The transcription factor Zfh1 acts as a wing-morph switch in planthoppers

Zhang, Jin-Li; Chen, Sun-Jie; Liu, Xin-Yang; Moczek, Armin P.; Xu, Hai-Jun

doi:10.1038/s41467-022-33422-6

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Published: 27 September 2022

The transcription factor Zfh1 acts as a wing-morph switch in planthoppers

Jin-Li Zhang¹,
Sun-Jie Chen¹,
Xin-Yang Liu¹,
Armin P. Moczek² &
…
Hai-Jun Xu ORCID: orcid.org/0000-0002-7314-377X¹

Nature Communications volume 13, Article number: 5670 (2022) Cite this article

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Abstract

Insect wing polyphenism is characterized by its ability to produce two or more distinct wing morphs from a single genotype in response to changing environments. However, the molecular basis of this phenomenon remains poorly understood. Here, we identified a zinc finger homeodomain transcription factor Zfh1 that acts as an upstream regulator for the development of long-winged (LW) or shorted-winged (SW) morphs in planthoppers. Knockdown of Zfh1 directs SW-destined nymphs to develop into LW morphs by down-regulating the transcriptional level of FoxO, a prominent downstream effector of the insulin/IGF signaling (IIS) pathway. The balance between transcriptional regulation via the Zfh1-FoxO cascade and post-translational regulation via the IIS-FoxO cascade provides a flexible regulatory mechanism for the development of alternative wing morphs. These findings help us understand how phenotypic diversity is generated by altering the activity of conserved proteins, and provide an extended framework for the evolution of wing morphological diversity in insects.

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Introduction

The evolution of wings was one of the most important events in the diversification of insects^1,2. Yet despite the evolutionary advantages of flight, nearly all originally winged insect orders possess many partially winged or secondarily wingless lineages^3,4,5. Wing polymorphism is an especially common version of this phenomenon, in which both flying and flightless forms are contained within the same population. Typically, the flying morph possesses fully developed wings and flight muscles, suited for long-distance dispersal. By contrast, the flightless morph possesses vestigial or no wings as well as undeveloped flight muscles, and does not disperse. Winged and wingless morphs maintain high fitness by specializing on different life histories: a winged morph able to disperse when local conditions are declining, and a wingless and more sedentary morph able to invest heavily into reproduction when local conditions are favorable^6,7. Wing polymorphic insects therefore provide many promising opportunities to investigate the adaptive significance of dispersal polymorphisms and more generally the evolution of alternative life histories. Doing so, however, is hampered by our incomplete understanding of the molecular basis underlying the development of alternative winged and wingless morphs.

Studies in diverse taxa including aphids, crickets, and planthoppers indicate that insect wing polymorphism can be caused by environmental cues encountered during particular developmental stages (polyphenism), by different genotypes, or by a combination of both^7,8,9. Juvenile hormone (JH), in particular, has been the main subject of studies into the endocrine regulation of wing polyphenism in diverse insect species, yet persuasive direct evidence documenting a functional role of JH in wing polyphenisms is still lacking^10,11,12. In contrast, growing evidence indicates that insect wing polyphenism might be regulated by diverse genes and gene regulatory networks whose identities may depend greatly on the taxa under study: for example, winged or wingless morphs are regulated, at least in part, by ecdysone signaling in female aphids Acyrthosiphon pisum¹³, whereas the male dimorphism is genetically controlled by a single locus on the X chromosome^14,15,16. In another aphid species, Aphis citricidus, small RNAs (e.g., miR-9b) also contribute significantly to wing dimorphism¹⁷.

These results contrast to findings in the wing polymorphic brown planthopper (BPH), Nilaparvata lugens (Hemiptera: Delphacidae), a destructive rice pest in Asia. During nymphal development, comprised of five nymphal stages, BPH wing buds grow roughly proportionally with body size. Nymphs eventually mature into distinct, short (SW)- and long-winged (LW) morphs which become externally distinguishable for the first time following the terminal nymphal-to-adult molt. SW adults possess vestigial forewings and rudimentary hindwing buds, in contrast to fully developed forewings and hindwings typical of LW adults (Fig. 1a, Supplementary Fig. 1a). Diverse environmental cues, including crowding, host plant quality, photoperiod, and temperature, have been shown to influence BPH wing-morph switching, although dominant environmental regulators—if they indeed exist—remain yet to be identified^12,18. The ability to develop into the LW morph enables BPHs to migrate northward from tropical to subtropical areas in the spring, followed by returning migrations in the autumn, causing severe rice yield loss throughout Asia^19,20,21. Previous studies showed that the forkhead transcription factor subgroup O (FoxO) serves as a key regulator that directs the development of LW or SW morphs^22,23, and may play an important role in interfacing with environmental cues such as wounding and stress^24,25. Additional studies indicated that the insulin/IGF signaling (IIS) pathway affects BPH wing dimorphism via modulating the phosphorylation level of FoxO^22,26,27, resulting in phosphorylated FoxO to be excluded from the nucleus and thus unable to inhibit wing development, thereby leading to LW morphs. Interestingly, the IIS pathway has also been implicated in the regulation of wing dimorphism in both the red-shouldered soapberry bug Jadera haematoloma²⁸ and the European firebug Pyrrhocoris apterus²⁹, likely constituting independent yet parallel co-option events. Yet despite the prominence of the IIS-FoxO signaling axis in the regulation of wing dimorphism, we still know little about how it interfaces with the gene regulatory networks underpinning wing formation in insects.

**Fig. 1: Knockdown of BPH *Zfh1* leads to long-winged morphs.**

In this work, we performed comparative transcriptomic analysis on SW-destined and LW-destined BPH nymphs, and identified a zinc-finger homeodomain transcription factor Zfh1 that acts as a wing-morph switch in planthoppers. We find that knockdown of Zfh1 directs SW-destined nymphs to develop into LW morphs by affecting the transcription of FoxO. In addition, we find that Zfh1 also acts independently of both the IIS and Ultrabithorax pathways in regulating wing-morph development. These findings deepen our understanding of the genetic bases underpinning insect wing polymorphism.

