Results and discussion

Insect wings vary dramatically in their size and morphology. Butterflies and moths in particular have attracted attention for their rapidly evolving wings that show extreme variation in shape and color pattern. Interestingly, the diversity of lepidopteran color patterns appears to follow relatively simple rules, wherein an evolutionarily conserved set spot and stripe pattern elements can vary in size, color, and presence/absence along the antero-posterior (AP) axis.^1,2 Surveys of wing pattern diversity across butterflies, considering both natural variation and genetic mutants, suggest that wings can be subdivided into five AP domains, bounded by the M1, M3, Cu2, and 2A veins as summarized in Figure 1A.³ Within each domain there is a strong correlation between pattern variation and wing morphology, and, conversely, between domains there is a relative degree of independence in morphological variation. We know little about how these different AP domains are differentiated during wing development, however.

Figure 1.

Some aspects of the AP wing differentiation process have been extrapolated from D. melanogaster wings to butterflies. The transcription factors engrailed and spalt define AP domains in developing fly wings^4-6 and exhibit expression patterns in butterflies that suggest some functions may be conserved. In butterflies engrailed transcription across the entire region posterior to the M1 vein, while its paralog invected marks a largely overlapping region between M1 and 2A.^7-9 spalt expression is more dynamic and complex in butterflies, but appears to consistently mark a region between the R2 and M3 veins.⁹ These observations, coupled with the overall size and morphological complexity of butterfly wings, have spurred interest in the question of whether additional AP differentiation mechanisms may underlie AP domain specification in Lepidoptera and other insects.^10,11 The identification of additional domain-determining transcription factors in broad winged insects would provide a model for the modular diversification of insect wing morphology.

The goal of this study was to identify the molecular basis of AP domain specification in the butterfly wing, posterior of the M1 engrailed boundary. We initially identified the transcription factor mirror as a potential candidate selector gene based on previous mRNA-seq studies,¹² coupled with D. melanogaster work showing that mirror specifies the alula – a small membranous lobe at the posterior base of the wing in some dipterans (Figure 1C,D).^13,14 We used in situ hybridization to visualize mirror mRNA localization in the last-instar imaginal discs of the common buckeye butterfly, Junonia coenia, and observed mirror expression throughout the wing domain posterior to the 2A vein boundary (Figures 2A and S1). This expression precisely marks the vannus, or anal region (Figures 1B, E), of the butterfly wing – a distinct posterior domain of the wing blade populated by the anal (A) veins and bordered anteriorly by the claval fold between the cubital (Cu) and A veins in the hindwing (Figure 2B). As observed in various representative species across different butterfly families, this region of the hindwing is distinct from the rest of the wing; the color patterns defining the wing blade terminate at the 2A vein, and in many species it is marked by scales that differ in morphology and lack color patterns found in anterior regions of the wing (Figures 1B and S2A). This region has been proposed to represent the posterior-most color pattern domain in butterflies (Figure 1A, Figure 2B).³

Figure 2.

To functionally assess the role of mirror in wing development we generated CRISPR/Cas9 mosaic knockouts (mKOs) targeting the coding region of J. coenia mirror. We recovered many mKO mutants that consistently exhibited several distinctive phenotypes (Table S1), but only ever in the mirror-expressing posterior vannus region:

1. Venation: In mKO mutant hindwings we observed frequent truncation or complete loss of the loss of the 3A vein, as well as discontinuity and ectopic bifurcation of 2A veins (hindwings: Figures 2C-G, S2, S3; forewings: Figure S4).

2. Wing shape: mKOs consistently showed shape change along the posterior edge of the wing, especially the hindwing where the projecting lobe characteristic of the vannus was reduced and replaced by a more even, regular wing margin. The anal fold between the 2A and 3A veins, which marks the edge of the vannal lobe, was also lost in mutant clones affecting this wing region (Figures 2C-G, S2-S3).

