Juvenile hormone degradation enzymes have shared and unique requirements in Drosophila development

Juvenile hormone, Juvenile hormone epoxide hydrolase, Juvenile hormone esterase, 26 ecdysone, metamorphosis. 27 This PDF file includes: 28 Main Text 29 Figures 1 to 5 30 31 32 33 34 35 36 37 .CC-BY-NC 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted June 12, 2025. ; https://doi.org/10.1101/2025.06.09.657647doi: bioRxiv preprint 2

38 Precise control of hormones is essential to development and reproduction. Hormone 39 bioavailability is regulated by their synthesis and transport, as well as sequestration, natural 40 turnover, and programmed degradation. Here, we use Drosophila melanogaster to investigate 41 developmentally programmed degradation of the retinoic acid-like Juvenile hormones (JHs) that 42 functionally oppose the steroid hormone, ecdysone. Elevated JH titers promote growth and 43 prevent metamorphosis. To understand the role of programmed JH degradation, we generated 44 double and triple knockout animals lacking one of two classes of JH degradation enzymes: JH 45 esterases (JHEs) and JH epoxide hydrolases (JHEHs). While both classes of enzymes degrade 46 JHs, we found they have separate developmental functions such that JHEs restrain growth while 47 JHEHs support growth and ensure timely metamorphosis. To investigate the nature of these 48 unique developmental requirements, we performed genetic and hormone rescue experiments, 49 analyzed hormone-producing glands, and characterized dysregulated gene networks by RNA-50 sequencing. Together, these data revealed unique dysregulated networks consistent with the 51 separate developmental requirements, as well as shared regulation of Cytochrome P450 gene 52 expression and the size of the ecdysone-producing gland. Strikingly, the four-day delay in 53 metamorphosis in the absence of JHEHs is rescued by ecdysone despite increased ecdysone 54 synthesis gene expression and larger glands in these animals. Together, this study provides new 55 genetic tools and insights into the complexities of programmed hormone degradation in 56 development. 57 58 Main Text 59 60

61 62 Development, physiology and reproduction are controlled by hormones. Hormones are a diverse 63 class of secreted, small molecules that include peptides and lipids that impart dramatic and 64 systemic responses. Hormones often act in concert or in opposition in cycles of stasis and 65 progression. With simplified signaling cascades, broad receptor expression, and a high degree of 66 potency, the precise and rapid control of hormones presents a significant biological challenge. 67 68 A wealth of insights into hormone regulation has been gained using insect model organisms. 69 Insect development and physiology are driven by the steroid hormone ecdysone, and the retinoic 70 acid-like sesquiterpenoid Juvenile hormones (JHs). Whereas much is known about ecdysone, 71 large gaps in knowledge persist for JHs. JH synthesis begins with the conversion of acetyl-CoA to 72 farnesyl-PP through the mevalonate pathway and ends with the downstream JH-specific 73 enzymatic cascade that includes epoxidation of farnesoic acid (1), and methylation by JH acid 74 methyltransferase (Jhamt) (2). JH synthesis primarily occurs in a neuroendocrine corpora allatum, 75 which is part of the neuroendocrine ring gland in larvae (3). In juvenile and adult animals, JHs 76 circulate through the hemolymph via JH binding proteins, enter responding cells, and bind to 77 transcription factors, Methoprene-tolerant (Met) or Germ cell expressed (Gce) to induce 78 transcription-dependent responses (Fig 1a) (4). Decreased JH signaling shortens the juvenile 79 period resulting in premature metamorphosis, smaller pupae and pupal lethality (5). Additionally, 80 increased JH signaling is also detrimental to development. Increased or prolonged JH extends 81 the juvenile period, leading to larger animals and either a delayed or absent metamorphosis (6). 82 Indeed, artificially increasing JH signaling is the basis for JH-based insect growth regulators used 83 around the world. Together, these observations highlight the importance of balancing the 84 synthesis and degradation of JH in development. 85 86 While our understanding of JH synthesis enzymes is increasing, JH degradation enzymes are 87 poorly understood. Insects and crustaceans have two major classes of enzymes that 88 endogenously degrade JH: JH esterases (JHEs) and JH epoxide hydrolases (JHEHs) (Fig 1a). 89 .CC-BY-NC 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted June 12, 2025. ; https://doi.org/10.1101/2025.06.09.657647doi: bioRxiv preprint 3 These two classes of enzymes differ by structure, mechanism, reversibility, and location of action. 90 JHEs function as monomers extracellularly, often within the circulatory system, and inactivate JHs 91 by removing the methyl group from the carboxylic ester to form JH acid (Fig 1a) (7). JHE-92 dependent demethylation is presumed to be reversible by Jhamt (7). JHEHs are intracellular 93 dimers, that irreversibly degrade JHs by hydrolyzing the epoxide groups of JHs or JH acid to 94 generate JH-diol or JH acid-diol (8). While the biochemistry of JHEs and JHEHs has been 95 examined in vitro, the roles and contributions of these classes of degradation enzymes in vivo is 96 poorly understood, in part due to genetic redundancy and compensatory feedback (5, 9, 10). 97 98 The Drosophila melanogaster genome encodes two JH esterases, Jhe and JheDup, and three JH 99 epoxide hydrolases, Jheh1, Jheh2, and Jheh3. The sequence homology for these paralogs 100 ranges between 71-77% similarity and 41-46% identity (Fig 1b). Yet, their predicted tertiary 101 structures, binding pockets, and catalytic amino acids are nearly identical (Fig 1b and c, Fig S1). 102 Paralogs are positioned in tandem within the genome and share similar spatiotemporal 103 expression (Fig 1d). However, JHE and JHEH genes are expressed in largely distinct 104 spatiotemporal patterns except in the larval ring gland (11, 12). Recent work has shown 105 neofunctionalization of JheDup in olfaction and requirements of Jheh 1 and Jheh2 in glucose 106 metabolism (13, 14). However, much remains to be understood about whether each class of 107 enzymes has shared or unique requirements in development. 108 109 Here, we wholistically define the developmental requirements of JHE and JHEH enzymes in two 110 classic JH functions: regulating growth and developmental timing. We generated double 111 jhe,jheDup knockout (JHE DKO) animals and triple jheh1,2,3 knockout (JHEH TKO) Drosophila 112 melanogaster strains. Using these models, we found that loss of each enzyme class differentially 113 affected developmental timing and growth such that JHEs are required to restrain body size, 114 whereas JHEHs are required for normal developmental timing in Drosophila. Genetic rescue, 115 epistasis, and transcriptomic analyses revealed that JHEs and JHEHs have shared requirements 116 in regulation of Cytochrome P450 gene expression and ring gland size, while JHEHs have unique 117 requirements in the regulation of the ecdysone axis. Together, our findings provide the first report 118 of the shared and unique requirements for JH degradation enzymes in Drosophila development, 119 shedding light on the diversity of mechanisms that control hormone dynamics and homeostasis in 120 development. 121 122 123