Results

Knockdown of Zfh1 results in LW morph development

To screen potential wing-morph regulators, we conducted RNA sequencing (RNA-seq) on thoracic nota (mesonotum and metanotum) dissected from 0–72 h fifth-instar nymphs of wild-type SW-BPH (Wt^SW, SW ratio > 90%) and LW-BPH (Wt^LW, LW ratio > 80%) strains, representing SW-destined and LW-destined BPHs, respectively. The fifth-instar stage was chosen because it is the instar during which SW and LW developmental fates are determined³⁰. We identified 123 putative transcription factors (TFs) including FoxO and Zfh1 (Fig. 1b, Nl.chr09.837 in Supplementary Data 1) that were significantly differentially expressed in Wt^LW versus Wt^SW nymphs. Next, we performed RNA interference (RNAi)-mediated gene silencing by microinjection of Wt^SW nymphs with double-stranded RNA (dsRNA) targeting each TF gene except for FoxO which has been examined previously^22,26. Microinjection of dsRNA targeting Zfh1 (dsZfh1, Fig. 1b) reduced the transcriptional level of Zfh1 by ~40% compared to dsRNA targeting green fluorescent protein (dsGfp, Fig. 1c). Both female (Fig. 1d, e) and male (Supplementary Fig. 1b, c) nymphs treated with dsZfh1 exhibited a strong and comparable bias towards LW morphs (dsZfh1^LW), relative to control individuals injected with dsGfp (dsGfp^SW). In contrast, no effect on wing morph determination was detected following dsRNA-mediated knockdown of any of the other 121 TF genes (Supplementary Data 2). Hypomorphic knockdown phenotypes were detectable in a small fraction of dsZfh1-treated adults with intermediate-size wings (dsZfh1^IMW, Fig. 1d, e, Supplementary Fig. 1b, c). Notably, dsZfh1 had no detrimental effect on nymph development and survival (Supplementary Fig. 2). Additionally, dsZfh1^LW BPHs exhibited hind tibia lengths, a common metric for body size in BPH, comparable to Wt^LW BPHs (Fig. 1f, Supplementary Fig. 1d), suggesting that dsZfh1 had no obvious effect on body size. However, the forewing size of dsZfh1^LW BPHs was slightly but significantly smaller than that of Wt^LW BPHs although the vein patterning was within the normal range of variation observed in Wt^LW BPHs (Fig. 1f, g, Supplementary Fig. 1d, e). In addition to wings, dsZfh1^LW BPHs resembled Wt^LW BPHs with regards to indirect flight muscle (IFM) development. Specially, immunohistochemistry (IHC) staining showed that IFM was abundant only in the thorax of dsZfh1^LW and Wt^LW BPHs, but not in Wt^SW BPHs (Fig. 2a). Further, examination by transmission electron microscopy (TEM) showed that dsZfh1^LW and Wt^LW but not Wt^SW BPHs contained well-organized sarcomeres with visible Z and M discs (Fig. 2b). This observation indicates that Zfh1 inhibits IFM differentiation in BPHs, in line with the inhibitory role of Zfh1 in muscle differentiation described in Drosophila and vertebrates^31,32,33.

**Fig. 2: Knockdown of *Zfh1* induces the development of indirect flight muscles.**

To confirm the dsZfh1 phenotype, we conducted Zfh1 knockdown using a second non-overlapping dsRNA targeting Zfh1 (dsZfh1_2, Fig. 1b). Microinjection with dsZfh1_2 reduced the transcriptional level of Zfh1 by ~50% compared to dsGfp treatment (Fig. 1c). As with dsZfh1, the majority of dsZfh1_2-treated Wt^SW nymphs developed into LW adults (Fig. 1e, Supplementary Fig. 1c). Additionally, quantitative real-time PCR (qRT-PCR) analysis showed that Zfh1 expression exhibited a distinct spatiotemporal pattern, with low level of Zfh1 detected in the thorax and thoracic nota of fifth-instar Wt^LW relative to Wt^SW nymphs (Supplementary Fig. 3), indicating that the expression of Zfh1 is inversely correlated with LW development.

To investigate whether Zfh1 was functionally conserved in the planthopper family Delphacidae, we performed RNAi-mediated knockdown of the Zfh1 homolog in the planthopper Laodelphax striatellus (LsZfh1). Microinjection with dsRNA targeting LsZfh1 (dsLsZfh1) significantly decreased the transcriptional level of LsZfh1 (Supplementary Fig. 4a), and significantly increased the LW ratio (dsLsZfh1^LW, Fig. 3a, b) relative to dsGfp treatment, consistent with the phenotype observed in BPHs. Notably, hypomorphic knockdown of LsZfh1 also led to adults with intermediate-size wings (dsLsZfh1^IMW). Together, these findings indicate that Zfh1 acts as a molecular switch that regulates alternative wing morphs in planthoppers.

**Fig. 3: Knockdown of *L. striatellus Zfh1* homolog (*LsZfh1*) leads to long-winged (LW) morphs.**

Functional specificity of Zfh1 among BPH homeobox genes

BPH Zfh1 contains a homeodomain in addition to nine Cys2His2 type zinc-finger motifs (Fig. 1b), and as such belongs to the zfh family of homeobox genes across metazoan phyla³⁴. As does Drosophila, BPH possesses two zfh family members, Zfh1 and Zfh2, which differ in the number and position of homeodomains and zinc-finger motifs. Studies in Drosophila indicated that both Zfh1 and Zfh2 are involved in neurogenesis and cell differentiation^35,36,37. In addition, Drosophila Zfh1 functions in the positioning of mesoderm derivatives including somatic musculature³⁸, whereas Zfh2 is involved in the proximal-distal patterning of Drosophila wing discs^39,40. Phylogenetic analyses show that both Zfh1 and Zfh2 are conserved across animals (Supplementary Fig. 5a): invertebrate Zfh1 homologs cluster with two vertebrates Zfh1 orthologues (Zeb1 and Zeb2)³⁸, whereas Zfh2 and other zfh family members cluster separately (Supplementary Fig. 5a).

To test whether BPH Zfh2 is functionally redundant to Zfh1 with respect to the regulation of wing dimorphism, we conducted Zfh2-specific knockdowns by microinjection of fourth-instar Wt^SW nymphs with dsRNA targeting Zfh2 (dsZfh2). Notably, Zfh2 knockdown caused ~50% mortality of nymphs before adult eclosion, while surviving adults exhibited curved wings (Supplementary Fig. 5b). However, we found that surviving dsZfh2-treated BPHs produced a high proportion of SW morphs, similar to Wt^SW BPHs (Supplementary Fig. 5c), indicating that Zfh2 is not involved in BPH wing dimorphism.

To assess whether any additional homeobox genes might be involved in LW development, we conducted RNAi targeting the remaining 87 potential BPH homeobox genes in Wt^SW nymphs. We found that knockdown of any of these 87 genes did not change wing-morph ratios (Supplementary Data 3). Thus, our results suggest that Zfh1 may be the sole BPH homeobox gene that functions specifically in the regulation of wing dimorphism.

