suggesting activation of hb by Bcd as in D. melanogaster (Lemke et al., 2008). Moreover, the regulatory role of hb seems to be similar as well, since cuticle phenotypes of hb RNAi-treated M. abdita embryos resemble hb hypomorphs in D. melanogaster (Stauber et al., 2000). Finally, Kr expression has been mentioned to be similar in both species (Rafiqi et al., 2008), although no data were shown since the study focused on extraembryonic tissues at later stages of development.

The similarity of gap gene expression across cyclorrhaphan species—despite considerable variation in maternal gradients—provides an excellent opportunity to study developmental system drift in an experimentally tractable set of laboratory models. A comparative, network-level comparison of gap gene regulation would reveal how the network compensates for variable upstream inputs and inter-species differences in initial expression patterns. As a foundation for such an analysis we have previously sequenced the early embryonic transcriptome of M. abdita (Jiménez-Guri et al., 2013), and have established a precise staging scheme for embryogenesis—with a particular focus on the blastoderm stage (Wotton et al., 2014).

In this paper, we present a comprehensive analysis of gap gene expression in M. abdita, based on quantitative data with high spatial and temporal resolution. We compare wild-type expression dynamics to an equivalent dataset from D. melanogaster (Crombach et al., 2012a). Our comparison reveals that gap gene expression differs significantly between the two species during early to mid blastoderm stage but converges to equivalent patterns before the onset of gastrulation. In particular, we find that the broader influence of Bcd in M. abdita causes gap domains to emerge at more posterior positions compared to D. melanogaster. In addition, the absence of maternal Cad causes a delay in the onset of anterior shifts of posterior gap domains in M. abdita. This delay is later compensated by stronger shifts compared to D. melanogaster. To investigate the regulatory changes underlying these altered dynamics, we use previously established RNAi protocols (Stauber et al., 2000; Lemke et al., 2008) to systematically knock-down trunk and terminal gap genes. We then assay expression of the remaining trunk gap genes in each genetically perturbed background using the same high-resolution quantitative approach as for wild-type patterns. Our knock-down analysis reveals that qualitative regulatory structure—the type of interactions between gap genes—is strongly conserved between species. We do, however, detect changes significantly affecting the strength of gap gene cross-repression. Based on this, we suggest quantitative changes in regulatory mechanisms for the altered dynamics of gap gene expression in M. abdita. Our evidence suggests that such quantitative developmental system drift is a common feature of evolving developmental systems.

M. abdita fly culture and embryo collection/fixation were carried out as described in (Rafiqi, at al. 2011b; Rafiqi et al., 2011c). Blastoderm stage embryos of M. abdita—corresponding to embryonic stages 4 and 5 (Campos-Ortega and Hartenstein, 1997; Wotton et al., 2014)—were collected 4 hr (hrs) after egg laying. We visualised maternal-coordinate and gap gene mRNA expression patterns using an enzymatic (colorimetric) in situ hybridisation protocol as previously described (Crombach et al., 2012a). We used single probes for the majority of stains. Embryos double-stained for gap genes or for a gap gene along with the pair-rule gene even-skipped (eve) were used to confirm the position of expression domains relative to each other (see http://superfly.crg.eu for our full dataset; Cicin-Sain et al., 2015). We took four images of each stained embryo (Crombach et al., 2012b): a bright-field image from which the expression profile was extracted, a DIC image used to create the whole-embryo mask, and two images of the fluorescently counterstained nuclei and membrane morphology respectively, which were used to determine the time class of each embryo based on the staging scheme described in Wotton et al. (2014). These images were processed to extract the position of expression domain boundaries as described in Crombach et al. (2012b) and shown in Figure 2A,B.

RNAi treatment was carried out as described in Stauber et al. (2000); Lemke et al. (2008); Rafiqi et al. (2011a). Two distinct dsRNA constructs were injected per gene, one being the knock-down agent, the other one—with conserved domains excluded—serving as a control. Full length bcd (CAB40892.1), cad (ABY25304.1), hb (CAC00483.1), Kr (ABX89307.1), and eve (AAT92572.1) open reading frames (ORFs) were amplified from cDNA using primers based on published sequences (Stauber et al., 1999, 2000, 2008; Bullock et al., 2004; Rafiqi et al., 2008). tll, (KM489059), gt (KM489058), and kni (KM489060) (ORFs) were cloned from cDNA using degenerate and RACE PCR. hkb (KM489061) was cloned using sequence data from our published early embryonic transcriptome (http://diptex.crg.es; gene ID: comp8024) (Jiménez-Guri et al., 2013). cad germ-line clone females were generated as described in Wu and Lengyel, (1998) using fly stocks cad(2L-264-12-3)FRT40A/CyO,hs-hid, w*; P{w⁺ ovo^D1} P{neo FRT}40A/T(1,2)OR64/SM6a, and y*w*,P{hs-Flp}122; If/CyO,hs-hid (kind gift from Uwe Irion). These females were mated to wild-type males to obtain embryos, all of which lack maternal cad activity but carry one zygotically active wild-type cad gene.