124 125 The unique and shared developmental requirements for JH degradation enzymes are unknown. 126 Since the genes encoding each class of JH degradation enzyme have distinct spatiotemporal 127 expression profiles, we hypothesized that each class of JH degradation enzyme has distinct 128 requirements in regulating JH levels and development. To test this hypothesis, we generated 129 deletions in Jhe, JheDup and jheh1, jheh2, jheh3 by CRISPR gene editing (Fig 2a). We screened 130 founders by PCR using primers flanking desired cut sites and recovered five lines lacking jhe and 131 jheDup originating from a single founder, which we will be referred to as DKO, for JHE double 132 knockout. We also generated three lines lacking jheh1, jheh2, jheh3 from three separate 133 founders, which will be referred to as TKO for JHEH triple knockout. Each line was sequenced for 134 precise breakpoints and analyzed in silico and in all cases edited genomes do not encode 135 functional Jhe and JheDup, or Jheh1, Jheh2, and Jheh3. 136 137 To define any requirements in development, we first assessed the viability of animals lacking 138 JHEs or JHEHs. To avoid confounding effects of second site mutants, we crossed our newly 139 generated JHE DKOs to a deficiency line lacking both Jhe and JheDup. As at the time of these 140 studies a deficiency line lacking all three JHEHs was not available, we crossed two newly 141 generated JHEH TKO lines from two independent founders to each other. Under non-crowding 142 laboratory conditions, an expected percentage of heterozygous and homozygous mutant adult 143 .CC-BY-NC 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted June 12, 2025. ; https://doi.org/10.1101/2025.06.09.657647doi: bioRxiv preprint 4 progeny eclosed into adults (Fig S2a). To test whether maternally provided JHEs or JHEHs 144 contribute to embryonic development, we generated embryos lacking maternal and zygotic JHE 145 or JHEHs and found that the percentage of embryos lacking either class of JH degradation was 146 comparable to wild type levels (Fig S2b). Together, these data indicate that neither JHEs nor 147 JHEHs are required for viability in Drosophila melanogaster. 148 149 JHEs and JHEHs have opposing requirements in developmental growth 150 151 In many insects, elevated JH titers coincide with larger body size (6, 15-17). Thus, we predicted 152 that loss of JH degradation enzymes would increase Drosophila body size. To test this prediction, 153 we measured pupal length in DKO, TKO and control animals. As Drosophila size is highly 154 sensitive to animal density during development, we transferred 25 embryos to individual vials (18, 155 19). We found that JHEH TKO animals were smaller, and JHE DKO animals were larger than wild 156 type controls across both sexes (Fig 2b,c). To determine whether differences in pupal length were 157 due to loss of JHEs or JHEHs, we generated JHE DKO and JHEH TKO animals that carried 158 exogenous Bacterial Artificial Chromosomes (BACs) containing either both jhe, jheDup loci, or all 159 three jheh1, 2, 3 genes. Because these BAC rescue animals were white+ and white- flies carry 160 numerous defects (20), we included a second control, OregonR crossed to w1118 (OreRx w1118), 161 for this and all subsequent experiments. We found that JHEH TKO animals carrying a BAC with 162 jheh1,2,3 genes were significantly larger than JHEH TKOs and not statistically different from wild 163 type controls (Fig 2c). We noted that while JHE DKO pupae were larger, JHE DKO carrying a 164 BAC with jhe, jheDup genes partially rescued the larger size of DKO females but not males (Fig 165 2c). To further explore whether the larger size of JHE DKO animals was related to a dysregulated 166 JH axis, we conducted an epistasis experiment. We reasoned that if increased growth was due to 167 increased JH titers, then this phenotype might be altered by reducing JH synthesis. To reduce JH 168 synthesis, we incorporated loss-of-function mutations in jhamt, a key JH synthesis gene, into the 169 JHE DKO and JHEH TKO background and measured pupal length. We found that pupal length of 170 jhamt, Jhe, JheDup triple homozygous mutant animals was restored to wild type levels in both 171 sexes, supporting the prediction that the larger body size of JHE DKO animals results from 172 increased JH signaling (Fig 2d). Together, these data demonstrate that JHEs and JHEHs have 173 opposite requirements in developmental growth. 174 175 JHEHs, but not JHEs, are required for normal developmental timing: 176 177 Over the course of our viability and size studies, we noted that homozygous JHEH TKO animals 178 eclosed much later than heterozygous siblings (Fig S3). Elevated JH titers are predicted to 179 extend the juvenile period of development (6). Thus, we quantified the time it took DKO, TKO, 180 and control animals to pupariate. To control the effect of animal density on developmental timing 181 (21), we transferred 75 embryos laid over a four-hour time period to individual wide vials and 182 quantified pupae every six hours until five days after the first adult progeny eclosed to avoid 183 quantifying the F2 pupae. JHE DKO displayed no delay in pupariation (Fig 3a). In contrast, JHEH 184 TKO took seven days longer to pupariate (Fig 3b), however in subsequent experiments in larger 185 incubators which held temperature longer, JHEH TKOs took an average of four days longer to 186 pupariate (Fig 3c and Fig 5a). To determine if any observed prolonged juvenile stages were due 187 to loss of JHEH genes, second site mutations, or a gain-of-function phenotype from the intact 188 jheh 3 promoter (Fig 2a), we measured pupariation timing in JHEH TKO animals carrying an 189 exogenous BAC with all three jheh1, 2, 3 genes. We found that JHEH TKO, BAC [jheh1,2,3] 190 animals pupariated at the same time as wild type controls, indicating that the prolonged juvenile 191 .CC-BY-NC 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted June 12, 2025. ; https://doi.org/10.1101/2025.06.09.657647doi: bioRxiv preprint 5 stage is due to loss of JHEHs (Fig 3b). Together, these data suggest that Drosophila 192 melanogaster JHEHs are required to maintain normal developmental timing. 193 194 Transcriptomics reveals both shared and unique requirements for JHEs and JHEHs 195 196 Our data thus far indicate that JHEs and JHEHs have unique requirements in juvenile 197 developmental growth and timing. While these phenotypes are consistent with dysregulated JH 198 signaling, incorporation of jhamt mutations did not rescue the smaller body size nor prolonged 199 development in JHEH TKO animals (Fig 2c and Fig 3c), suggesting JHEH may have additional 200 requirements. To understand the nature of these multiple JHE and JHEH developmental 201 requirements, we profiled transcriptomes of JHE DKO, JHEH TKO and control males and females 202 by bulk RNA sequencing. Given the dramatic developmental delay of JHEH TKOs, we isolated 203 RNA from animals in the highly dynamic but 15 minute-long white prepupae stage to ensure 204 comparison of equivalent developmental stages (22). Principal component analyses of three 205 biological replicates per sex per genotype highlighted a high degree of similarity within JHE DKO 206 and JHEH TKO biological replicates, both of which clustered separately from wild type samples 207 (Fig S4a). The number of differentially expressed genes (DEGs) ranged from 699 to 1,841, with 208 more DEGs in females than males among both JH degradation class mutants (Fig 4a, Datasets 209 S1 and S2). Between JHEH TKO and JHE DKO animals, JHEH TKO animals of both sexes had 210 more DEGs, consistent with their more severe developmental phenotypes (Fig 4a). 211 212 We next performed gene ontology analysis to gain insights into processes dysregulated in JHE 213 DKO and JHEH TKO animals. Comparison of GO terms revealed shared gene regulation 214 requirements between JHE and JHEH enzyme classes, which was more prominent in females. 215 Indeed, 29% of DEGs in JHE DKO white prepupae are shared with JHEH TKO females (Fig 4b). 216 Among shared upregulated DEGs, we found a significant enrichment for genes encoding 217 Cytochrome P450s (Cyp450s) (Fig 4c). Also shared was upregulation of genes expressed in fat 218 body and yolk nuclei genes, tissues regulated by JH (Fig S4c, (23)), as well as metabolism genes 219 (Dataset S3). Together, these findings indicate that despite unique requirements in 220 developmental growth and timing, JHE and JHEH have shared transcriptional signatures. 221 222 The gene ontology analysis also revealed unique DEG signatures (Dataset S3). Downregulated 223 DEGs in JHE DKO males were significantly enriched for spermatogenesis-related genes (Fig 224 S4e). In contrast, downregulated DEGs in JHEH TKO animals of both sexes were significantly 225 enriched for iron binding, small molecule transport, metabolism, seminal vesicle biology, 226 andribosomal protein genes (Fig S4d). Additional Cyp450s were also significantly enriched in 227 JHEH TKO DEGs, as twelve Cyp450s were uniquely upregulated (Fig 4c). Thus, together with 228 the ten Cyp450s upregulated in both classes of JH degradation mutants, 22 out of the 83 total 229 Cyp450s genes in the Drosophila genome are dysregulated in JHEH TKO animals. Together, 230 these transcriptomic results support our phenotypic findings that together indicate JHEH and JHE 231 enzymes have shared and unique requirements in Drosophila development. 