Zfh1 regulates FoxO transcriptional activity

To further detail the role of Zfh1 in the development of wing dimorphism, we collected the mesonotum and metanotum of dsZfh1- and dsGfp-treated fifth-instar nymphs for RNA-seq (Fig. 4a). Interestingly, FoxO was one of the 478 genes downregulated following dsZfh1 injection (Fig. 4b, Supplementary Data 4). This is notable because dsZfh1 phenocopied the effects of FoxO knockdown (dsFoxO) on LW development²². qRT-PCR analysis confirmed that dsZfh1 significantly reduced FoxO mRNA level (Fig. 4c).

**Fig. 4: Zfh1 regulates the transcription of *FoxO*.**

To further assess whether FoxO is indeed a target gene transcriptionally regulated by Zfh1, we examined FoxO translation following Zfh1 knockdown, using a fusion protein consisting of FoxO and Human influenza hemagglutinin (HA) as a reporter. To do so, we microinjected eggs with a mix of Cas9, single-guide RNA (sgRNA), and single-strand DNA (ssDNA) donors to construct a knock-in BPH strain (FoxO::HA) using clustered regularly interspaced palindromic repeats/CRISPR-associated (CRISPR/Cas9)-mediated homology-directed repair. The resulting FoxO::HA mutant then allowed us to detect FoxO protein levels using commercially available anti-HA antibodies. We found that 13.2% (n = 1023) injected egg successfully molted into nymphs (G0), among which 11 out of 44 examined nymphs (25%, Supplementary Table 1) contained an HA-tag at the C terminus of FoxO. To examine germline transmission, G0 adults were mated with Wt^SW adults to produce G1 offspring. We found that 26.7% G1 individuals (32 out of 120, Supplementary Table 1) contained FoxO with an HA insertion (Fig. 4d). Crossing G1 siblings then allowed us to obtain a FoxO::HA homozygous mutant. To verify the FoxO::HA mutant, whole bodies of FoxO::HA mutants and Wt^SW adults were homogenized for western blot assay using an HA antibody. We found that the FoxO-HA fusion protein was specifically detected in FoxO::HA mutants rather than in Wt^SW BPHs (Fig. 4e). To further confirm the FoxO::HA mutant, genomic DNA of FoxO::HA mutants was extracted and then used for Sanger sequencing with primers (FoxO-KI-F/FoxO-KI-R, Fig. 4d) located outside of homology arms. Sanger sequencing showed that the HA-tag was correctly inserted before the stop codon (TAG) of FoxO (Fig. 4f). Next, we performed Zfh1 knockdown in FoxO::HA mutants, followed by western blot assay using an HA antibody. In line with the qRT-PCR data, dsZfh1 significantly reduced the FoxO::HA protein expression compared to dsGfp (Fig. 4g).

Given that FoxO is transcriptionally regulated by Zfh1, we asked whether Zfh1 protein could bind to the FoxO promoter, similar to the vertebrate Zfh1 homolog which represses gene expression by binding to the consensus sequence (CACCT and CACCTG)⁴¹. We identified three high-affinity Zfh1 binding sites in the first 2.5 kb fragment of the FoxO promoter (ProFoxO, Fig. 4h). Using an immunoprecipitation (IP) protein-expressing Zfh1-his in human embryonic kidney 293 cells (HEK293T), we found that the FoxO promoter sequence was significantly enriched by anti-His antibodies when ProFoxO was co-transfected with pCMV-Zfh1-His (Fig. 4i), confirming Zfh1 binding to the FoxO promoter.

To further ascertain whether FoxO is positioned downstream of Zfh1, thoraxes dissected from dsFoxO-, dsZfh1-, and dsGfp-treated fifth-instar nymphs were subjected to RNA-seq. The comparative transcriptomic analysis revealed that 55% of genes (546 out of 997) regulated by dsFoxO were also regulated by dsZfh1 (Fig. 4j, Supplementary Data 5). Among these, 96% (522 out of 546) showed comparable changes in expression (Fig. 4j). Gene ontology (GO) analysis showed important similarities as well as differences among the genes whose expression was significantly altered following dsFoxO and/or dsZfh1: genes up-regulated by both dsFoxO and dsZfh1 were significantly enriched for 9 GO terms associated with cellular processes (Supplementary Fig. 6a), whereas genes downregulated by both dsFoxO and dsZfh1 were significantly enriched for 10 GO terms associated with chitin metabolic processes (Supplementary Fig. 6b). In contrast, the top GO terms specific to genes whose expression was altered following dsFoxO were ‘mitochondrial ribosome’, ‘organellar ribosome’ and ‘mitochondrial translation’, whereas the top-ranked GO terms specific to dsZfh1 regulation were ‘structural constituent of cuticle’, ‘extracellular region’ and ‘muscle system process’ (Supplementary Fig. 6c, d). Taken together, these findings suggest that Zfh1 operates as an upstream regulator for wing dimorphism by affecting FoxO transcriptional activity.

Zfh1 regulates wing morphs in parallel to the IIS pathway

Inactivation of genes encoding positively acting components (InR1 or Akt) of the IIS pathway can lead to SW morphs by dephosphorylation of FoxO; however, inactivation of InR2, a negative regulator of InR1, enhances the phosphorylation of FoxO, leading to LW morphs²². Because FoxO is an effector downstream of the IIS pathway and because FoxO is transcriptionally regulated by Zfh1, we asked whether Zfh1 regulates the expression of FoxO by relaying IIS activity. If this is the case, the inactivation of Zfh1 would abolish any effects of the IIS pathway on wing dimorphism in the same manner as done by FoxO²². Thus, we performed a double-gene knockdown using dsFoxO and dsZfh1 in combination with a dsRNA targeting InR1 (dsInR1) or Akt (dsAkt). Although dsZfh1 combined with dsGfp led to LW morphs, Wt^SW nymphs developed into SW adults when dsZfh1 was combined with either dsInR1 or dsAkt (Fig. 5a, Supplementary Fig. 7). As a control, Wt^SW nymphs developed into LW adults when dsFoxO was combined with dsInR1 or dsAkt (Fig. 5a, Supplementary Fig. 7). These data indicate that Zfh1 is not a TF downstream of the IIS pathway.