Appendix I: M. abdita segmentation gene expression

In this section, we describe the quantified wild-type mRNA expression patterns of M. abdita maternal co-ordinate and gap genes in detail and compare them to the published literature in D. melanogaster.

Maternal co-ordinate genes

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bicoid (bcd)

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Our data confirm earlier reports that bcd transcripts are distributed more broadly in M. abdita than in D. melanogaster (Stauber et al., 2002, 2000, 1999). At C12, bcd transcripts extend from the anterior pole to 23% A–P position (Figure 10). This is 9–12%EL further to the posterior, or almost twice a far, as in D. melanogaster embryos at any time during the blastoderm stage (Berleth et al., 1988; Little et al., 2011; Crombach et al., 2012b). At subsequent stages, the distribution of bcd transcripts becomes more restricted and its posterior boundary moves to 16% A–P position at C13 and to 12% at C14A-T1/T2 (Figure 10). In comparison, we do not detect transcripts in D. melanogaster beyond 11% A–P position during C14A, in rough agreement with previous studies that reported a posterior boundary at around 14% (Berleth et al., 1988; Little et al., 2011). bcd transcripts in M. abdita disappear at C14A-T2, earlier than in D. melanogaster where they can still be detected up to C14A-T4 (Figure 10) (Crombach et al., 2012a).

Figure 10

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caudal (cad)

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Unlike D. melanogaster, M. abdita does not show any maternal cad expression (Stauber et al., 2008). We first detect zygotic cad transcripts at C12 extending from around 34% A–P position to the posterior pole (Figure 10). At C13, the anterior boundary of cad expression is at a similar position in M. abdita and D. melanogaster (31 vs 33% A–P position; Figure 10) (Crombach et al., 2012a). However, at subsequent stages, cad retracts towards the posterior in D. melanogaster (39–42% A–P position during C14A-T2 to T5), while it maintains a stationary boundary at around 31–32% A–P position in M. abdita (until around C14A-T5; Figure 10; see also Stauber et al., 2008). At C14A-T5, cad transcripts start to disappear in most of the central region of the embryo in both species (Figure 10) (Mlodzik and Gehring, 1987; Schulz and Tautz, 1995). Around the same time, cad also retracts from the posterior pole forming a posterior stripe at 79–90% A–P position in M. abdita (Figure 10) (Stauber et al., 2008). This domain is much wider than its equivalent in D. melanogaster (about 11 vs 4% EL) (Mlodzik and Gehring, 1987; Moreno and Morata, 1999; Crombach et al., 2012a). It remains larger despite the posterior boundary shifting to the anterior at C14A-T8 causing its width to shrink to about 7% (vs. 4% EL in D. melanogaster; Figure 10) (Crombach et al., 2012a).

Terminal gap genes

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tailless (tll)

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We first detect tll expression at C12 in a domain at the posterior pole extending to 90% A–P position (Figure 10). This boundary remains more or less stationary up to C14A-T6, after which posterior tll expression appears to decrease. Despite beginning as a slightly larger domain at C13 than its equivalent in D. melanogaster, the posterior tll domain narrows and comes to occupy a slightly smaller region of the embryo (boundary at 84 vs 89% A–P position; Figure 10) (Pignoni et al., 1990; Crombach et al., 2012a). Expression dynamics in the anterior differ more markedly: while the anterior tll domain in D. melanogaster appears in C13, its equivalent in M. abdita does not appear until around C14-T4 (Figure 10) (Pignoni et al., 1990; Crombach et al., 2012a). At later stages, this domain retracts from the anterior pole and expands slightly to the posterior in both species.

huckebein (hkb)

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We first detect hkb expression at C12 in two domains, one at the anterior pole extending to 8% A–P position, and one at the posterior pole extending to 90% A–P position (Figure 10). The boundary of the anterior domain remains at 7–8% A–P position through C13 and most of C14A, narrowing to 6% at C14A-T8. The extent of the posterior domain reaches up to around 89–90% A–P position between C14A-T1 and T6. In contrast to D. melanogaster, this domain narrows slightly during C14A-T7 and T8 to 92% A–P position. Other than that, expression dynamics and boundary positions are very similar to those reported for D. melanogaster (Figure 10) (Brönner and Jäckle, 1996; Crombach et al., 2012a).