232 233 Our transcriptomic data hinted at the activation of compensatory feedback mechanisms, 234 especially upon the loss of JHEHs. Due to the serendipitous fact that the JHEH3 promoter is 235 intact in our JHEH TKO animals (Fig 2a), we found a 13-fold upregulation of transcriptions from 236 the intact JHEH3 promoter (Fig 4d), suggesting feedback-driven autoregulation of JHEHs in vivo. 237 We did not detect upregulation of Kr-h1 in neither JHE DKO nor JHEH TKO animals, expected 238 because we matched mutant samples to metamorphosis onset rather than hours post egg lay. 239 Indeed, JH titers must fall for ecdysone to initiate pupariation (24-27). However, we found that 240 several ecdysone-responsive genes were dysregulated in both JHE DKO and JHEH TKO 241 animals, although the directionality of change differed depending on JH degradation class, sex, 242 and gene (Fig S4b). Further investigation into genes within the ecdysone axis revealed a 243 significant upregulation of several ecdysone synthesis genes in JHEH TKO animals (Fig 4c). 244 .CC-BY-NC 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted June 12, 2025. ; https://doi.org/10.1101/2025.06.09.657647doi: bioRxiv preprint 6 Together, these data indicate ecdysone-related factors are significantly dysregulated, especially 245 in the absence of JHEHs. 246 247 Developmental timing requirements for JHEH are linked to ecdysone signaling 248 249 Our transcriptomic analysis suggested that ecdysone signaling may be dysregulated in JHEH 250 TKO animals. To test whether delayed pupariation is due to dysregulated ecdysone signaling, we 251 fed wild type and JHEH TKO larvae the active form of Ecdysone, 20E, and measured time to 252 pupariation. Strikingly, feeding exogenous 20E significantly rescued delayed pupariation in JHEH 253 TKO animals (Fig 5a). These data suggest that delayed pupariation is linked to a lack of sufficient 254 ecdysone in JHEH TKO animals despite increased expression of ecdysone synthesis genes. 255 Thus, we analyzed the morphology of the ecdysone-producing prothoracic gland (PG), which 256 contributes the majority of the volume of the larval ring gland (RG) (28). We dissected RGs from 257 wild type, JHE DKO, and JHEH TKO wandering third instar larvae and stained them for DNA (Fig 258 5b). Quantification of RG volume revealed both JHE DKO and JHEH TKO RGs were significantly 259 larger than wild type controls despite opposing requirements in developmental growth (Fig 5c). 260 Ecdysone synthetic capacity scales with the number of endocycles in the prothoracic gland (29). 261 To estimate endocycling status, we measured total PG nuclei volume using DAPI signal and 262 found DAPI volume per PG was either the same or significantly decreased in JHE DKO and 263 JHEH TKO animals of both sexes (Fig 5d). These results suggest that increases in RG size in 264 both JHE DKO and JHEH TKO animals is likely due to defects other than an increase in 265 endocycling. Altogether, our transcriptomic, rescue, and RG morphological data support our 266 phenotypic findings indicate JHEH and JHE enzymes have shared and unique requirements in 267 Drosophila development (Fig 5e). 268 269 270

271 Developmental timing, growth, and reproduction require precise spatiotemporal regulation of 272 hormones through the balance of synthesis and degradation. Here, we use JHs in Drosophila 273 melanogaster as a model to understand the developmental requirements for programmed 274 hormone degradation in vivo. By generating strains lacking both JHE enzymes or all three JHEH 275 enzymes, we found these distinct JH degradation pathways have separate contributions to 276 developmental growth and timing yet have shared requirements in regulating prothoracic gland 277 size and Cyp450 gene expression. Together, these efforts provide the first steps toward better 278 understanding of the complexity of programmed hormone degradation. 279 280 JH signaling promotes juvenile growth and developmental timing. We find that these two classic 281 JH functions are differentially regulated by JHE and JHEH degradation enzymes. JHEs restrain 282 body size, while JHEHs restrain the juvenile period (Fig 2 and Fig 3). These results are consistent 283 with previous studies showing that JHEs restrainsize. 284 285 JHE knockdown causes increase in cell size of adult intestinal stem cells in Drosophila (30) and 286 body size in Bombyx mori (31, 32) while overexpression of JHE has been used as a pesticide in 287 Heliothis virescens and leads to smaller body size (7). Indeed, jheh knockdown leads to 288 decreased body size and longer developmental time in Colorado potato beetle, Leptinotarsa 289 decemlineata (33) and RNAi of jheh induces the short wing morph in brown planthoppers (34). 290 Several scenarios could lead to separation of developmental requirements. Each enzyme class 291 might have JH-dependent and JH-independent functions. While JHEs and JHEHs are both 292 capable of degrading JHs, these enzymes may act on additional, unshared substrates (14, 35, 293 36). Separate requirements could reflect differences in the reversibility of JH degradation, as 294 JHE-dependent demethylation can be reversed by Jhamt activity (7). Alternatively, the 295 degradation products made by each type of enzyme may have unique biological activities that 296 contribute to either developmental growth or timing. Indeed, JHEs produce JH acids which are 297 proposed to have hormone functions (37). Finally, these enzymes might be active at different 298 .CC-BY-NC 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted June 12, 2025. ; https://doi.org/10.1101/2025.06.09.657647doi: bioRxiv preprint 7 times or tissues, consistent with differential spatiotemporal gene expression (Fig 1). While our 299 study is a first wholistic step in defining developmental requirements for JHE and JHEH enzymes, 300 additional biochemical and developmental studies will be needed to explore each of these 301 possibilities. 302 303 Like many hormones, the JH axis possesses genetic redundancies and compensatory feedback. 304 Here, we overcame genetic redundancies by generating double JHE and triple JHEH knockout 305 animals, which revealed developmental requirements not found by loss of individual enzymes. 306 While Jheh1 and Jheh2 loss leads to a four-hour delay in pupariation (13), loss of all three JHEHs 307 causes a four-to-seven-day delay in pupariation (Fig 3). Together, these data indicate that either 308 that Jheh3 is the most critical JHEH to regulate developmental timing or that loss of Jheh1 and 309 Jheh2 leads to compensatory upregulation of Jheh3. We favor the latter, as we serendipitously 310 found an autoregulation mechanism at the level of JHEH transcription (Fig 4), adding to the 311 several examples of compensatory feedback in the JH axis (5, 9, 10, 38-40). A recent study found 312 compensatory feedback between the two classes of JH degradation enzymes (41). While we did 313 not find evidence for JHE to JHEH cross-regulation, this may reflect the fact that our dataset 314 focuses on a very narrow developmental stage. Consistent with the existence of cross-regulation, 315 we failed to generate animals lacking all five degradation enzymes, suggesting that even 316 heterozygous loss of all JH degradation enzymes significantly compromises development. Future 317 studies will be needed to determine whether compensatory feedback exists within JHEs or 318 between each JH degradation class and whether lack of feedback between JHE and JHEHs in 319 Drosophila reflect functional specialization. 