**Fig. 5: Zfh1 regulates wing dimorphism independent of the IIS pathway.**

Next, we sought to determine whether Zfh1 is positioned upstream of the IIS pathway. If correct, inactivation of Akt would abolish the regulatory role of Zfh1 on FoxO protein expression. To test these predictions, we microinjected FoxO::HA nymphs with dsZfh1, dsAkt, a dsRNA targeting InR2 (dsInR2), a dsRNA mixture targeting Zfh1 and Akt (dsZfh1;dsAkt), or dsGfp, followed by the dissection of the nota of fifth-instar nymphs for western blot assay. Compared to dsGfp, double-gene knockdown using dsZfh1;dsAkt significantly reduced FoxO::HA fusion protein levels, as did dsZfh1 alone (Fig. 5b), indicating that Zfh1 is also not a factor upstream of the IIS pathway. Notably, both dsInR2 and dsAkt resulted in comparable levels of FoxO::HA protein relative to dsGfp (Fig. 5b). This supports previous reports that the IIS pathway may regulate FoxO protein function via post-translational modification⁴².

To exclude any possible crosstalk between the IIS pathway and Zfh1, we generated a Zfh1::HA knock-in mutant using CRISPR/Cas9-mediated homology-directed repair, which would allow us to detect Zfh1 protein with anti-HA antibodies since Zfh1 was fused an HA-tag with at the C terminus (Fig. 5c). The CRISP/Cas9-medicated mutagenesis rate and germline transmission for Zfh1 reached 11.6% (n = 64) and 18.9% (n = 90), respectively. The Zfh1::HA homozygous mutant was verified by western blot using anti-HA antibodies (Fig. 5d) and by Sanger sequencing (Fig. 5e) with primers (Zfh1-KI-F/Zfh1-KI-R, Fig. 5c) located outside of the homology arms. The Zfh1::HA mutant allowed us to detect Zfh1 protein in the context of perturbation of the IIS pathway using anti-HA antibodies. We found that knockdown of components of the IIS pathway (InR1, InR2, and Akt) did not affect Zfh1 protein level (Fig. 5f). Meanwhile, knockdown of Zfh1 did not affect the transcriptional level of InR1, InR2, and Akt (Fig. 5g). Thus, these findings indicate that Zfh1 regulates wing dimorphism independent of the IIS pathway in BPHs.

Zfh1 functions synergistically with the IIS pathway

Given that Zfh1 regulates LW development in parallel to the IIS pathway, we asked whether Zfh1 and the IIS pathway may interact synergistically in the regulation of LW development. For this purpose, we performed Zfh1 knockdown in the context of an InR2-null mutant (InR2^E4)⁴³, which by itself results in 100% LW morphs (Fig. 5h) due to accumulated phosphorylated FoxO in the cytoplasm via activating the IIS pathway. We found that dsZfh1-treated InR2^E4 mutants (InR2^E4::dsZfh1) developed forewings that were 7% larger than the InR2^E4 mutants treated with dsGfp (InR2^E4::dsGfp, Fig. 5i), indicating a moderate synergistic effect of Zfh1 and the IIS pathway on LW development.

Zfh1 is functionally distinct from Ultrabithorax

The presence of a homeodomain in BPH Zfh1 combined with the Zfh1 knockdown phenotype are reminiscent of the ‘four-winged Drosophila’ phenotype derived from Ultrabithorax (Ubx) mutants⁴⁴. This phenotype results from the homeotic transformation of the third thoracic segment (T3) to the identity of the second thoracic segment (T2), and the concomitant transformation of halters into hindwings in Drosophila⁴⁴, or membranous hindwings into elytra in the flour beetle Tribolium castaneum⁴⁵.

To exclude the possibility that dsZfh1 produced LW morphs via homeotic transformation, we compared and contrasted the morphological characteristics of wings and thoracic segments between dsZfh1 and Ubx-RNAi (dsUbx) adults, focusing on three aspects: (i) dsUbx adults that were derived from RNAi in SW-destined nymphs developed only small wing-like appendages on T3 and SW forewings at T2 (Fig. 6a), in strikingly contrast to the fully developed forewings and hindwings of dsZfh1 BPHs (Fig. 1d); (ii) dsZfh1 adults that were derived from RNAi in LW-destined nymphs developed membranous and fully transparent hindwings similar to the control BPHs (dsGfp, Fig. 6b), whereas dsUbx adults that were derived from RNAi in LW-destined nymphs developed heavily-pigmented hindwings (Fig. 6b), resembling forewings; and (iii) dsZfh1 adults developed T2 and T3 nota identical to those of dsGfp-treated BPHs (Fig. 6c), whereas Ubx knockdown led to an expansion of the T3 notum (Fig. 6c), thereby causing it to resemble the characteristics of the T2 notum. In addition, Zfh1 and Ubx differed strikingly in terms of tissue-specific expression. qRT-PCR assay showed that hindwing buds had a slightly but significantly higher level of Zfh1 relative to forewing buds (Fig. 6d). In contrast, Ubx expression was strongly biased toward hindwing buds relative to forewing duds (Fig. 6d). Together, these observations indicate that the SW-to-LW transition derived from Zfh1 knockdown is most likely not due to homeotic transformation.

**Fig. 6: Phenotypic comparison of dsZfh1 and dsUbx BPHs.**

A molecular model of wing dimorphism in planthoppers

Our data revise and significantly extend our understanding of how distinct wing morphs may result from modifications of the same gene regulatory networks (Fig. 7). Under normal conditions, BPHs maintain a basic level of Zfh1, which in turn maintains FoxO transcription at low levels. As a result, low activity of the Zfh1-FoxO signaling cascade prevents FoxO from inhibiting wing development, thus leading to LW morphs. However, in response to certain environmental inputs, BPHs raise Zfh1 transcription, thereby raising FoxO levels, which in turn suppresses LW development and therefore leads to SW morphs. In addition to this Zfh1-FoxO signaling cascade, wing dimorphism is regulated by a second, and independent, signaling axis at the post-translational level: activation of the IIS pathway phosphorylates FoxO and thus exports FoxO from the nucleus to the cytoplasm, leading to LW morphs. In contrast, inactivation of the IIS pathway dephosphorylates FoxO and thus accumulates FoxO in the nucleus, leading to SW morphs.