Trunk gap genes

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hunchback (hb)

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Maternal hb transcripts have been reported to be uniformly distributed in the pre-blastoderm embryo of M. abdita (Stauber et al., 2000). We have not examined this maternal pattern any further. Earlier qualitative studies reported that zygotic expression of hb is similar in M. abdita and D. melanogaster (Stauber et al., 2000; Lemke et al., 2008). In contrast to these reports, we detect significant differences in expression dynamics between the two species. We first detect the formation of an anterior domain of zygotic hb expression in M. abdita at C11. This domain extends from 0 to 56% A–P position, reaching significantly further posterior than in D. melanogaster, where it is located from 0 to 50% A–P position (Figure 11) (Tautz, 1988; Crauk and Dostatni, 2005). In M. abdita, the posterior boundary of the anterior hb domain steadily shifts to the anterior over time (Figure 11). This shift covers 8% EL by C14A-T8. In contrast, the equivalent boundary in D. melanogaster remains stationary through C14A (see Figure 3) (Surkova et al., 2008a; Crombach et al., 2012a). By C13, some M. abdita embryos show a decrease of expression levels at the anterior pole (Figure 11). This process continues through C14A and results in the formation of a broad anterior stripe clearly visible at C14A-T4 (Figure 11). A slightly more intense stripe within the broad anterior hb domain in M. abdita may correspond to the parasegment 4 (PS4) stripe in D. melanogaster (Tautz et al., 1987; Crombach et al., 2012a).

Figure 11

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Around C12 or C13, a small posterior hb domain appears in M. abdita, extending to 90% A–P position as in D. melanogaster (Figure 11) (Tautz et al., 1987; Crombach et al., 2012a). By C14A-T2, the anterior expression border has shifted anteriorly to 84% A–P position, leading to an expansion of the posterior hb domain. In D. melanogaster, this domain retracts from the posterior pole around C14A-T2 (Crombach et al., 2012a). This retraction is delayed in M. abdita. Expression levels of hb start to drop in the posterior pole region around C14A-T6 but full retraction only occurs at C14A-T8 (Figure 11). At this stage, the posterior hb domain of M. abdita is positioned slightly further anterior (72–84 vs 75–85% A–P position), and is somewhat wider (12 vs 10%EL), than in D. melanogaster (Crombach et al., 2012a). This difference in domain width can be explained by the increased shift of the anterior boundary of posterior hb in M. abdita compared to D. melanogaster (18 vs 14% EL overall; see also Figure 3).

giant (gt)

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We first detect gt expression in M. abdita at C12 in an anterior stripe at 14–34% A–P position, and a posterior domain extending from 73% A–P position to the posterior pole (Figure 11). In D. melanogaster, gt expression is initiated at a similar time, during cycles C11 and C12 (Jaeger et al., 2007). During C13, the anterior domain is located more anteriorly in M. abdita than in D. melanogaster (15–35 vs 19–40% A–P position) but is of comparable width (20 vs 21%EL) (Jaeger et al., 2007; Crombach et al., 2012a). In D. melanogaster, this domain starts to split around C14A-T3 and forms two clearly resolved stripes at C14A-T4, and an additional expression domain (called the head patch) appears at the anterior pole at C14A-T5 (Mohler et al., 1989; Kraut and Levine, 1991a; Crombach et al., 2012a; Becker et al., 2013). These three anterior expression stripes persist until C14A-T8, and their posterior-most boundary shifts by about 3%EL to 37% A–P position at the onset of gastrulation (Crombach et al., 2012a; Becker et al., 2013). In M. abdita, gt expression starts to decrease in the anterior part of the anterior domain around C14A-T4 forming a single narrow stripe at C14A-T5 (Figure 11). At the same time, a domain comparable to the head patch of D. melanogaster appears at the anterior pole (Figure 11). In contrast to D. melanogaster, the posterior boundary of anterior gt expression does not shift significantly in M. abdita (see also Figure 3).