320 321 Our transcriptomic data provide important clues about these unique functions. JHE DKO white 322 prepupae exhibited a downregulation of spermatogenesis genes, suggesting JHE may function in 323 male reproduction consistent with several recent reports implicating JH signaling in male 324 reproduction (42-46). In contrast, JHEH TKO animals exhibited an upregulation of five ecdysone 325 biosynthesis enzymes (Fig 4). Upregulation of ecdysone synthesis genes could be the result of 326 enlarged prothoracic glands. However, prothoracic glands of JHE DKO animals were larger yet 327 only had one upregulated ecdysone synthesis gene and the ecdysone synthesis gene expressed 328 in the fat body, shade, is also upregulated in JHEH TKO animals (Figs 4 and 5). Alternatively, 329 both ring gland enlargement and increased expression of ecdysone synthesis genes could reflect 330 global activation of homeostatic feedback to counteract elevated JH titers. This postulate is 331 supported by the absence of JH inducible gene misexpression (Fig S4). Our observation that 332 ectopic 20E rescues delayed pupariation despite increased expression of ecdysone synthesis 333 genes is indicative of compensatory program to counteract elevated JH titers. The mechanisms of 334 this feedback loop and whether a similar loop exists for JHE-mediated degradation will be an 335 exciting area of future exploration. 336 337 Dysregulated ecdysone signaling alone cannot account for all JHEH requirements in Drosophila 338 development. Indeed, we noted several defects. 20E-rescued JHEH TKO animals were still 339 delayed approximately 30 hours relative to wild type animals (Fig 5), pupae were significantly 340 smaller (Fig S5), and unlike 20E-fed wild type or vehicle control JHEH TKO animals, only 50% of 341 20E-fed JHEH TKO animals eclosed (Fig S5). Slower development and smaller body size in 342 JHEH TKO animals could be due to increased expression of lipid signaling and metabolism 343 genes (Dataset S3). In Drosophila, lipid metabolism is required for proper developmental timing 344 (47). In C. bowringi and H. axyridis, JHEHs regulate lipid storage (48, 49). Consistently, loss of 345 the lipid binding protein CG9186/Strukopf phenocopies JHEH TKO mutants and physically 346 interacts with all three JHEHs (50). Jheh1 and 2 were recently shown to regulate glucose 347 metabolism (13), thus one unique function of JHEHs may be to regulate the balance between 348 glucose and lipid metabolism independent of JH. While functions in metabolism may underlie our 349 observations that JHEH TKO are smaller despite prolonged juvenile development, we noted that 350 ribosomal protein genes are highly enriched among JHEH TKO DEGs (Fig S4d). Based on 351 findings that JH signaling promotes ribosomal gene expression in the mosquito (23), we postulate 352 .CC-BY-NC 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted June 12, 2025. ; https://doi.org/10.1101/2025.06.09.657647doi: bioRxiv preprint 8 that reduction of ribosomal proteins in JHEH TKO restricts animal growth and cannot be 353 compensated by increased developmental time (51, 52). Together, these findings suggest that 354 regulation of ribosomal protein genes may be a conserved feature of JH signaling. 355 356 Despite unique requirements in juvenile growth and developmental timing, our studies also 357 suggest JHE and JHEHs have shared functions. Loss of either class of degradation enzymes 358 significantly altered expression of Cyp450s (Fig 4). Cyp450s are a diverse class of genes 359 involved in the synthesis of hormones and fatty acids, as well as the metabolism and 360 detoxification of many small molecules. Indeed, several insecticides induce Cyp450 expression 361 (53). Consistent with findings in termites in which ectopic JH III induces Cyp450 expression (54), 362 our results suggest increased endogenous JHs also induce Cyp450 expression (Fig 4). 363 Upregulation of Cyp450s could facilitate JH degradation in the absence of JHEHs or JHEs. For 364 instance,Cyp4C7 was shown to convert JH III to 12-trans-hydroxy JH III and expression of this 365 Cyp450 was induced after bursts of JH synthesis (55). The mechanistic and functional 366 relationship between JH levels and Cyp450s will be important to elucidate in future studies. 367 368 Here, we defined the role of JH degradation enzymes in classic JH-functions: developmental 369 timing and growth. However, JHs also have essential roles in reproduction, diapause, and sexual 370 dimorphic structures and behaviors (56-59). The importance of JH degradation in controlling 371 these post-juvenile functions is currently unclear. JH mimics are used around the world as insect 372 growth regulators for pest control (60-62). The expression JH degradation enzymes are induced 373 upon application of JH mimics (63-65). The extent to which JH degradation enzymes and 374 associated pathways are involved in resistance to JH mimics remains unknown. Our findings on 375 programmed JH degradation in the genetically tractable Drosophila melanogaster model provide 376 a platform to shed light on the mechanisms that control hormone dynamics and homeostasis in 377 development and beyond. 378 379 380

381 382 Animal rearing: 383 Stocks used in this study are listed in Supplemental Table 1 were reared on a molasses/cornmeal 384 medium with methyl 4-hydroxybenzoate (Tegosept) as a mold inhibitor. Embryo collections were 385 conducted using cages with removable apple juice plates. The apple juice plates were made of 386 25% apple juice, 2.5% sucrose, 2.25% Bacto-agar, and 0.15% Tegosept. Experiments were 387 carried out at 25°C, 70% humidity with a 12-hour day-night cycle unless otherwise specified. 388 389 Predicted Structures 390 Jhe and JheDup (Uniprot IDs: A1ZA98 and A1ZA97 respectively) predicted structures were 391 retrieved from the AlphaFold Protein Database and created using the AlphaFold Monomer v2.0 392 pipeline. Jheh1,2,3 (Uniprot IDs: Q7JRC3, Q7KB18, Q7K1W4 respectively) predicted dimer 393 structures were retrieved from the SWISS-MODEL repository and were created using ProMod3. 394 395 Generation of double and triple JH degradation gene knock out animals: 396 JHE DKO and JHEH TKO animals were generated by non-homologous end joining of CRISPR-397 mediated breakpoints flanking jhe and jheh loci. Guide RNAs were designed based on 398 sequencing relevant PAM sites in the vas-Cas9 injection stock (Bloomington Stock Center 399 #51323). For each knockout, two guides were injected per PAM site for a total of four injected 400 guides each for jhe,jheDup and jheh1,2,3 deletion (Supplemental Table 2). Founders were 401 screened by PCR detection of small products generated by primers flanking predicted cut sites. 402 For jheh 1,2,3 loci, six promising founders were identified. For jhe,jheDup loci, eight promising 403 founders were identified. For each promising founder, twelve F1 crosses to Sp,hs-hid/CyO, ft-lacZ 404 were set up and desired deletions were screened by PCR with genomic DNA templates isolated 405 .CC-BY-NC 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted June 12, 2025. ; https://doi.org/10.1101/2025.06.09.657647doi: bioRxiv preprint 9 by QuickExtract using primers listed in Supplemental Table 2. Precise breakpoints were then 406 identified by Sanger sequencing using the same primers used to screen for deletions. 407 408 For sequence-verified JHEH TKO lines 1-6-2; 1-18-8; and 1-19-4, the large deletion encodes a 409 frameshift starting at amino acid 53 of Jheh3, upstream of the hydrolase domain which starts at 410 amino acid 154. Beyond amino acid 53 of Jheh3, peptides of varying lengths (25 for 1-6-2; 93 for 411 1-18-8; and 193 for 1-19-4) are possible but do not match any Jheh sequence. For sequence-412 verified JHE DKO lines 2-1-1; 2-1-3; 2-1-5; 2-1-9; and 2-1-12, the large deletion encodes a 413 frameshift starting at amino acid 62 of JheDup, downstream of the carboxylesterase domain 414 starting at amino acid 30 but upstream of the acetyl esterase/lipase domain at 103 and upstream 415 of the catalytic triad at amino acid 207, 342 and 466. For all lines, there is a potential additional 416 15 amino acids not matching any Jhe sequence. 417 418 Percent expected class: 419 The percent expected class of zygotic JHE DKO and JHEH TKO mutants was calculated as done 420 previously (66). Briefly, three virgin heterozygous, balanced females were crossed to two 421 heterozygous, balanced males for three days for each of three biological replicates. To allow for 422 sufficient time to eclose, balanced and unbalanced progeny were counted for up to twenty days 423 after the cross was set up. 424 425 Hatch rate: 426 To determine the hatch rate of maternally and zygotically JH degradation mutants, less than 3-427 day old homozygous JHE DKO and JHEH TKO virgin females were crossed to homozygous 428 mutant males in vials for 24 hours alongside control crosses. Then, parents were transferred to 429 egg collection cages for 24 hours before collecting embryos every 24 hours thereafter. Each plate 430 represented one out of three biological replicates. Embryos/eggs were counted upon plate 431 collection and then incubated at 25°C, 70% relative humidity for 36 hours, afterwhich the number 432 of unhatched embryos was counted. 