3. Color pattern expansion: In mutant hindwings we consistently observed extension of wing margin color patterns across the 2A vein boundary into the posterior region of the wing. In wild-type wings the submarginal and central symmetry system color patterns terminate at the 2A vein (dorsal: Figure 2C-G; ventral: Figure S3). Remarkably, in mirror mKO mutants these entire color pattern systems extend past the 2A vein into the posterior regions of the wing on both dorsal and ventral wing surfaces. Specifically, submarginal bands continue past 2A to follow the posterior wing margin (Figures 2C-G, S2), while central symmetry patterns extend from 2A down to the posterior margin (Figure S3).

4. Scale morphology: In most mutants vannus scales lost their distinctive silver structural color and rounded apical edges, and transformed into brown, serrated scales typical of the remigium (Figure S2A). We also observed that scales along mutant wing margins often displayed elongated scales, often manifesting as a dense hair-like matt across the posterior margin (Figure S2A).

The knockout phenotypes we observed illustrate several things about mirror’s function in wing development. First, across all mKO replicates we never observed mutant phenotypes anterior to the 2A vein, which suggests that mirror effects are limited to the far posterior domain in which we observed mirror expression. Second, mirror knockouts appear to affect multiple aspects of wing morphology, including scale shape and coloration, overall wing shape, and wing venation. Finally, the loss of mirror function resulted in transformation, not loss, of the posterior domain. Even in mKO individuals that appeared to have a mutant phenotype across the entire posterior region, the mirror-expressing cells posterior to the 2A vein were not simply lost – the wing field continues well past the 2A vein. Instead, the identities of the post-2A scale cells changed in color and structure. In this respect, the expansion of multiple color pattern systems into the posterior domain of mutants was particularly compelling because the loss of central symmetry system and submarginal patterns in the post-2A vannus region is commonly observed in Lepidoptera (Figure 1B). The ability to reconstitute these complex color pattern elements in the posterior domain by knocking out a single transcription factor leads us to speculate that their patterning information may occur in a latent form that is masked by mirror effects. Together, our expression and knockout data suggest that mirror acts as a selector gene necessary to define the identity of the far posterior wing domain.

Identifying mirror as a vannus-specifying factor is of interest for a few reasons. First, it provides a molecular explanation for how different AP domains of the butterfly wing may be individuated to independently evolve their own shape and color pattern variations. Previous authors have proposed the existence of such individuated domains, and speculated that they may be specified by selector genes.^3,10 Our data provide experimental support for this model, and now motivate us to identify additional factors that may specify other domain boundaries between the M1 and A2 veins.

Next, the role of mirror in specifying the vannus provides an important link between molecular developmental biology and Snodgrass’ classical anatomical designations of the insect wing fields – i.e., the remigium, the vannus, and the jugum (Figure 1E).¹⁵ The remigium and the vannus are dominant features of most insect wings, while the jugum is smaller lobe-like feature that is reduced or lost in many clades. The remigium is the major anterior wing blade that is directly articulated by thoracic motor musculature to power flight, and is typically divided from the vannus by a distinct wing fold (e.g. the “anal fold” in butterflies). In many insect orders the vannus is a prominent fan-like structure, which can be a greatly enlarged gliding surface some hemimetabolous clades such as Orthoptera (crickets and grasshoppers), Dictyoptera (mantids and roaches), Phasmatodea (stick insects), and Plecoptera (stoneflies). Importantly, mirror knockdown in the hemimetaboulous milkweed bug Oncopeltus fasciatus causes developmental defects in the claval and anal furrows in the posterior wing,¹⁶ which leads us to infer that mirror’s role in determining the vannus may be deeply conserved in insects. This prompts us to speculate that the developmental individuation of the posterior domain by mirror facilitated the remarkable diversification of the vannus across the insects.

Finally, the function of mirror in determining the alula in D. melanogaster¹³ suggests that the alula may represent an evolutionarily reduced vannus. The ramifications of this are significant for reconstructing the history of the insect wing, because they suggest that the D. melanogaster wing blade is a lone remigium – only one-third of the archetypal insect wing (Figure 1E). The dipteran jugum appears to have been lost entirely, except perhaps as a membranous haltere cover (calypter) in the Calyptratae clade, as speculated by Snodgrass.¹⁵ Thus, while flies have been an important model for characterizing genes and processes that build insect wings, a more comprehensive understanding of the development and evolution of insect wings requires work in species that have wings more representative of the complete ancestral blueprint.