At C13, the anterior boundary of the posterior gt domain is at a slightly more anterior position in M. abdita than in D. melanogaster (72 vs 74% A–P position) (Crombach et al., 2012a). It also shifts further to the anterior by C14A-T8 in this species (10 vs 9% EL to 62 vs 65% A–P position). Furthermore, retraction from the posterior pole occurs slightly earlier in M. abdita than in D. melanogaster. Some M. abdita embryos (2 out of 13) show the first signs of retraction at C13, and by C14A-T1, the posterior boundary of posterior gt is found at 86% A–P position (Figure 11). In D. melanogaster, this domain only retracts at C14A-T1 with the posterior boundary located at 82% A–P position (Crombach et al., 2012a; Becker et al., 2013). Although the posterior gt domain of M. abdita remains wider for most of C14A, it narrows in both species to a width of about 8% EL by C14A-T8 (Figure 11). This is caused by a very rapid and strong shift of its posterior boundary during late C14A in M. abdita, which results in a larger overall anterior shift (18 vs 9% EL). During this process, the M. abdita boundary ‘overtakes’ its equivalent in D. melanogaster such that it is found further anterior (71 vs 73% A–P position) at C14A-T8 (Figure 11; see also Figure 3) (Crombach et al., 2012a; Becker et al., 2013).

krüppel (Kr)

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In M. abdita, we first detect Kr expression at C12, as is the case in D. melanogaster (Knipple et al., 1985; Jaeger et al., 2007). Kr appears in a central domain between 40 and 63% A–P position (Figure 11). This domain is much wider at this stage than in D. melanogaster (23 vs 12%EL), extending further to the anterior (by 8.6% EL) and to the posterior (by 2.6% EL) (Knipple et al., 1985; Jaeger et al., 2007). The posterior boundary of the central Kr domain shifts further in M. abdita than in D. melanogaster (12 vs 8% EL). In contrast, the anterior boundary shifts somewhat less (3% vs 6% EL). This results in a narrowing of the domain over time in both species (Figure 11). At all times, however, the central Kr domain remains wider (14 vs 10% EL at C14A-T8), and its final position at the onset of gastrulation lies further anterior (37–51 vs 43–53% A–P position), in M. abdita than in D. melanogaster (see also Figure 3) (Crombach et al., 2012a; Becker et al., 2013).

In addition to its central domain, Kr shows two additional domains during the blastoderm stage (Knipple et al., 1985; Harding and Levine, 1988). Neither of these are involved in gap gene cross-regulation. The head domain appears earlier in M. abdita than in D. melanogaster (C12 vs C14A-T4/T5), while the posterior domain appears slightly later (C14A-T8 vs T6; Figure 11) (Crombach et al., 2012a; Becker et al., 2013). These domains exhibit similar expression dynamics in the two species, although both extend somewhat further into the central region of the embryo in D. melanogaster than in M. abdita (Figure 11).

Knirps (kni)

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The expression of kni in M. abdita is very similar to D. melanogaster (see Figure 3). We first detect kni expression at C12 in the anterior pole region extending to 7% A–P position and in an abdominal domain at 61–78% A–P position (Figure 11). While some studies have reported kni expression in D. melanogaster at C12 (Rothe et al., 1989, 1994), more recent quantitative measurements could not detect kni before the mitotic division preceding the interphase of C13 (Jaeger et al., 2007). Throughout C13 and C14A, the abdominal kni domain shifts to the anterior and narrows progressively (Figure 11). The extent of the domain shift during C14A is slightly greater in M. abdita than in D. melanogaster: 4 vs 2% EL for the anterior and 8 vs 7% EL for the posterior boundary (Crombach et al., 2012a; Becker et al., 2013). The abdominal kni domain is also consistently wider and narrows more in M. abdita than in D. melanogaster.

In the anterior, expression at the pole remains at around 8–10% A–P position as in D. melanogaster (Crombach et al., 2012a). A head stripe domain appears around C14A-T3 or T4 at 24–27% A–P position (Figure 11). This is slightly further to the anterior than the equivalent head stripe in D. melanogaster (located at 27–30% A–P position), which appears at C14A-T4 (Crombach et al., 2012a). Expression in the head stripe is maintained up to C14A-T8.

Appendix II: trunk gap gene RNAi cuticle phenotypes

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In this section, we describe the gap gene RNAi cuticles of M. abdita and compare them to the published literature on gap gene mutants in D. melanogaster.

In M. abdita, hb knock-down (as previously published in Stauber et al., 2000) results in the deletion of the thorax, with additional defects in abdominal segments A1–8 and A8, and a reduced cephalopharyngeal skeleton, suggesting that the posterior gnathocephalon is also missing. This corresponds to a hypomorphic hb mutant phenotype in D. melanogaster (Nüsslein-Volhard and Wieschaus, 1980; Jürgens et al., 1984; Lehmann and Nüsslein-Volhard, 1987).

Other trunk gap gene knock-downs resemble D. melanogaster null mutants (Figure 12). M. abdita gt RNAi resulted in cuticles lacking abdominal segments A5–7 (Figure 12B) rather than just the denticle bands of abdominal segments A5–7 as in D. melanogaster (i.e., no naked cuticle is formed) (Wieschaus et al., 1984a; Gergen and Wieschaus, 1986; Petschek et al., 1987). Additionally, head segments are affected, and A8 is slightly reduced with filzkörper not fully present.