433 434 Developmental timing assays: 435 Wild type and JHE(H) KO/CyO, Tubby1::RFP virgin female flies were collected for no more than 3 436 days before setting up the mating cross, and male flies were also collected no more than 24 437 hours before the crosses were set. Parental ages were controlled to mitigate any effects of age 438 on progeny development. All genetic crosses were set in vials for 24 hours before transferring to 439 cages with apple juice plates to ensure mating occurred prior to collection of the progeny. The 440 flies were kept in cages for 24 hours to adjust the flies to the collection process after which, fresh 441 cage plates were collected every 6 hours to synchronize the vials and set the time resolution of 442 the experiment. As larval density is known to impact pupariation timing in Drosophila (19), 75 443 embryos were placed in each vial for all control and experimental conditions. To prevent larval 444 starvation from being unable to break the surface of the food, the food in each vial was mashed 445 before embryos were placed inside (18). Starting on day three after embryo transfer, vials were 446 checked every 6 hours, and the number of Tubby and non-Tubby pupa were counted. Pupa were 447 only counted when the prepupal case had formed (P0 or later), and the center of each pupa was 448 marked on the outside of the vial to prevent recounting of the same pupa. Data was collected for 449 12 days, after which remaining larva were assumed to be from the next (F2) generation. 450 451 Pupal and adult tissue size measurements: 452 For pupal and adult size experiments, controls and mutant genotypes were crossed in vials for 24 453 hours and transferred to cages. 25 embryos were placed into each wide vial to prevent 454 overcrowding from affecting larval size. The food in the vials was mashed to allow the larva to 455 feed even in low density conditions. After the larva developed, the pharate adult pupa were sexed 456 by the presence of sex combs and separated. Images of each genotype and sex were taken 457 using the Nikon Digital Sight 1000 mounted to a Nikon SMZ1500 trinocular microscope at 75X 458 .CC-BY-NC 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted June 12, 2025. ; https://doi.org/10.1101/2025.06.09.657647doi: bioRxiv preprint 10 magnification. The length of the pupa was measured in ImageJ using a reference ruler in the 459 image. 460 461 Jhamt epistasis experiment: 462 To generate jhamt, jhe, jheDup triple mutants and jhamt, jheh1,2,3 quadruple mutants, we first 463 crossed jhamt8F2 or jhamtDef (38) to JHEH TKO or JHE DKO flies (this paper). Resulting F1 464 virgin females heterozygous for jhamt and JHE(H) KO were crossed to hs-hid, Sp/CyO,ft-lacZ 465 males. Single male F2 CyO, ft-lacZ progeny were mated to females carrying a CyO, Tubby1::RFP 466 balancer for 4 days and then collected for genotyping by PCR for jhamt mutation, as well as 467 jheh1,2,3 or jhe, jheDup deletions. DNA was extracted using QuickExtract DNA Extraction 468 Solution (Biosearch Techologies QE0905T) and jhamt, jheh1,2,3, jhe, jheDup loci were amplified 469 by PCR (Denville Scientific C788T64) To detect the presence of jhamt mutations, PCR products 470 were purified using a DNA gel extraction kit (NEB T1020) and sanger sequenced (Azenta). A lack 471 of PCR product was used to detect the presence of jheh1,2,3 or jhe, jheDup deletions. 472 Developmental timing and pupal length were measured as described above. 473 474 RNA isolation, library preparation and sequencing: 475 To avoid overcrowding, control and heteroallelic mutant crosses were set up in embryo collection 476 cages. Seventy-five embryos were transferred to wide vials to prevent overcrowding. White 477 prepupa were collected from control and mutant genotype vials, which allowed for the pupa to be 478 synchronized to within a 15-minute window of development. As prepupae were collected, they 479 were homogenized in TRIzol (Invitrogen, ThermoFisher #15596018) and stored at -80°C for later 480 RNA extraction. RNA was isolated from three biological replicates, each containing five white 481 prepupa, by TRIzol extraction followed by isopropanol precipitation. Seven micrograms of total 482 RNA from each replicate were DNaseI-treated and cleaned up using Monarch Total RNA 483 Miniprep Kit (NEB T2010S). One microgram of total RNA was used for RNA-Seq library 484 preparation, using NEBNext Ultra II Directional RNA Library Prep Kit for Illumina (NEB E7760) 485 with NEBNext Poly(A) mRNA Magnetic Isolation Module (NEB E7490). Samples were indexed 486 with unique dual index primer pairs from NEBNext Multiplex kit (NEB E6440) and processed 487 according to the manufacturers’ instructions, with 8 cycles of amplification. Individual libraries 488 were sized and quantified using Tapestation 4200 and High Sensitivity D1000 ScreenTape 489 (Agilent 5067-5584), and Qubit fluorometer with dsDNA HS Assay kit (Thermo Q32854). 490 Equimolar amounts of individual libraries were combined in one pool and sequenced by a 491 commercial provider (Azenta Inc.) using 150 bp paired-end chemistry on NovaSeq X Plus with 492 5% phiX spike-in. 493 494 RNA sequencing analyses: 495 Demultiplexed .fastq files were aligned to BDGP dm6 using HISAT v 2.2.1 (67), converted to 496 .bam, sorted and indexed using SAMtools 1.21 view, sort and index commands with default flags 497 (68), and visualized as .bigwig using deepTools 3.5.6 bamCoverage “--binSize 1” flag (69). Read 498 quality and alignment statistics were assessed with FastQC and multiQC (70). Read counts were 499 summarized from sorted .bam files using HTSeq 2.0.3 “htseq-count” command using BDGP 500 dm6.46 113 release .gtf reference (71). Differentially expressed genes were identified using 501 DESeq 2 (72), with thresholds of fold change >1.5 and p value <0.05 adjusted for multiple testing 502 using Benjamini-Hochberg correction. Raw reads, counts, and .bw tracks are deposited to Gene 503 Expression Omnibus under accession number GSE286242. The functional enrichment analysis 504 was performed using g:Profiler (version e111_eg58_p18_f463989d) with g:SCS multiple testing 505 correction method applying significance threshold of 0.05 (73). 506 507 Immunohistochemistry: 508 Wild type and mutant wandering third instar larva were collected from 6 replicate vials seeded 509 with 75 embryos each as described elsewhere. Male and female ring glands dissected according 510 to (74). Briefly, the anterior third of the larvae were dissected and inverted, then fixed in 0.4% 511 PFA for 30 minutes. Samples were then stained with DAPI and mouse anti-1b1 (1:20, AB528070, 512 .CC-BY-NC 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted June 12, 2025. ; https://doi.org/10.1101/2025.06.09.657647doi: bioRxiv preprint 11 Developmental Studies Hybridoma Bank) to visualize the nucleus and cell membrane then 513 dissected further to get the CNS and Ring Gland. Images were collected using a Dragonfly 200 514 spinning disk confocal microscope (Andor/Oxford Instruments) at 20x magnification and two-515 micron step size. Images were analyzed for ring gland volume and the volume of prothoracic 516 gland cell nuclei using IMARIS. Two machine learning models were trained on control images, 517 one to identify ring glands and another for PG nuclei. Surfaces were generated using these 518 models with manual refinement, and the volume of ring gland and prothoracic gland nuclei were 519 obtained. 