Materials and methods

Identity of Iroquois family gene mirror ortholog in J. coenia

We performed a reciprocal BLAST of amino acid sequences of D. melanogaster Iroquois Complex genes araucan, caupolican, and mirror to identify their orthologs in the latest J. coenia and Heliconius erato lativitta genome sequences on lepbase.org and Tribolium castaneum and Apis mellifera on NCBI. The amino acid sequences of the top hits – from J. coenia (JC_02265-RA, JC_02269-RA), H. erato lativitta (HEL_009655-RA, HEL_009656-RA), A. mellifera (LOC412840, LOC412839) and T. castaneum (LOC652944, LOC660345) were aligned to the Iroquois genes of D. melanogaster using MUSCLE.¹⁷ We used these alignments to build a maximum likelihood gene phylogeny on IQ-TREE 2¹⁸ using ModelFinder¹⁹ for estimating the most accurate substitution model for our amino acid sequences. We used FigTree to visualize the tree depicted in Figure S5.

HCR fluorescent in situ hybridization of mirror mRNA

This protocol was adapted and modified from Bruce et al. (dx.doi.org/10.17504/protocols.io.bunznvf6). The coding sequence of JC_02269-RA was used by Molecular Instruments, Inc. (Los Angeles, CA, USA) to design a mirror-specific hybridization chain reaction (HCR) probe set (Lot #PRQ901). The probes are unique to the JC_02269-RA transcript as verified by BLAST search using the latest J. coenia genome assembly (v2) on lepbase.org, and do not share significant sequence similarity with any other transcripts, including the other Iroquois family member JC_02265-RA (see Figure S5). Last (5^th) instar larval wing imaginal discs were dissected in cold 1x PBS. The dissected discs collected from left and ride sides of the larvae were randomly assigned to control or treatment batches. Both controls and treatment batches were fixed in 1x cold fix buffer (750uL PBS, 50mM EGTA, 250uL 37% formaldehyde) on ice. The discs were then washed three times in 1x PBS with 0.1% Tween 20 (PTw) on ice spending approximately 30sec – 2min for each wash. Tissues were then gradually dehydrated by washing in 33%, 66%, and 100% MeOH (in PTw) for 2-5min / wash on ice, and stored in 100% MeOH in -20°C for days until HCR protocol was started. On the day of the HCR protocol, the wing discs were progressively rehydrated in cold 75% MeOH, 50% MeOH, and 25% MeOH in PTw spending 2-5min per wash. After rehydrating, discs were washed once for 10min and twice for 5min with PTw. The discs were then permeabilized in 300-500μL of detergent solution (1.0% SDS, 0.5% Tween, 50mM Tris-HCL pH 7.5, 1mM EDTA pH 8.0, 150mM NaCl) for 30min at room temperature. While waiting, hybridization buffer (Molecular Instruments) was warmed to 37°C (200μL/tube). After permeabilizing with detergent solution, each tube of wing discs was incubated in 200μL of pre-warmed hybridization buffer for 30min at 37°C. Probe solution was prepared by adding 0.8pmol (0.8 μl of probe from 1uM stock solution) of each probe mixture to 200μL of probe hybridization buffer at 37°C. The pre-hybridization buffer was removed, and the probe solution was added. For negative control tissues, hybridization buffer without probes was added instead. All wing discs were incubated overnight (12-16hr) at 37°C. Before resuming the protocol, probe wash buffer (Molecular Instruments) was warmed to 37°C, amplification buffer (Molecular Instruments) was calibrated to room temperature, and a heat block was set to 95°C. The probe solution was removed and saved at -20°C to be reused. Wing discs were then washed four times in 1mL pre-warmed probe wash buffer at 37°C for 15min per wash. After the last wash step, discs were washed twice (5min/wash) with 1mL of 5x SSCT (5X sodium chloride sodium citrate, 0.1% Tween 20) at room temperature. The wing discs were pre-amplified with 1mL of equilibrated amplification buffer for 30min at room temperature. During this step, 2μL of each of hairpin h1 and 2uL of each hairpin h2 were mixed in 100μL of amplification buffer at 95°C for 90sec and cooled to room temperature in the dark for 30 min. For mirror-only in situ hybridizations, we used B1 hairpin amplifiers tagged with AlexaFluor 594 fluorophore (Molecular Instruments). For mirror and wingless double stains, we used mirror probes with B1 amplifier tagged with AlexaFluor 647 fluorophore and wingless probes (Lot #PRG129) with B3 hairpin amplifiers tagged with AlexaFluor 546 fluorophore. After 30min, the pre-amplification buffer was removed from the discs, hairpin solution added, and the tissues were then incubated in the dark at room temperature for 2-16hr. The hairpins were removed and saved at -20°C to be reused later. Excess hairpins were removed by washing 5 times (twice for 5min, twice for 30min and once for 5min) with 1mL of 5x SSCT at room temperature. The wing discs were then incubated in 50% glycerol solution (in 1X PBS) with DAPI (0.01μg/mL) overnight at 4°C. The wing discs were then mounted and visualized on a Zeiss 710 or Leica Stellaris 5 confocal microscope.

