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20-Hydroxyecdysone (20E) signaling as a promising target for the chemical control of malaria vectors


With the rapid development and spread of resistance to insecticides among anopheline malaria vectors, the efficacy of current World Health Organization (WHO)-approved insecticides targeting these vectors is under threat. This has led to the development of novel interventions, including improved and enhanced insecticide formulations with new targets or synergists or with added sterilants and/or antimalarials, among others. To date, several studies in mosquitoes have revealed that the 20-hydroxyecdysone (20E) signaling pathway regulates both vector abundance and competence, two parameters that influence malaria transmission. Therefore, insecticides which target 20E signaling (e.g. methoxyfenozide and halofenozide) may be an asset for malaria vector control. While such insecticides are already commercially available for lepidopteran and coleopteran pests, they still need to be approved by the WHO for malaria vector control programs. Until recently, chemicals targeting 20E signaling were considered to be insect growth regulators, and their effect was mostly studied against immature mosquito stages. However, in the last few years, promising results have been obtained by applying methoxyfenozide or halofenozide (two compounds that boost 20E signaling) to Anopheles populations at different phases of their life-cycle. In addition, preliminary studies suggest that methoxyfenozide resistance is unstable, causing the insects substantial fitness costs, thereby potentially circumventing one of the biggest challenges faced by current vector control efforts. In this review, we first describe the 20E signaling pathway in mosquitoes and then summarize the mechanisms whereby 20E signaling regulates the physiological processes associated with vector competence and vector abundance. Finally, we discuss the potential of using chemicals targeting 20E signaling to control malaria vectors.

Graphical abstract


Malaria is spread when Plasmodium parasites are transmitted between humans via hematophagous female anopheline mosquitoes. While the 2019 statistics (409,000 deaths globally and ± 3.0 billion USD invested in malaria control and elimination programs) already reveal the high socio-economic impact of this disease [1], recent models predict that with the unprecedented coronavirus disease 2019 (COVID-19) pandemic, deaths due to malaria in low- and middle-income countries could increase by 36% over the next 5 years [2]. Therefore, African countries, where currently more than 90% of all malaria deaths worldwide occur [1], are the most at risk.

The burden of malaria is managed by a multi-disciplinary approach which combines targeting the parasite (artemisinin combination therapy) [3, 4], the vector (World Health Organization [WHO]-approved insecticides and long-lasting insecticide treated bednets) [5, 6] and, to some extent, the environment (habitat modification or larval source management) [7,8,9]. Additionally, two vaccines are currently under trial: RTS,S and AGS-v [10,11,12,13]. The pre-erythrocytic stage antimalarial vaccine RTS,S targets the circumsporozoite surface protein of Plasmodium falciparum and is currently in phase IV clinical trials in Ghana, Kenya, and Malawi [1, 11, 12]. In contrast, the AGS-v vaccine targets four conserved saliva peptides in Anopheles spp., Aedes spp. and Culex spp., and has shown promising results in terms of safety and immunogenicity during its phase I clinical trial in humans [10]. Of all these interventions, vector control plays a central role. In fact, the WHO has stated that “vector control is a vital component of malaria prevention, control, and elimination strategies because it can be highly effective in providing personal protection and/or reducing disease transmission” [6].

The WHO-approved vector control strategies can be divided into two categories, referred to as core interventions and supplementary interventions [6]. Core interventions comprise incorporating insecticides into bednets (long-lasting insecticide-treated nets [LLINs]) or spraying insecticides onto the walls of houses (indoor residual spraying [IRS]). At present, four WHO-approved classes of insecticides are used in IRS interventions, namely pyrethroids, carbamates, organophosphates and organochlorines, as opposed to LLINs for which only pyrethroids have been approved owing to their relative safety [6]. Collectively, LLINs and IRS have led to an 18% global reduction in malaria cases over the past 8 years [14]. Supplementary interventions, on the other hand, include larval source management (LSM) via biological or chemical larvicides, as well as the disruption of breeding sites [6]. Although there are reports of highly successful LSM programs [15], in reality most malaria-endemic countries have so many breeding sites that LSM becomes both expensive and impractical.