**Fig. 7: Model of the molecular regulation of wing dimorphism in planthoppers.**

Discussion

We identified the zinc-finger homeodomain transcription factor Zfh1 as a wing-morph molecular switch that determines long or short wings by regulating the transcriptional level of a second transcription factor FoxO in BPHs. Previous studies indicated that Zfh1 is an evolutionarily conserved gene critical for counteracting the myogenic differentiation program^31,34,46. In Drosophila, embryos with a Zfh1 loss-of-function mutation show alterations in somatic muscles³⁸. In the same way, a vertebrate homolog of Zfh1 (ZEB) acts as a negative regulator of muscle differentiation³¹. Here, we found that knockdown of BPH Zfh1 promoted IFM development, consistent with myogenesis suppression by Zfh1 in Drosophila and vertebrates. However, in addition to these common phenotypes reported previously, this study demonstrates for the first time that knockdown of Zfh1 directs SW-destined nymphs to develop into LW in wing dimorphic BPHs, and likely planthoppers broadly. Given that the development of LW in BPHs mainly relies on cell proliferation^23,43, this phenotype indicates that Zfh1 might act as an inhibitor of wing cell proliferation. However, whether suppression of wing cell via Zfh1 is species-dependent or conserved among insects remains unknown. Additionally, a recent study showed that the maintenance of Drosophila muscle progenitors involves a switch between Zfh1-long and Zfh1-short RNA isoforms³³. The Zfh1-long isoform is subjected to miR-8 microRNA-mediated downregulation that enables muscle progenitors to differentiate in early pupal stages, whereas the Zfh1-short isoform cannot be targeted by miR-8, thus allowing these cells to persist as muscle progenitors in the adult stage³³. In this study, only one Zfh1 isoform was identified after examining all BPH transcriptomes prepared by our lab. Thus, whether multiple BPH Zfh1 isoforms, if they do indeed exist, possess distinct roles in the regulation of wing development and muscle homeostasis needs to be clarified in further studies.

Our previous studies showed that FoxO plays a pivotal role in controlling the development of alternative wing morphs in BPHs via relaying the activity of the IIS pathway²². Activation of the IIS pathway phosphorylates and deactivates FoxO, leading to LW morphs, while IIS inactivation and FoxO dephosphorylation promote SW morphs. Here, we found that knockdown of Zfh1 stimulates LW development by decreasing FoxO transcriptional activity. These findings highlight that the Zfh1 and IIS cascades may converge on FoxO to regulate wing dimorphism in BPHs. This assumption is supported by a synergistic effect of both pathways on LW development. However, exactly how BPHs balance the relative activities of the Zfh1-FoxO and IIS-FoxO cascades to regulate wing developmental plasticity remains to be determined. More generally, our results suggest that this regulatory system may provide a multi-tiered regulatory machinery able to facilitate evolutionary changes in threshold responses separating wing morphs and increase the ways in which population and species may diverge genetically in the precise regulation of wing polyphenism.

Methods

Insects

Laboratory populations of N. lugens were established from colonies collected in Hangzhou, China, in 2008. The Wt^SW colony was purified by inbreeding for more than 13 generations⁴⁷. The Wt^LW strain was provided by Dr. Hong-Xia Hua (Huazhong Agricultural University, China). The planthopper L. striatellus was collected in Hangzhou, China, 2018. All insects were reared at 26 ± 0.5 °C under a photoperiod of 16:8 h (L:D) at a relative humidity of 50 ± 5% on rice seedlings (rice variety: Xiushui 11).

Sample preparation for RNA-seq

To screen for potential wing-morph regulators, mesonotum and metanotum were dissected from 0–72 h fifth-instar Wt^SW (n = 20) and Wt^LW nymphs (n = 20), denoted as 5th-SW and 5th-LW respectively (BioProject ID: PRJNA805393). To identify downstream effectors targeted by Zfh1, third-instar Wt^SW nymphs were microinjected with 50 ng dsZfh1 or dsGfp. Subsequently, mesonotum and metanotum were dissected from 36–48 h fifth-instar nymphs with three replicates (n = 30 for each replicate), denoted as Zfh1-nota and Gfp-nota (BioProject ID: PRJNA805395). To investigate common genes regulated by Zfh1 and FoxO, third-instar Wt^SW nymphs were microinjected with 50 ng dsZfh1, dsFoxO, or dsGfp. Thoraxes were dissected from 48–54 h fifth-instar nymphs with three replicates (n = 30 for each replicate), denoted as Zfh1-T, FoxO-T and Gfp-T (BioProject ID: PRJNA805400). Samples were homogenized for total RNA isolation using RNAiso Plus (Cat#9109, Takara) according to the manufacturer’s protocol.

cDNA library preparation and Illumina sequencing

A total of 1 µg RNA per sample was used to construct a sequencing library using a NEBNext Ultra RNA library prep kit for Illumina (Cat#E7770, NEB) according to the manufacturer’s recommendations, and index codes were added to each sequence. Library fragments of 250–300 bp in length were preferentially purified using an AMPure XP system (Beckman Coulter). Clustering of the index-coded samples was performed on a cBot Cluster generation system using a TruSeq PE cluster kit v3-cBot-HS (Cat#PE-401-3001, Illumina) according to the manufacturer’s instructions. The cDNA libraries were sequenced on an Illumina Novaseq 6000 platform and 150 bp paired-end reads were generated.

Read mapping and differentially expressed genes (DEGs)

After Illumina sequencing, clean reads were generated by removing adapters, poly-N, and low-quality reads from the raw data using fastp algorithm (v0.12.4)⁴⁸. Clean reads were mapped against the N. lugenes genome v4.0⁴⁷ using hisat2 (v2.1.0)⁴⁹, and transcript abundance was quantified using StringTie (v1.3.5)⁵⁰. Fragments Per Kilobase of transcript per Million mapped reads (FPKM) was used to quantify the expression level of each transcript. The read count information of each transcript was extracted directly from the files generated by StringTie using a python script of prepDE.py (http://ccb.jhu.edu/software/stringtie/dl/prepDE.py). Differential expression analysis of 5th-SW and 5th-LW samples was conducted using the edgeR package (v3.38.4)⁵¹, and genes with a false discovery rate (FDR) < 0.01 were considered to be significant. Differential expression analysis of Zfh1-nota versus Gfp-nota, Zfh1-T versus Gfp-T, and FoxO-T versus Gfp-T was conducted using the DESeq2 package (v1.36.0)⁵² and gene expression change was considered to be significant under the FDR < 0.05. To generate heat map, FPKM of each replicate of Zfh1-T, FoxO-T and Gfp-T were z-score transformed and clustered using the online OmicShare tool (https://www.omicshare.com/tools/Home/Soft/heatmap). GO enrichment analysis of DEGs was performed using the online OmicShare tool (https://www.omicshare.com/tools/home/report/goenrich.html), and the enriched GO terms with FDR ≤ 0.05 were considered as significant.