CRISPR-Cas9 mediated mutagenesis of mirror in J. coenia

Two single-guide RNAs (sgRNAs; sgRNA1-5’-GAATGGACTTGAACGGGGCA; sgRNA2-5’-AGAAACAGGGTCGATGATGA) targeting the homeobox domain of mirror were mixed with 500ng/μl of Cas9 nuclease and injected to J. coenia eggs 0.5-4hr after oviposition (n = 1042 eggs) as previously described. ²⁰ Injected G0 individuals were reared on standard artificial diet, until they emerged and were immediately frozen in -20^°C upon emergence. Wings of surviving G0 adults (n=99) were assayed for anomalies to detect mosaic knockout (mKO) phenotypes (n=29). Injection results and mutation phenotype frequencies are detailed in Table S1.

Validating mirror mutants by genotyping

DNA was extracted using E.Z.N.A. Tissue DNA Kit (Omega Bio-Tek) from the thorax of individuals that showed mutations in the wings. Extracted genomic DNA was amplified using a pair of primers flanking the sgRNA cut sites (Forward: 5’-CGCTTGTGCCCACCTTAAAC, Reverse: 5’-GTATGGCTCGGGGGATTCTG). Amplified DNA was run on a 2% agarose gel and were excised and purified using the MicroElute Cycle-Pure Kit (Omega Bio-Tek). Purified DNA was Sanger sequenced by Cornell Institute of Biotechnology. Example mutant alleles of mirror are shown in Figure S6.

Supplemental information

Figure S1.

Figure S2.

Figure S3.

Figure S4.

Figure S5.

Figure S6.

Table S1.

Acknowledgements

We thank Kate Siegel, Nigel Williams, and Rick Fandino for assistance with rearing butterflies; Johanna Dela Cruz for help with confocal microscopy at Biotechnology Resource Center (BRC) Imaging Facility (RRID:SCR_021741) at the Cornell Institute of Biotechnology. We also thank Dr. Eirene Markenscoff-Papadimitriou for use the Leica Stellaris 5 confocal microscope for imaging. This work was supported by the United States National Science Foundation grants NSF IOS-1753559 and IOS-2128164 awarded to R.D.R., and a Cornell Summer Experience Grant and an Office of Undergraduate Biology fellowship to X.Y.

Additional information

Author contributions

Conceptualization, M.C and R.D.R; Methodology, M.C and R.D.R; Investigation, X.Y.Y, M.C, N.K.B , G.C.H and C.A.A; Writing-Original Draft, M.C, X.Y.Y and R.D.R; Writing-Review & Editing, M.C and R.D.R; Visualization, M.C and X.Y.Y; Funding Acquisition, R.D.R and X.Y.Y; Supervision, M.C and R.D.R