Unfortunately, field resistance to insecticides is common and widespread, with the result that the efficacy of the core interventions has been drastically impaired. In particular, pyrethroid resistance has been detected in all the major African malaria vectors, including Anopheles gambiae, An. coluzzii, An. arabiensis and An. funestus [16, 17]. This has created an urgent need for enhanced insecticides, such as those carrying synergists (e.g. piperonyl butoxide) [18,19,20], antimalarials (e.g. atovaquone) [21] or chemicals with novel targets. The latter would ideally target a mosquito pathway that is essential for vector competence and vector abundance, have minimal effect on non-target species and be effective against mosquitoes that are resistant to the classes of insecticides currently under use. One such pathway of interest is that of the steroid hormone, 20-hydroxyecdysone (20E) [22]. Indeed, studies in An. gambiae suggest that chemicals which target the 20E signaling pathway have the potential to control malaria vectors, both at the adult [23,24,25] and immature stages [26, 27]. This is because the 20E pathway regulates several key physiological processes in mosquitoes, such as blood-feeding, insecticide resistance, pathogen development, molting, mating, fecundity and fertility (Table 1). In this review, we first describe the 20E signaling pathway in mosquitoes, then discuss the mechanisms whereby 20E signaling regulates the physiological processes associated with vectorial capacity, such as susceptibility to Plasmodium infection, egg production and development. Finally, we discuss the potential of chemical control interventions targeting 20E signaling to reduce the burden of malaria.

Table 1 Phenotypes associated with manipulating 20-hydroxyecdysone titers, activity or signaling in mosquitoes

An overview of the 20E signaling pathway in mosquitoes

20E biosynthesis is a multi-enzyme process

From the food they ingest, mosquitoes obtain cholesterol, the precursor molecule for 20E biosynthesis [28, 29]. In larvae and pupae, the conversion of cholesterol to 20E takes place in the prothoracic glands [30, 31], while in adults 20E biosynthesis occurs in the ovaries and fat body (females) and in the the accessory glands (males) [32,33,34,35]. Although most knowledge related to 20E biosynthesis comes from studies in Drosophila, orthologues of the enzymes involved in this process have been characterized in mosquitoes. The first enzyme in this process, neverland, catalyzes the conversion of dietary cholesterol to 7-dehydrocholesterol [36,37,38], which is in turn metabolized to 5ß-ketodiol via Δ4-diketol, 5β-diketol and a few uncharacterized intermediate metabolites (Fig. 1a). Hence, the term “black box” has been used to describe this part of the 20E biosynthesis pathway [37]. Nonetheless, research has shed some light on the intermediate steps and characterized the enzymes, spook, shroud, spookier and spookiest in the Drosophila melanogaster model [39,40,41,42]. Of these, spook and shroud orthologues have been identified in Ae. aegypti and/or An. gambiae [38, 43]. In particular, spook knockdown by RNA interference in An. gambiae decreased the production of 20E in the ovaries, confirming that spook has the same function in both Drosophila and An. gambiae [44]. The metabolite 5β-ketodiol is further converted to 5β-ketotriol and then transformed to 2-deoxyecdysone before it is finally changed to ecdysone (Fig. 1a); these three steps are catalyzed by cytochrome P450 (CYP) enzymes CYP306a1 (phantom), CYP302a1 (disembodied) and CYP315A1 (shadow), respectively [45,46,47,48]. Finally, ecdysone is secreted from the prothoracic glands or ovaries into the hemolymph. It then enters the fat body where it is oxidized to the active form 20E, by another cytochrome p450 enzyme, namely CYP314a1 (shade) [49]. 20E is then release from the fat body and transported to different cells and tissues, as needed. Orthologues of these four enzymes have been identified and functionally characterized in An. gambiae, confirming their roles in 20E biosynthesis in the mosquito [32].