RNAi and RNAi efficiency

DsRNA synthesis and injection were performed as previously described²². Briefly, primers were synthesized with T7 RNA polymerase promoter at both ends (Supplementary Data 6), and then dsRNAs were synthesized using a T7 high yield RNA transcription kit (Cat#TR101-02, Vazyme) according to the manufacturer’s instructions. Microinjection was performed using a FemtoJet microinjection system (Eppendorf). Each third-instar or fourth-instar nymph was microinjected with approximately 50 ng or 100 ng dsRNAs, respectively. Two days after microinjection, insects (n = 5 for each of three replicates) were collected for RNAi efficiency examination by qRT-PCR. The ribosomal protein S11 gene (rps11)⁵³ and ribosomal protein L5 gene (rpl5)⁵⁴ were used as the internal reference gene in BPH and L. striatellus planthopper, respectively.

qRT-PCR assay

Total RNAs were isolated from BPHs using RNAiso Plus (Cat#9109, Takara) according to the manufacturer’s protocol. The first-strand cDNA was synthesized from total RNAs (900 ng) using HiScript QRT super mix (Cat#R123-01, Vazyme). The qRT-PCR was conducted on a CFX96 real-time PCR detection system (Bio-Rad) with the following conditions: denaturation for 3 min at 95 °C, followed by 40 cycles at 95 °C for 10 s, and then 60 °C for 30 s. The rps11 and rpl5 were used as the internal reference gene in BPH and L. striatellus planthopper, respectively. The 2^−∆∆Ct method (Ct represents the cycle threshold) was used to measure relative expression levels. Three biological replicates were used for statistical comparison between samples.

Nymphal duration and survival rate

Third-instar Wt^SW nymphs were microinjected with 50 ng dsZfh1 or dsGfp. The developmental times of fourth- and fifth-instar stages and survival rate were monitored every 12 h. To determining the nymphal duration, 13 females and 19 males were used for dsZfh1, and 17 females and 11 males were used for dsGfp. To conduct the survival assay, 50 nymphs were used for dsZfh1 and dsGfp.

Spatiotemporal expression of Zfh1

To investigate the temporal expression of Zfh1 in Wt^LW and Wt^SW BPHs, total RNAs were isolated from the thorax of first-instar (n = 100), second-instar (n = 50), third-instar (n = 50), fourth-instar (n = 30), fifth-instar nymphs (n = 15), and female adults (n = 15). To investigate the spatial expression of Zfh1, fifth-instar Wt^LW and Wt^SW nypmphs (n = 50) were collected for tissue dissection, and then total RNA was isolated from head, fat body, the abdominal cuticle, six legs, the whole digestive tract (gut), and nota (mesonotum and metanotum), respectively. To investigate the expression pattern of Zfh1 across the fifth-instar stage, mesonotum and metanotum were dissected from 24 h-, 48 h-, and 72 h-fifth-instar Wt^LW and Wt^SW nymphs (n = 50), and then used for total RNA isolation. Three independent biological replicates were used for RNA isolation, and the first-strand cDNA was synthesized for Zfh1 quantification. The relative expression level of Zfh1 was normalized to that of the rps11 gene.

IHC staining

IHC staining was performed as previously described⁵⁵. Briefly, third-instar nymphs were microinjected with 50 ng dsZfh1. Samples were fixed using 4% paraformaldehyde in PBS overnight at 4 °C, following by blocking with Tissue-Tek O.C.T. Compound (Cat#4583, Sakura Finetek) at −80 °C. Samples were longitudinally cut into ~30 μm sections using a Lecia CM1900 cryotome (Leica Microsystems) at −20 °C, and then transferred to Superfrost⁺ slides (Cat#12-550-15, Thermo Fisher Scientific). The cytoskeleton and nucleus were stained using 100 nM rhodamine-labeled phalloidin (Cat# 40734ES75, Yeasen) and 100 nM DAPI (Cat#D9542, Sigma Aldrich), respectively. Fluorescence images were acquired using a Zeiss LSM 800 confocal microscopy (Carl Zeiss MicroImaging).

TEM assay

Third-instar nymphs were microinjected with dsZfh1. Thoraxes were dissected from 24 h dsZfh1^LW females for transmission electron microscopy as previously described⁵⁵. Briefly, samples were fixed in 2.5% glutaraldehyde overnight at 4 °C, followed by post-fixation in 1% osmium tetroxide for 1.5 h. The samples were then dehydrated in a standard ethanol/acetone series, infiltrated and embedded in Spurr medium, and then superthin sections (~70 nm) were cut. The sections were stained with 5% uranyl acetate followed by Reynolds’ lead citrate solution and observed under a JEM-1230 transmission electron microscope (JEOL) at the Analysis Center of Agrobiology and Environmental Sciences of Zhejiang University.

Phylogenetic analysis of Zfh1 and Zfh2

BlastP (v2.2.31) was used to search Zfh1 and Zfh2 homologs using the N. lugens Zfh1 and Zfh2 as query sequences. Multiple amino acid sequence alignment was performed using the MUSCLE algorithm with MEGA-X (v10.1.8) software. The phylogenetic tree was constructed by MEGA-X using maximum-likelihood methods under the JTT + G model with 1000 bootstrap replicates. Phylogenetic tree was annotated and edited using the iTOL tools (https://itol.embl.de/) and Adobe Photoshop CC (v19.1.9) program.

In vitro synthesis of sgRNA and Cas9 mRNA

sgRNA was designed as previously reported⁵⁶. Briefly, sgRNA was designed by manually searching genomic sequence around the region of the FoxO and Zfh1 stop codon for the sequences corresponding to 5′-N_17-20NGG-3′, where NGG is the protospacer-adjacent motif (PAM) of SpCas9 and N is any nucleotide. For in vitro transcription of sgRNA, a DNA fragment was amplified by PCR from pMD19-T sgRNA scaffold vector with a forward primer containing a T7 promoter and a reverse primer containing a partial sgRNA sequence. The PCR products were used as a template for in vitro transcription using a T7 high yield RNA transcription kit (Cat#TR101-02, Vazyme) at 37 °C overnight according to the manufacturer’s instruction.

In vitro synthesis of Cas9 mRNA was prepared as previously reported⁵⁷. Cas9 mRNA was transcribed from plasmid pSP6-2sNLS-SpCas9 vector using the mMESSAGE mMACHINE SP6 transcription kit (Cat#AM1340, Thermo Scientific) and Poly(A) tailing kit (Cat#AM1350, Thermo Scientific).