520 521 Ecdysone rescue experiments 522 Wild type and JHEH TKO embryos were collected within four-hour time windows for synchronous 523 development. Animals were then left on apple juice plates supplemented with ample mashed 524 normal food for 72 hours. After 72 hours, fifty larvae were transferred to each vial, containing 525 either 0.5mg/g of 20-Hydroxyecdysone (20E, dissolved in 100% Ethanol, Sigma Cat #H5142) or 526 the equivalent amount of 100% ethanol. All experiments contained three biological replicates. To 527 measure time to pupariation, pupae were quantified every 12 hours until five days after the first 528 two adults eclosed to avoid quantifying the next generation. To measure eclosion rates, we 529 separated pupae into 96-well plates and quantified adult animals after 10 days. Pupal length was 530 measures as described elsewhere in these Methods. 531 532 Statistics 533 For the developmental timing experiments, the time at which 50% of the larva had pupariated was 534 found using linear interpolation between the nearest time points. Normality and variance of 535 samples were assessed using Shapiro and Levine tests in R, t tests were used when data was 536 parametric and a Mann–Whitney U test was used when data was non-parametric, as noted in the 537 figure legends. 538 539 540 Data and Resource Availability 541 542 All relevant data and resources generated from this study can be found within the article and its 543 supplemental material. RNA-Seq data have been uploaded to the Gene Expression Omnibus 544 database (Accession Number GSE286242). 545 546 547 Acknowledgments 548 549 We thank Dr. Brian Brigham for statistics consultation and members of the Barton Lab and Dr. 550 Pamela Geyer for feedback. We thank Dr. Ruth Lehmann for advice and generous support at the 551 beginning of this study. The mouse anti-1B1 antibody developed by Dr. Howard Lipshitz was 552 obtained from the Developmental Studies Hybridoma Bank, created by the NICHD of the NIH and 553 maintained at The University of Iowa, Department of Biology, Iowa City, IA 52242. Stocks 554 obtained from the Bloomington Drosophila Stock Center (NIH P40OD018537) were used in this 555 study. Gene function obtained from FlyBase (NSF 2039324). This work was supported by R00 556 HD097306 from the Eunice Kennedy Shriver National Institute for Child Health and Development 557 to LJB. This research was conducted while Lacy J Barton was an AFAR Grant for Junior Faculty 558 awardee. Lacy Barton, PhD holds a Voelcker Fund Young Investigator Award from the MAX AND 559 MINNIE TOMERLIN VOELCKER FUND. 560 561 562 563 564 565 566 .CC-BY-NC 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted June 12, 2025. ; https://doi.org/10.1101/2025.06.09.657647doi: bioRxiv preprint 12

567 568 1. Q. Jia et al., Cyp6g2 is the major P450 epoxidase responsible for juvenile hormone 569 biosynthesis in Drosophila melanogaster. BMC Biol 22, 111 (2024). 570 2. R. Niwa et al., Juvenile hormone acid O-methyltransferase in Drosophila melanogaster. 571 Insect Biochem Mol Biol 38, 714-720 (2008). 572 3. V. B. Wigglesworth, Secretion of Juvenile Hormone by the Corpus Allatum of Calliphora. 573 Nature 174, 556 (1954). 574 4. M. Jindra, M. Uhlirova, J. P. Charles, V. Smykal, R. J. Hill, Genetic Evidence for Function 575 of the bHLH-PAS Protein Gce/Met As a Juvenile Hormone Receptor. PLoS Genet 11, 576 e1005394 (2015). 577 5. M. A. Abdou et al., Drosophila Met and Gce are partially redundant in transducing 578 juvenile hormone action. Insect Biochem Mol Biol 41, 938-945 (2011). 579 6. T. Zhang et al., Kruppel homolog 1 represses insect ecdysone biosynthesis by directly 580 inhibiting the transcription of steroidogenic enzymes. Proc Natl Acad Sci U S A 115, 581 3960-3965 (2018). 582 7. S. G. Kamita, B. D. Hammock, Juvenile hormone esterase: biochemistry and structure. J 583 Pestic Sci 35, 265-274 (2010). 584 8. D. Borovsky et al., Cloning and Characterization of Drosophila melanogaster Juvenile 585 Hormone Epoxide Hydrolases (JHEH) and Their Promoters. Biomolecules 12 (2022). 586 9. A. C. LeBoeuf et al., Molecular evolution of juvenile hormone esterase-like proteins in a 587 socially exchanged fluid. Sci Rep 8, 17830 (2018). 588 10. V. Smykal, D. Dolezel, Evolution of proteins involved in the final steps of juvenile 589 hormone synthesis. J Insect Physiol 145, 104487 (2023). 590 11. B. Cho et al., Single-cell transcriptome maps of myeloid blood cell lineages in Drosophila. 591 Nat Commun 11, 4483 (2020). 592 12. D. Christesen et al., Transcriptome Analysis of Drosophila melanogaster Third Instar 593 Larval Ring Glands Points to Novel Functions and Uncovers a Cytochrome p450 594 Required for Development. G3 (Bethesda) 7, 467-479 (2017). 595 13. F. Rogalski et al., Epoxide hydrolases JHEH1 and JHEH2 deficiency impairs glucose 596 metabolism in Drosophila. Biochem Biophys Res Commun 748, 151313 (2025). 597 14. C. Steiner, F. Bozzolan, N. Montagne, M. Maibeche, T. Chertemps, Neofunctionalization 598 of "Juvenile Hormone Esterase Duplication" in Drosophila as an odorant-degrading 599 enzyme towards food odorants. Sci Rep 7, 12629 (2017). 600 15. C. K. Mirth et al., Juvenile hormone regulates body size and perturbs insulin signaling in 601 Drosophila. Proc Natl Acad Sci U S A 111, 7018-7023 (2014). 602 16. H. F. Nijhout, G. Davidowitz, D. A. Roff, A quantitative analysis of the mechanism that 603 controls body size in Manduca sexta. J Biol 5, 16 (2006). 604 17. D. E. Wheeler, H. F. Nijhout, Soldier determination in ants: new role for juvenile hormone. 605 Science 213, 361-363 (1981). 606 18. M. Magee, R. Spokony, Effect of D. melanogaster larval density on pupal size. MicroPubl 607 Biol 2023 (2023). 608 19. M. Santos, K. Fowler, L. Partridge, Gene-environment interaction for body size and larval 609 density in Drosophila melanogaster: an investigation of effects on development time, 610 thorax length and adult sex ratio. Heredity (Edinb) 72 ( Pt 5), 515-521 (1994). 611 20. M. Galloni, B. A. Edgar, Cell-autonomous and non-autonomous growth-defective mutants 612 of Drosophila melanogaster. Development 126, 2365-2375 (1999). 613 21. P. Klepsatel, E. Prochazka, M. Galikova, Crowding of Drosophila larvae affects lifespan 614 and other life-history traits via reduced availability of dietary yeast. Exp Gerontol 110, 615 298-308 (2018). 616 22. S. P. Bainbridge, M. Bownes, Staging the metamorphosis of Drosophila melanogaster. J 617 Embryol Exp Morphol 66, 57-80 (1981). 618 .CC-BY-NC 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted June 12, 2025. ; https://doi.org/10.1101/2025.06.09.657647doi: bioRxiv preprint 13 23. J. L. Wang, T. T. Saha, Y. Zhang, C. Zhang, A. S. Raikhel, Juvenile hormone and its 619 receptor methoprene-tolerant promote ribosomal biogenesis and vitellogenesis in the 620 Aedes aegypti mosquito. J Biol Chem 292, 10306-10315 (2017). 621 24. Q. He, Y. Zhang, Kr-h1, a Cornerstone Gene in Insect Life History. Front Physiol 13, 622 905441 (2022). 623 25. H. F. Nijhout, C. M. Williams, Control of moulting and metamorphosis in the tobacco 624 hornworm, Manduca sexta (L.): cessation of juvenile hormone secretion as a trigger for 625 pupation. J Exp Biol 61, 493-501 (1974). 626 26. J. W. Truman, L. M. Riddiford, Physiology of insect rhythms. 3. The temporal organization 627 of the endocrine events underlying pupation of the tobacco hornworm. J Exp Biol 60, 628 371-382 (1974). 629 27. L. M. Riddiford, How does juvenile hormone control insect metamorphosis and 630 reproduction? Gen Comp Endocrinol 179, 477-484 (2012). 631 28. R. C. King, S. K. Aggarwal, D. Bodenstein, The comparative submicroscopic morphology 632 of the ring gland of Drosophila melanogaster during the second and third larval instars. Z 633 Zellforsch Mikrosk Anat 73, 272-285 (1966). 634 29. Y. Ohhara, S. Kobayashi, N. Yamanaka, Nutrient-Dependent Endocycling in 635 Steroidogenic Tissue Dictates Timing of Metamorphosis in Drosophila melanogaster. 636 PLoS Genet 13, e1006583 (2017). 637 30. M. M. Rahman, X. Franch-Marro, J. L. Maestro, D. Martin, A. Casali, Local Juvenile 638 Hormone activity regulates gut homeostasis and tumor growth in adult Drosophila. Sci 639 Rep 7, 11677 (2017). 640 31. M. Kamimura, M. Takahashi, K. Kikuchi, A. M. Reza, M. Kiuchi, Tissue-specific regulation 641 of juvenile hormone esterase gene expression by 20-hydroxyecdysone and juvenile 642 hormone in Bombyx mori. Arch Insect Biochem Physiol 65, 143-151 (2007). 643 32. Z. Zhang et al., Depletion of juvenile hormone esterase extends larval growth in Bombyx 644 mori. Insect Biochem Mol Biol 81, 72-79 (2017). 