Fig. 1
figure 1

20-Hydroxyecdysone (20E) signaling in insects. a 20E biosynthesis from dietary cholesterol, based on studies in Drosophila melanogaster. Metabolites and enzymes are indicated in black and pink, respectively. The “black box” (where the exact metabolites/enzymes are unknown) is indicated by the grey area. Orthologues of these enzymes have been characterized in mosquitoes. b Once 20E binds to its heterodimer EcR/USP receptor, the latter is activated and acts as a transcription factor, binding to an enhancer region known as the ecdysone response elements (EcRE). Binding of EcR/USP to EcRE activates the transcription of early genes (E75, E74, HR3 and Broad-Complex). These four early genes in turn act as transcription factors, inducing or repressing the expression of downstream genes involved in vector competence and vector abundance. Br-C Broad complex, EcR ecdysone receptor, USP ultraspiracle

Activation and regulation of the 20E signaling cascade

In mosquitoes, the 20E signaling pathway is activated when 20E binds to its nuclear receptor, the ecdysone receptor complex (Fig. 1b). The ecdysone receptor complex is a heterodimer consisting of the ultraspiracle protein (USP) and the ecdysone receptor protein (EcR) (Fig. 1b). USP and EcR are orthologues of the mammalian retinoid-X receptor (RXR) and farnesoid X receptor (FXR), respectively [50, 51]. Both EcR and USP are members of the steroid receptor superfamily, which is characterized by five domains: A/B (transactivation), C (DNA-binding), D (hinge), E (ligand-binding) and F (transactivation) [52, 53]. Recently, the F-domain of Ae. aegypti EcR was shown to bind to the metal ions Cu2+ and Zn2+, thereby inducing a helical structure in the protein and promoting ligand binding specificity [54, 55]. Only EcR binds to 20E, but a study in Drosophila revealed that EcR requires heterodimerization with USP to successfully bind the hormone [56]. It has also been reported that USP might be involved in the allosteric regulation of EcR, altering its conformation to favor the hormone- and DNA-binding properties of EcR [57, 58]. The EcR–USP complex acts as a transcription factor, binding with high affinity to the ecdysone response elements (EcRE) [59], an enhancer located in the upstream regulatory regions of ecdysone-responsive genes. In Ae. aegypti, EcRE are composed of DNA motifs that are either inverted or direct repeats [60]. Binding of EcR–USP to EcRE activates the transcription of “early genes” such as E75, E74, HR3 and Broad-Complex [61,62,63,64,65,66,67]. These early genes in turn also act as transcription factors, inducing or repressing the expression of several downstream genes which control reproduction, immunity and development (Fig. 1b) [68]. However, there are some cases where the EcR–USP transcription factor directly binds to the EcRE regions of downstream genes, such as the vitellogenin gene [69].

Two EcR isoforms (A and B) have been identified in Ae. aegypti, and they appear to vary in biological function as well as spatial and/or temporal expression [63, 70,71,72]. For example, in the fat body, EcR-A expression increases during the vitellogenic period from 12 h post-blood meal (hPBM) to 24 hPBM, and then decreases by 36 hPBM; while EcR-B is most abundant in the pre-vitellogenic and post-vitellogenic period [72]. Similarly, two USP isoforms have been described in Ae. aegypti [53, 71, 73]. The abundance of USP-A in the fat body is highest in the pre-vitellogenic and late vitellogenic period, while USP-B is highly expressed during vitellogenesis [73]. In the midgut of Ae. aegypti larvae, the EcR-B and USP-A isoforms are more abundant than EcR-A and USP-B [74]. In addition to the mosquito fat body and midgut, isoforms of the ecdysone receptor subunits have also been detected in the ovaries and male accessory glands [32, 70]. Also, while there is currently no experimental evidence in mosquitoes, to our knowledge, EcR isoforms have also been detected in the central nervous system of Agrotis ipsilon [75], Apis mellifera [76] and Bombyx mori [77].