Construction of ssDNA donors

The ssDNA was produced via lambda exonuclease digestion from PCR amplicons according to a previous description⁵⁸ with minor modifications. Briefly, a dsDNA fragment containing the whole ssDNA donor cassette for HA-tag knock-in was de novo synthesized and then subcloned into a cloning vector. For FoxO::HA knock-in mutants, the ssDNA donor cassette consisted of a 125 nt left homology arm (LHA) upstream of the FoxO stop codon, a 136 nt right homology arm (RHA) downstream of the FoxO stop codon, and a HA-tag (5′-TACCCATACGATGTTCCAGATTACGCT-3′) in the middle region. For Zfh1::HA knock-in mutant, the ssDNA donor cassette consisted of a 142 nt LHA upstream of the Zfh1 stop codon, a 133 nt RHA downstream of the Zfh1 stop codon, and a HA-tag in the middle region. Then, the dsDNA fragments were used as a template for PCR amplification with a forward primer and a 5′ phosphorylated reverse primer (Supplementary Data 6). PCR products were treated with lambda exonuclease (Cat#EN6501, Thermo Scientific) to remove the 5′-phosphorylated DNA. The ssDNAs were purified by phenol-chloroform extraction, followed by isopropanol precipitation, and then dissolved nuclease-free water.

Embryo microinjection and CRISPR/Cas9-mediated homology-directed repair

Microinjection of BPH embryos was performed as previously described⁵⁷ with minor modifications. Briefly, pre-blastoderm eggs were collected within 1 h after oviposition, and microinjected with a solution mixture (~0.5 nl) containing 500 ng/μl Cas9 mRNA, 300 ng/μl sgRNA and 300 ng/μl ssDNA using a FemtoJet microinjection system (Eppendorf). To minimize mechanical damage, the posterior end of the eggs was pierced with glass needles. After microinjection, eggs were transferred to a sterile petri dish covered with filter papers that were moistened with an antibiotic solution containing tebuconazole (20 ng/ml) and kanamycin (50 ng/ml). The eggs were incubated at 30 °C and allowed to hatch into nymphs (G0).

Crossing and genotyping

DNA typing of FoxO:: HA and Zfh1::HA heterozygous or homozygous mutants was determined as described previously⁵⁷. Newly enclosed G0 adults were mated with Wt^SW adults to produce G1 progeny. After mating, one individual G0 adult was homogenized in a 1.5-ml sterile Eppendorf tube containing 100 μl of DNA extraction buffer (10 mM Tris-HCl pH 8.2, 1 mM EDTA, 25 mM NaCl, and 0.2 mg/ml proteinase K) as reported previously⁵⁹. The body lysate was incubated at 37 °C for 30 min, followed by incubation at 95 °C for 2 min to inactivate proteinase K. Then, the precipitation was removed by centrifuging at 15,000 × g for 2 min at room temperature, and the supernatant was kept for PCR amplification.

To determine the genotyping of G1 BPHs, wings were dissected from a single adult and then incubated in 100 μl wing gDNA extraction buffer (10 mM Tris-HCl pH 8.0, 10 mM EDTA, 100 mM NaCl, 39 mM DTT, 2% SDS, and 0.02 mg/ml proteinase K) overnight at 37°C. gDNA was precipitated in 100 μl isopropanol, followed by washing with 500 μl ethanol (80%). The gDNA precipitation was dissolved in ddH₂O, and then used for PCR amplification.

To confirm the correct insertion of the HA sequence in FoxO::HA mutants, PCR was conducted with primer pairs of FoxO-KI-F/HA-R or FoxO-KI-F/FoxO-KI-R (Supplementary Data 6). To confirm the correct insertion of the HA sequence in Zfh1::HA mutants, PCR was conducted with primer pairs of Zfh1-KI-F/HA-R or Zfh1-KI-F/Zfh1-KI-R (Supplementary Data 6). The FoxO-KI-F/FoxO-KI-R and Zfh1-KI-F/Zfh1-KI-R primer pairs were located outside of homology arms and the HA-R primer was located inside the HA encoding sequence. The PCR products were either directly used for Sanger sequencing or subcloned into cloning vectors, and then single clones were picked for Sanger sequencing. Those G1 individuals with the correct HA-tag insertion were collected and sibling-crossed to obtain homozygous FoxO::HA and Zfh1::HA strains.

Western blot analysis

To detect FoxO::HA or Zfh1:HA fusion protein, whole body of fourth-instar nymphs (n = 40) or thorax (mesonotum and metanotum) dissected from 48 h fifth-instar nymphs (n = 35) was pooled for homogenization. Samples were homogenized in RIPA lysis and extraction buffer (Cat#89900, Thermo Scientific), and then denatured in SDS-PAGE loading buffer. Equal amounts of protein were loaded for each lane on SDS-PAGE gel and then transferred to a polyvinylidene difluoride membrane (Cat#IPVH00010, Millipore). The membranes were blocked in TBST (TBS with 0.1% Tween-20) containing 5% non-fat dried milk powder overnight at 4 °C. After washing, the membranes were incubated with anti-HA mAb (Cat# M180-3, MBL, 1: 2000) for 1.5 h at 35 °C, followed by incubation with horseradish peroxidase (HRP) conjugated goat anti-mouse antibody (Cat#SA00001-1, Proteintech, 1: 5000) for 1 h at 35 °C. The antibody against β-actin (Cat#M1210-2, Huabio, 1:5000) was used as the loading control. Secondary antibody binding was detected using the chemiluminescence HRP substrate detection method (Cat#34577, Thermo Scientific) and imaged with the Molecular Imager ChemiDoc XRS system (Bio-Rad).

Determination of the full-length cDNA of Zfh1

Total RNA was isolated from the whole body of fifth-instar Wt^SW BPHs (n = 10) using RNAiso plus (Cat#9109, Takara). First-strand cDNA was synthesized using a HiScript-TS 5′/3′ RACE kit (Cat#RA101-01, Vazyme) according to the manufacturer’s instruction. To obtain the 5′ end sequence of Zfh1, PCR was conducted with primers of Universal-primer and Zfh1-5RACE-GSP1, followed by a nested PCR with primers of nested-primer and Zfh1-5RACE-GSP2 (Supplementary Data 6). To obtain the 3′ end sequence of Zfh1, PCR was conducted with primer of Universal-primer and Zfh1-3RACE-GSP1, followed by a nested PCR with primers of Nested-primer and Zfh1-3RACE-GSP2 (Supplementary Data 6). All amplicons were subcloned into cloning vectors and confirmed by Sanger sequencing (Tsingke Biological Technology). The cDNA sequence of Zfh1 was deposited in GenBank with accession number OM283826.