645 33. F. G. Lu, K. Y. Fu, W. C. Guo, G. Q. Li, Characterization of two juvenile hormone epoxide 646 hydrolases by RNA interference in the Colorado potato beetle. Gene 570, 264-271 647 (2015). 648 34. J. Zhao, Y. Zhou, X. Li, W. Cai, H. Hua, Silencing of juvenile hormone epoxide hydrolase 649 gene (Nljheh) enhances short wing formation in a macropterous strain of the brown 650 planthopper, Nilaparvata lugens. J Insect Physiol 102, 18-26 (2017). 651 35. D. H. Hopkins et al., The molecular basis for the neofunctionalization of the juvenile 652 hormone esterase duplication in Drosophila. Insect Biochem Mol Biol 106, 10-18 (2019). 653 36. A. Mackert, K. Hartfelder, M. M. Bitondi, Z. L. Simoes, The juvenile hormone (JH) 654 epoxide hydrolase gene in the honey bee (Apis mellifera) genome encodes a protein 655 which has negligible participation in JH degradation. J Insect Physiol 56, 1139-1146 656 (2010). 657 37. S. M. Ismail, K. Satyanarayana, J. Y. Bradfield, K. H. Dahm, G. Bhaskaran, Juvenile 658 hormone acid: evidence for a hormonal function in induction of vitellogenin in larvae of 659 Manduca sexta. Arch Insect Biochem Physiol 37, 305-314 (1998). 660 38. L. J. Barton et al., Juvenile hormones direct primordial germ cell migration to the 661 embryonic gonad. Curr Biol 34, 505-518 e506 (2024). 662 39. J. Leyria et al., Crosstalk between Nutrition, Insulin, Juvenile Hormone, and Ecdysteroid 663 Signaling in the Classical Insect Model, Rhodnius prolixus. Int J Mol Sci 24 (2022). 664 40. J. Leyria, I. Orchard, A. B. Lange, Impact of JH Signaling on Reproductive Physiology of 665 the Classical Insect Model, Rhodnius prolixus. Int J Mol Sci 23 (2022). 666 41. Y. Y. Li et al., Enhanced Degradation of Juvenile Hormone Promotes Reproductive 667 Diapause in the Predatory Ladybeetle Coccinella Septempunctata. Front Physiol 13, 668 877153 (2022). 669 42. A. M. Box et al., Cell cycle variants during Drosophila male accessory gland 670 development. G3 (Bethesda) 14 (2024). 671 .CC-BY-NC 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted June 12, 2025. ; https://doi.org/10.1101/2025.06.09.657647doi: bioRxiv preprint 14 43. Y. Kurogi et al., The seminal vesicle is a juvenile hormone-responsive tissue in adult 672 male Drosophila melanogaster. Open Biol 14, 240315 (2024). 673 44. X. Y. Lyu, X. L. Wang, D. Q. Geng, H. Jiang, Z. Zou, Juvenile hormone acts on male 674 accessory gland function via regulating l-asparaginase expression and triacylglycerol 675 mobilization in Aedes aegypti. Insect Sci 30, 81-94 (2023). 676 45. N. A. Ramesh, A. M. Box, L. A. Buttitta, Post-eclosion growth in the Drosophila 677 ejaculatory duct is driven by Juvenile hormone signaling and is essential for male fertility. 678 Dev Biol 519, 122-141 (2025). 679 46. T. G. Wilson, S. DeMoor, J. Lei, Juvenile hormone involvement in Drosophila 680 melanogaster male reproduction as suggested by the Methoprene-tolerant(27) mutant 681 phenotype. Insect Biochem Mol Biol 33, 1167-1175 (2003). 682 47. E. T. Danielsen et al., A Drosophila Genome-Wide Screen Identifies Regulators of 683 Steroid Hormone Production and Developmental Timing. Dev Cell 37, 558-570 (2016). 684 48. Q. Gao et al., Key role of juvenile hormone in controlling reproductive diapause in 685 females of the Asian lady beetle Harmonia axyridis. Pest Manag Sci 78, 193-204 (2022). 686 49. S. Guo et al., The limited regulatory roles of juvenile hormone degradation pathways in 687 reproductive diapause preparation of the cabbage beetle, Colaphellus bowringi. J Insect 688 Physiol 119, 103967 (2019). 689 50. M. Werthebach et al., Control of Drosophila Growth and Survival by the Lipid Droplet-690 Associated Protein CG9186/Sturkopf. Cell Rep 26, 3726-3740 e3727 (2019). 691 51. K. Kongsuwan et al., A Drosophila Minute gene encodes a ribosomal protein. Nature 317, 692 555-558 (1985). 693 52. S. J. Marygold et al., The ribosomal protein genes and Minute loci of Drosophila 694 melanogaster. Genome Biol 8, R216 (2007). 695 53. J. P. David, H. M. Ismail, A. Chandor-Proust, M. J. Paine, Role of cytochrome P450s in 696 insecticide resistance: impact on the control of mosquito-borne diseases and use of 697 insecticides on Earth. Philos Trans R Soc Lond B Biol Sci 368, 20120429 (2013). 698 54. X. Zhou, C. Song, T. L. Grzymala, F. M. Oi, M. E. Scharf, Juvenile hormone and colony 699 conditions differentially influence cytochrome P450 gene expression in the termite 700 Reticulitermes flavipes. Insect Mol Biol 15, 749-761 (2006). 701 55. T. D. Sutherland et al., A cytochrome P450 terpenoid hydroxylase linked to the 702 suppression of insect juvenile hormone synthesis. Proc Natl Acad Sci U S A 95, 12884-703 12889 (1998). 704 56. S. Easwaran, M. Van Ligten, M. Kui, D. J. Montell, Enhanced germline stem cell longevity 705 in Drosophila diapause. Nat Commun 13, 711 (2022). 706 57. H. Gotoh et al., Juvenile hormone regulates extreme mandible growth in male stag 707 beetles. PLoS One 6, e21139 (2011). 708 58. C. G. Santos, F. C. Humann, K. Hartfelder, Juvenile hormone signaling in insect 709 oogenesis. Curr Opin Insect Sci 31, 43-48 (2019). 710 59. B. Wu et al., Sexual dimorphism of sleep regulated by juvenile hormone signaling in 711 Drosophila. PLoS Genet 14, e1007318 (2018). 712 60. A. P. Cabral et al., Pyriproxyfen, villain or good guy? A brief review. Arch Endocrinol 713 Metab 68, e240154 (2024). 714 61. J. L. Martin et al., Bio-efficacy of field aged novel class of long-lasting insecticidal nets, 715 against pyrethroid-resistant malaria vectors in Tanzania: A series of experimental hut 716 trials. PLOS Glob Public Health 4, e0002586 (2024). 717 62. N. A. Nur Aliah, S. Ab-Rahim, H. E. Moore, C. C. Heo, Juvenile hormone: Production, 718 regulation, current application in vector control and its future applications. Trop Biomed 719 38, 254-264 (2021). 720 63. D. R. Kethidi et al., Identification and characterization of a juvenile hormone (JH) 721 response region in the JH esterase gene from the spruce budworm, Choristoneura 722 fumiferana. J Biol Chem 279, 19634-19642 (2004). 723 64. D. R. Kethidi, Z. Xi, S. R. Palli, Developmental and hormonal regulation of juvenile 724 hormone esterase gene in Drosophila melanogaster. J Insect Physiol 51, 393-400 (2005). 725 .CC-BY-NC 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted June 12, 2025. ; https://doi.org/10.1101/2025.06.09.657647doi: bioRxiv preprint 15 65. V. Venkataraman, P. J. O'Mahony, M. Manzcak, G. Jones, Regulation of juvenile 726 hormone esterase gene transcription by juvenile hormone. Dev Genet 15, 391-400 727 (1994). 728 66. L. J. Barton et al., Unique and shared functions of nuclear lamina LEM domain proteins in 729 Drosophila. Genetics 197, 653-665 (2014). 730 67. D. Kim, J. M. Paggi, C. Park, C. Bennett, S. L. Salzberg, Graph-based genome alignment 731 and genotyping with HISAT2 and HISAT-genotype. Nat Biotechnol 37, 907-915 (2019). 732 68. P. Danecek et al., Twelve years of SAMtools and BCFtools. Gigascience 10 (2021). 733 69. F. Ramirez et al., deepTools2: a next generation web server for deep-sequencing data 734 analysis. Nucleic Acids Res 44, W160-165 (2016). 735 70. P. Ewels, M. Magnusson, S. Lundin, M. Kaller, MultiQC: summarize analysis results for 736 multiple tools and samples in a single report. Bioinformatics 32, 3047-3048 (2016). 737 71. G. H. Putri, S. Anders, P. T. Pyl, J. E. Pimanda, F. Zanini, Analysing high-throughput 738 sequencing data in Python with HTSeq 2.0. Bioinformatics 38, 2943-2945 (2022). 739 72. M. I. Love, W. Huber, S. Anders, Moderated estimation of fold change and dispersion for 740 RNA-seq data with DESeq2. Genome Biol 15, 550 (2014). 741 73. L. Kolberg et al., g:Profiler-interoperable web service for functional enrichment analysis 742 and gene identifier mapping (2023 update). Nucleic Acids Res 51, W207-W212 (2023). 743 74. E. Imura, Y. Yoshinari, Y. Shimada-Niwa, R. Niwa, Protocols for Visualizing 744 Steroidogenic Organs and Their Interactive Organs with Immunostaining in the Fruit Fly 745 Drosophila melanogaster. J Vis Exp 10.3791/55519 (2017). 746 75. J. B. Brown et al., Diversity and dynamics of the Drosophila transcriptome. Nature 512, 747 393-399 (2014). 748 76. V. R. Chintapalli, J. Wang, J. A. Dow, Using FlyAtlas to identify better Drosophila 749 melanogaster models of human disease. Nat Genet 39, 715-720 (2007). 