Several regulators coordinate the spatio-temporal expression and activation of EcR and USP. For example, before a blood meal, USP is bound to the nuclear factor HR38 in the fat body of Ae. aegypti, but HR38 is later displaced by EcR during the vitellogenesis period (12–24 hPBM) [78]. Another important regulator is the “early gene” E75. Three isoforms of E75 (E75A, E75B, and E75C) have been detected in the fat body of Ae. aegypti post-blood meal, and functional studies have revealed that silencing either E75A or E75C shifts the peak expression of EcR-A (which normally occurs 12–24 hPBM) to 24–30 hPBM [79]. Two additional proteins, FISC and βFTZ-F1, act as co-activators of EcR/USP in the fat body of blood-fed Ae. aegypti females, by recruiting and binding to the EcR/USP complex at the EcRE region of the vitellogenin promoter. This association was absent in the non-bloodfed cohorts [80]. Besides vitellogenesis, the timely regulation of metamorphosis also requires the presence cofactors to regulate EcR/USP. For example, in Ae. aegypti fourth instar larvae, the CREB-binding protein (CBP)—whose primary function is to loosen the chromatin structure to render the DNA regulatory regions accessible to transcription factors—suppresses the expression of EcR-A, to prevent premature molting. When CBP is silenced, EcR-A expression is elevated, and the larvae prematurely metamorphosed into pupae [63, 81].

20E signaling regulates multiple physiological processes at different stages of the mosquito life-cycle

Depending on the environmental conditions, mosquito eggs hatch into larvae within 2–3 days (reviewed in [82, 83]). The newly emerged larvae then undergo four successive molts from first to fourth instar larvae, lasting in total approximately 5–10 days, prior to becoming pupae. About 1–3 days later, adult mosquitoes emerge from their pupal cuticle. The 20E signaling pathway is an integral part of mosquitoes’ life-cycle (Fig. 2) as it regulates several physiological processes associated with development, reproduction or susceptibility to pathogen infection, as discussed below.

Fig. 2
figure 2

Manipulating 20E titers, activity or signaling affects several physiological processes at each stage of a mosquito life-cycle. Only processes that have been experimentally confirmed in mosquitoes are represented. The asterisk (*) indicates that this is not the role of 20E in males, but rather the role that the male-secreted 20E plays in females, once it is transferred to their atrium during mating

Egg development and oviposition

Egg development requires nutrients from blood

In anautogenous female mosquitoes, egg development requires nutrients obtained from a blood meal. After the ingested blood is digested to release cholesterol and proteins (Fig. 3), the cholesterol is used for 20E production, while the midgut proteases hydrolyze the proteins into amino acids. These amino acids are incorporated into various metabolic pathways in the fat body such as lipid and carbohydrate metabolic pathways, resulting in the production of lipids, yolk proteins and energy (Fig. 3) [84]. The pathways for carbohydrate metabolism, including glycogen metabolism, gluconeogenesis, the citric acid cycle and glycolysis, have previously been found to be upregulated at 18–24 hPBM, which is also the peak of 20E synthesis in females [85]. Further analysis revealed that silencing EcR downregulated the expression of several genes involved in glycolysis and glycogen metabolism, resulting in an increase in fat body glycogen, decreased ATP levels, and the accumulation of sugars (glucose and fructose) [85]. Dong et al. [86] later reported that 20E regulates carbohydrate metabolism via the HR38 nuclear transcription factor. Similarly, 20E signaling was also shown to regulate lipid metabolism in the fat body (Fig. 3), as silencing of EcR resulted in increased levels of triacylglycerols and decreased β-oxidation [87]. This allows the insect to store lipids as either an energy source for egg maturation or to incorporate these lipids in the developing oocytes [88].