Figure 12

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Following kni RNAi in M. abdita embryos, abdominal segments A1–7 merge (Figure 12C). In the most severe knock-down phenotypes, the fused segments are marked by a single, irregular denticle field. Intermediate phenotypes may exhibit a few remaining discernible abdominal denticle belts. In some kni RNAi cuticles, more posterior markers as well as the posterior portion of the T3 denticle belt were distorted as well. Head segments appear unaffected. This severe kni RNAi phenotype is very similar to the null mutant kni phenotype in D. melanogaster (Nüsslein-Volhard and Wieschaus, 1980; Jürgens et al., 1984; Nauber et al., 1988).

In severely affected M. abdita Kr knock-down cuticles, only the telson and A8, plus one or two additional abdominal segments are formed (putative A7, and sometimes A6; Figure 12D). Other abdominal segments and all thoracic segments are either missing or fused, and head involution fails to occur. Head segments were strongly affected. Unlike in D. melanogaster, where A6 is duplicated as a mirror image in Kr mutants (Nüsslein-Volhard and Wieschaus, 1980; Wieschaus et al., 1984b), we cannot identify clear instances of local inversions or duplications in M. abdita. However, it is possible that we have missed these phenotypes, due to the less obvious polarity of the segmental denticle patterns in M. abdita compared to D. melanogaster.

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Author details

1. Karl R Wotton

   1. European Molecular Biology Laboratory, CRG Systems Biology Research Unit, Centre for Genomic Regulation, Barcelona, Spain
   2. Universitat Pompeu Fabra, Barcelona, Spain

   Contribution

   KRW, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   For correspondence

   karl.wotton@crg.eu

   Competing interests

   The authors declare that no competing interests exist.

   "This ORCID iD identifies the author of this article:" 0000-0002-8672-9948

2. Eva Jiménez-Guri

   1. European Molecular Biology Laboratory, CRG Systems Biology Research Unit, Centre for Genomic Regulation, Barcelona, Spain
   2. Universitat Pompeu Fabra, Barcelona, Spain

   Contribution

   EJ-G, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

   "This ORCID iD identifies the author of this article:" 0000-0002-9592-1077

3. Anton Crombach

   1. European Molecular Biology Laboratory, CRG Systems Biology Research Unit, Centre for Genomic Regulation, Barcelona, Spain
   2. Universitat Pompeu Fabra, Barcelona, Spain

   Contribution

   AC, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

4. Hilde Janssens

   1. European Molecular Biology Laboratory, CRG Systems Biology Research Unit, Centre for Genomic Regulation, Barcelona, Spain
   2. Universitat Pompeu Fabra, Barcelona, Spain

   Contribution

   HJ, Acquisition of data, Analysis and interpretation of data

   Competing interests

   The authors declare that no competing interests exist.

5. Anna Alcaine-Colet

   1. European Molecular Biology Laboratory, CRG Systems Biology Research Unit, Centre for Genomic Regulation, Barcelona, Spain
   2. Universitat Pompeu Fabra, Barcelona, Spain
   3. Universitat de Barcelona, Barcelona, Spain

   Contribution

   AA-C, Analysis and interpretation of data

   Competing interests

   The authors declare that no competing interests exist.

6. Steffen Lemke

   Department of Organismal Biology and Anatomy, University of Chicago, Chicago, United States

   Present address

   Centre for Organismal Studies, Ruprecht Karls University, Heidelberg, Germany

   Contribution

   SL, Analysis and interpretation of data, Contributed unpublished essential data or reagents

   Competing interests

   The authors declare that no competing interests exist.

7. Urs Schmidt-Ott

   Department of Organismal Biology and Anatomy, University of Chicago, Chicago, United States

   Contribution

   US-O, Analysis and interpretation of data, Contributed unpublished essential data or reagents

   Competing interests

   The authors declare that no competing interests exist.

8. Johannes Jaeger

   1. European Molecular Biology Laboratory, CRG Systems Biology Research Unit, Centre for Genomic Regulation, Barcelona, Spain
   2. Universitat Pompeu Fabra, Barcelona, Spain

   Contribution

   JJ, Conception and design, Analysis and interpretation of data, Drafting or revising the article

   For correspondence

   yogi.jaeger@crg.eu

   Competing interests

   The authors declare that no competing interests exist.

   "This ORCID iD identifies the author of this article:" 0000-0002-2568-2103

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