Double-gene RNAi

Third-instar Wt^SW nymphs were microinjected with 50 ng dsZfh1 or dsFoxO, followed by microinjection with 50 ng dsInR1, dsAkt, or dsGfp at the fourth-instar stage. In parallel, fourth-instar Wt^SW nymphs were microinjected with dsRNA mixtures of dsAkt and dsGfp (50 ng each) or dsInR1 and dsGfp (50 ng each). All dsRNA-treated nymphs were raised to adults, and BPHs with different wing morphs (LW, SW, and IMW) were collected for statistical analysis.

IP assay

To express Zfh1 in HEK293T cell, the recombinant fragment contained Zfh1 ORF and 6*His tag were cloned downstream of a CMV promoter, designed pCMV-Zfh1-His. A genomic DNA fragment (ProFoxO) located 4,606-bp upstream of the start codon (ATG) of Zfh1 was amplified by PCR with primers of ProFoxO-F and ProFoxO-R (Supplementary Data 6), which contains a 607-bp 5ʹUTR of Zfh1 and putative FoxO promotor region. IP assays were performed using IP assay kit (Cat#P2078, Beyotime) as per the manufacturer’s protocol. Briefly, HEK293T cells were transfected with ProFoxO in combination with pCMV-Zfh1-his or pCMV vector alone. After 36 h, the transfected cells were cross linked using 1% formaldehyde before cell lysis. Sonicated lysates were incubated overnight at 4°C with 6*His tag monoclonal antibody (Cat#66005-1-Ig, 1: 650 (4 μg), Proteintech). The ProFoxO fragments were quantified by qRT-PCR with ChamQ SYBR color qPCR master mix (Cat# Q411-02, Vazyme) using primers of qChIP-F and qChIP-R (Supplementary Data 6).

Relative expression of InR1, InR2, Akt, and FoxO in the context of Zfh1 knockdown

Third-instar nymphs were microinjected with 50 ng dsZfh1 or dsGfp. Mesonotum and metanotum were dissected from 36–48 h fifth-instar nymphs (n = 25) and then used for total RNA isolation. The first-strand cDNA was synthesized with random primers and then used as templates for qRT-PCR. The relative expression level of InR1, InR2, Akt, and FoxO was normalized to that of the rps11 gene. Five independent biological replicates with three technical replicates were conducted for each experiment.

Relative expression of Zfh1 and Ubx in wing buds

Wing buds on the second and third thoracic segments were dissected from 48 h fifth-instar Wt^SW nymphs (n = 150), and then used for RNA isolation. The first-strand cDNA was synthesized with random primers and then used as templates for qRT-PCR with primers specific to Zfh1 and Ubx (Supplementary Data 6). The relative expression level of Zfh1 was normalized to that of the rps11 gene. Three independent biological replicates with three technical replicates were conducted for each experiment.

Scanning electron microscope (SEM)

Fourth-instar Wt^LW nymphs were microinjected with dsUbx, dsZfh1, or dsGfp, and allowed to molting into adults. After removing wings, female adults were placed on a stub and dried in an ion sputter (Hitachi) under vacuum. After gold sputtering, the samples were observed under SEM (TM-1000, Hitachi).

Image acquisition

Images of insects were taken using a DVM6 digital microscope (Leica Microsystems) with LAS X software. Images of wings and tibia were captured with a DFC320 digital camera attached to a Leica S8AP0 stereomicroscope using the LAS (v. 3.8) digital imaging system. Digital images of forewings (n = 20) and hind tibias (n = 20) were collected for measurement of forewing size and hind tibia length using ImageJ (v. 1.47). Plotting and statistical analysis were performed with GraphPad Prism (v8.0.1).

Reporting summary

Further information on research design is available in the Nature Research Reporting Summary linked to this article.

Data availability

All data are available in the manuscript or the supplementary materials. The DNA sequencing data generated in this study have been deposited in the GenBank database under the following accession codes: OM283826 (the Zfh1 gene), XP_039284941.1 (the Zfh2 gene), OM676634 (the LsZfh1 gene). The RNA sequencing data generated in this study have been deposited under the following accession coded: PRJNA805393, PRJNA805395, and PRJNA805400. Source data are provided with this paper.

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Acknowledgements

H.J.X. was supported by the National Natural Science Foundation of China (31972261, 32272519, and 31772158), Key Projects of Natural Science Foundation of Zhejiang Province (LZ21C140002), and Development and Demonstration of Comprehensive Technology for Prevention and Control of Major Diseases and Pests (2021YFD1401100). J.L.Z. was supported by the National Natural Science Foundation of China (32102186). We thank Prof. Hongxia Hua in Huazhong Agricultural University for providing the Wt^LW strain.

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Authors and Affiliations

State Key Laboratory of Rice Biology, Key Laboratory of Biology of Crop Pathogens and Insects of Zhejiang Province, Institute of Insect Sciences, College of Agriculture and Biotechnology, Zhejiang University, 866 Yu-Hang-Tang Ave, Hangzhou, China
Jin-Li Zhang, Sun-Jie Chen, Xin-Yang Liu & Hai-Jun Xu
Department of Biology, Indiana University, 915 East 3rd Street, Bloomington, IN, 47405, USA
Armin P. Moczek

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Jin-Li Zhang
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Sun-Jie Chen
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Xin-Yang Liu
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Armin P. Moczek
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Hai-Jun Xu
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Contributions

J.L.Z. and H.J.X. conceived the study and developed the initial study design. J.L.Z., S.J.C., and X.Y.L. performed the experiments. J.L.Z. and H.J.X. analyzed the experiments. A.P.M. and H.J.X. wrote the manuscript. All authors contributed intellectually to the experiments as well as editing and approval of the final version of the manuscript.

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Correspondence to Hai-Jun Xu.

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Nature Communications thanks Takahiro Ohde and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. Peer reviewer reports are available.

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Zhang, JL., Chen, SJ., Liu, XY. et al. The transcription factor Zfh1 acts as a wing-morph switch in planthoppers. Nat Commun 13, 5670 (2022). https://doi.org/10.1038/s41467-022-33422-6

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Received: 19 May 2022
Accepted: 16 September 2022
Published: 27 September 2022
DOI: https://doi.org/10.1038/s41467-022-33422-6

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