Fig. 3
figure 3

Anautogenous mosquitoes use the blood nutrients to produce egg components in the fat body. In Aedes aegypti, digestion of the blood meal involves several metabolic processes (indicated in boxes), many of which are regulated by 20E signaling, as indicated by the “20E” label. The carbohydrate-related metabolic pathways are indicated in green boxes, while the lipid-related metabolic pathways are indicated in red boxes. CoA Coenzyme A, TAGs triacylglycerols, TOR target of rapamycin

20E-mediated oviposition requires brain-secreted hormones

In Ae. aegypti, the ingestion of a blood meal triggers the brain to release two neurohormones—ovary ecdysteroidogenic hormone (OEH) and insulin-like peptide 3 (ILP3) — into the hemolymph [89, 90] (Fig. 4). ILP3 and OEH bind to their receptors (the insulin and OEH receptors, respectively), located on the follicle cells of the ovarioles [91, 92]. This binding triggers a phosphorylation cascade, which in turn activates the target of rapamycin (TOR) and insulin pathways, and ultimately blocks the activity of the glycogen synthase kinase 3 (GSK3) protein [93]. Blocking of GSK3 results in the proliferation of follicle cells, an indication that the ovarioles are ready to produce ecdysone [93]. Hence, the blood-derived cholesterol (transported by lipophorin, a carrier protein synthesized in the fat body) and amino acids (via amino acid transporters) enter the follicle cells where they serve as building blocks for ecdysone synthesis [94,95,96,97,98]. Ecdysone is then released from the ovaries and enters the fat body where it is converted into 20E (Fig. 4). In the fat body, 20E triggers the synthesis of yolk protein precursors (YPPs) such as vitellogenin, vitellogenin carboxypeptidase or cathepsin b-like protease [35]. These YPPs are released into the hemolymph and transported to the growing oocytes in the ovaries where they are taken up by receptor-mediated endocytosis (Fig. 4). Although the regulation of YPP transport has not been investigated in mosquitoes, Carney et al. [99] reported that Drosophila females with EcR mutations displayed decreased transport of YPPs to the ovaries compared to untreated controls. The oocytes, now fully developed into eggs, are laid by mosquitoes in aquatic environments.

Fig. 4
figure 4

20E signaling regulates oogenesis (for details see text). The steps regulated by 20E signaling in mosquitoes are indicated by orange asterisks (*) OEH Ovary ecdysteroidogenic hormone, ILP insulin-like peptide, YPPs yolk protein precursors


Mosquito larvae undergo four developmental stages, from the first instar to the fourth instar, all of which take place in aquatic environments. 20E signaling is essential to molting from one larval stage to the next, as indicated by the high levels of 20E and its receptor in Ae. aegypti during larval ecdysis [63, 100, 101]. In addition, it has been reported that manipulating 20E titers and signaling in Culex quinquefasciatus, Cx. pipiens, Ae. aegypti, and Anopheles gambiae impairs larval fitness, development, survival, cuticulogenesis and molting (Table 1).


Key events in mosquito pupal development include sexual dimorphism [102], programmed cell death and cell differentiation [71], ecdysis [63] and the formation of adult structures such as wings [103]. In Ae aegypti, both male and female pupae display an increase in titers of 20E, ecdysone, 2-deoxyecdysone and other steroid molting hormones. The ecdysteroid titers reach a much higher level in males than in females, and the peak appears earlier in male pupae than in female ones [102]. This difference may explain why males eclose into adults sooner than females, an important feature for mating success in adult mosquitoes [102]. In terms of programmed cell death and cell differentiation, Parthasarathy and Palli [71] observed that during the initial pupal stages in Ae. aegypti, both EcR isoforms (EcR-A and EcR-B) are present in larval cells undergoing apoptosis, while EcR-B is present in the imaginal diploid cells of pupae, indicating that both isoforms facilitate the turnover of larval cells while EcR-B plays an additional role in the development of imaginal cells. The role of 20E in the formation of wing structure has not yet been investigated in mosquitoes; however, in the domesticated silkworm Bombyx mori, manipulation of 20E titers suggests a pivotal role in healthy wing development [103]. Collectively, the roles of 20E signaling in different aspects of pupal development are in agreement with the observation that manipulating 20E signaling in the tsetse fly Glossina morsitans results in incomplete pupation [104].