750 77. D. P. Leader, S. A. Krause, A. Pandit, S. A. Davies, J. A. T. Dow, FlyAtlas 2: a new 751 version of the Drosophila melanogaster expression atlas with RNA-Seq, miRNA-Seq and 752 sex-specific data. Nucleic Acids Res 46, D809-D815 (2018). 753 754 755 .CC-BY-NC 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted June 12, 2025. ; https://doi.org/10.1101/2025.06.09.657647doi: bioRxiv preprint 16 Figures 756 757 758 759 Figure 1: Two classes of Juvenile hormone (JH) degradation enzymes in Drosophila. A 760 Schematic diagram of JH synthesis and degradation pathways. Left: JH synthesis from Acetyl-761 CoA precursors through methylation by Jhamt. Middle, JH signaling response in which JHs bind 762 transcription factors, Methoprene tolerant (Met) or Germ cell expressed (Gce), which translocate 763 and bind to response elements together with co-factor, Taiman (Tai). Right: JH degradation by 764 extracellular JH esterases (JHEs) to produce JH acids, or intracellular JH epoxide hydrolases 765 (JHEHs) to produce JH diols. B Line schematics of JHE and JHEH protein features and 766 sequence similarity. Jhe and JheDup share conserved esterase catalytic domains (pink) and 767 Jheh1,2, and 3 share alpha/beta hydrolase catalytic domains (purple). Catalytic triad residues are 768 highlighted in red and substrate specificity residues are in black. C Predicted catalytic pocket for 769 JH degradation enzymes with catalytic side chains shown. D Expression of JH degradation 770 .CC-BY-NC 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted June 12, 2025. ; https://doi.org/10.1101/2025.06.09.657647doi: bioRxiv preprint 17 enzymes across Drosophila development and tissues. w3rd – wandering third instar larvae, wpp 771 – white prepupal stage. Data retrieved from (75-77). 772 773 774 .CC-BY-NC 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted June 12, 2025. ; https://doi.org/10.1101/2025.06.09.657647doi: bioRxiv preprint 18 775 .CC-BY-NC 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted June 12, 2025. ; https://doi.org/10.1101/2025.06.09.657647doi: bioRxiv preprint 19 776 Figure 2: JHEs and JHEHs have opposing functions in developmental growth. A Schematic 777 of genes encoding JH degradation enzyme and knockout strains generated in this study. Exons 778 are shown as taller grey rectangles. guide RNAs used to delete both Jhe and JheDup and all 779 three copies of Jheh in respective mutants are shown underneath each locus as red lines. B 780 Representative images of wild type, JHE DKO, and JHEH TKO pupae, scale bars represent 1 781 mm. C Graph of pupal length in females (orange, top) and males (blue, bottom). Genotypes are 782 noted below. Each dot represents one pupa, with total sample size noted below each box plot. 783 Box plots represent median, 25th through 75th percentile with 1.5 IQR whiskers. D. Graph of pupal 784 length in females (orange, top) and males (blue, bottom) in each. Genotype is noted under the 785 male graph. Each dot represents one pupa, with total sample size noted below each box plot. Box 786 plots represent median, 25th through 75th percentile with 1.5 IQR whiskers. For B and C, a Mann-787 Whitney U test was used to test significance (* = p<0.05, ** = p<0.01, *** = p<0.001, n.s. = not 788 significant). 789 790 791 .CC-BY-NC 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted June 12, 2025. ; https://doi.org/10.1101/2025.06.09.657647doi: bioRxiv preprint 20 792 793 Figure 3: JHEHs, but not JHEs, are required for developmental timing. A Graph of pupal 794 timing in which the Y axis is the percent of animals that pupariated relative to days post egg lay 795 (X axis) in control, JHE DKO, and JHE DKO carrying a BAC that contains both Jhe and JheDup 796 loci. B Graph of the percent pupariation relative to days post egg lay in control animals, JHEH 797 .CC-BY-NC 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted June 12, 2025. ; https://doi.org/10.1101/2025.06.09.657647doi: bioRxiv preprint 21 TKO animals, or JHEH TKO animals carrying a BAC that contains Jheh1, Jheh2, and Jheh3 loci. 798 C Graph of percent pupariation relative to relative to days post egg lay in control animals, JHEH 799 TKO animals, or JHEH TKO; jhamt animals. All experiments were performed at 25°C. Each dot 800 indicates mean percent pupariated and whiskers represent standard deviation. The number of 801 biological replicates (rep) and total number of animals (n) are shown to the right of the genotype. 802 All 50% pupariation data was normally distributed and used for a two-tailed Student’s t-test was 803 used to test significance (* = p<0.05, ** = p<0.01, *** = p<0.001, n.s. = not significant). 804 805 806 .CC-BY-NC 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted June 12, 2025. ; https://doi.org/10.1101/2025.06.09.657647doi: bioRxiv preprint 22 807 808 Figure 4: JHE and JHEH mutants have shared and unique transcriptional signatures. A 809 Graph of the total number of Differentially Expressed Genes (DEGs) from bulk RNA sequencing 810 of white prepupae. Downregulated genes are in blue and upregulated genes are in orange. The 811 total number of DEGs is shown at the top of each bar B UpSet plot showing either DEGs that are 812 uniquely dysregulated in either JHE DKO or JHEH TKO animals (Left), or DEGs shared by JHE 813 DKO and JHEH TKO animals (Right). The genotypes are noted below, and which samples 814 display the shared DEGs is noted by solid circles with a connecting line. The number of unique or 815 shared DEGs is shown at the top of each bar. C Heatmap of differentially expressed Cytochrome 816 P450 genes (top) and ecdysone synthesis genes (bottom). Genes are noted to the left, heat map 817 scale is to the right, and genotype is noted below each column. D Raw tracks showing reads for 818 ecdysone synthesis genes, spok, shd, and phm, as well as Jheh1, Jheh2, and Jheh3 loci. 819 Genotype is noted to the left, the gene locus is shown below the tracks. M=male, F=female 820 821 822 .CC-BY-NC 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted June 12, 2025. ; https://doi.org/10.1101/2025.06.09.657647doi: bioRxiv preprint 23 823 .CC-BY-NC 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted June 12, 2025. ; https://doi.org/10.1101/2025.06.09.657647doi: bioRxiv preprint 24 824 Figure 5: Developmental requirements of JHEHs are linked to ecdysone signaling. A Graph 825 of the percent pupariation relative to days post egg lay in control and JHEH TKO fed either the 826 active form of Ecdysone (20E) dissolved in ethanol (EtOH) or fed an equivalent amount of vehicle 827 EtOH. Each dot indicates mean percent pupariated and whiskers represent standard deviation. 828 The number of biological replicates (rep) and total sample size (n) are shown to the right of each 829 genotype. All 50% pupariation data was normally distributed and used for a two-tailed Student’s t-830 test was used to test significance (* = p<0.05, ** = p<0.01, *** = p<0.001, n.s. = not significant). B 831 Representative images of ring glands from wandering third instar larval females (left) and males 832 (right) stained with DAPI (gray). Ring glands are outlined. Scale bars represent 50mm. C 833 Quantification of ring gland volume in wandering third instar females (orange, left) and males 834 (blue, right). D Quantification of the total volume of prothoracic gland nuclei in each ring gland in 835 wandering third instar females (orange, left) and males (blue, right). For C and D, dots indicate 836 the measurement of an individual ring gland and sample size for each group is shown under the 837 respective box plot. Box plots represent median, 25th through 75th percentile with 1.5 IQR 838 whiskers. A Mann–Whitney U test was used to test significance (* = p<0.05, ** = p<0.01, *** = 839 p<0.001, n.s. = not significant). E Schematic summarizing the working model based on data from 840 this study, in which JHEs restrain growth during development, while JHEHs restrict the time it 841 takes to undergo developmental transitions from larvae (light beige) to pupae (brown). Key 842 transcriptomic signatures shared and unique for each class of JH degradation enzyme are 843 summarized as Venn diagram. 844 845 846 .CC-BY-NC 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted June 12, 2025. ; https://doi.org/10.1101/2025.06.09.657647doi: bioRxiv preprint