Female adults

In female adults, several physiological parameters are affected by manipulating 20E signaling, including susceptibility to Plasmodium infection [105, 106], longevity [23], insecticide resistance [107, 108], blood-feeding and vitellogenesis [34, 52, 64, 109] (Fig. 4). As vitellogenesis has been discussed in above (section Egg development and oviposition), in this section we focus on the other phenotypes.

The sporogonic cycle of malaria parasites begins when Anopheles mosquitoes ingest Plasmodium gametocytes while feeding on infected hosts [110]. In the insect midgut, these gametocytes rapidly differentiate into male and female gametes. The zygotes that are formed from fertilization develop into motile ookinetes which, upon crossing the midgut epithelium and its basal membrane, transform into oocysts that remain fixed at the interface of the midgut and hemolymph [111]. Once fully matured (~ 14 hPBM), the oocyst “bursts” and releases sporozoites in the hemolymph [112]. These navigate to the salivary glands where they are ready to be injected into the next host during a following blood meal [111]. For the development of transmission-blocking interventions, three parameters related to the sporogonic cycle are relevant: (i) oocyst prevalence, which refers to the percentage of mosquitoes with contain oocysts after taking an infectious blood meal [113]; (ii) oocyst intensity, which is the average number of oocysts per mosquito; these are measured as functions of transmission-reducing activity and transmission blocking activity respectively, with TBA considered to be a more useful assessment of efficacy given that even just one oocyst can produce many infectious sporozoites [114]; (iii) duration of the sporogonic cycle, known as the extrinsic incubation period (EIP), which is a measure of the time needed for a mosquito to become infectious [115]. The EIP can be affected by factors such as environmental temperature and by underlying genetic features of both the vector and parasite [115].

Manipulating the titers, activity or signaling of 20E affects the parasite’s oocyst prevalence (Plasmodium falciparum and P. berghei), oocyst intensity (P. falciparum and P. berghei), and EIP (P. falciparum) [105, 106]. However, the molecular mechanisms by which these parasite parameters are regulated are poorly understood. From an immune response perspective, it is possible that 20E signaling regulates P. berghei development via several immune effectors, including antimicrobial peptides, prophenoloxidases, CLIP serine proteases or lysozymes [106]. In addition, given the increase in the number of phagocytic cells and activity after 20E injection [106], it is possible that the 20E pathway reduces susceptibility to P. berghei infection by increasing the phagocytic defense mechanism.

In terms of longevity, boosting 20E signaling by applying methoxyfenozide (see section 20E agonists) onto the thorax of Anopheles mosquitoes resulted in a reduced lifespan with increasing concentrations of methoxyfenozide [23]. This is important because if the mosquito lifespan becomes shorter than the parasite EIP, malaria transmission is effectively reduced (reviewed in [116]). In the context of insecticide resistance, previous reports have shown that silencing spookiest or shade in deltamethrin-resistant Cx. pipiens mosquitoes rendered these mosquitoes more susceptible to the pyrethroid [107, 108]. To date, there is no report, to our knowledge, directly linking 20E biosynthesis enzymes to Anopheles resistance to insecticides. However, previous studies have shown that shade is overexpressed in dichlorodiphenyltrichloroethane (DDT)-resistant An. gambiae [117], while shade and phantom are overexpressed in DDT- and pyrethroid-resistant An. funestus [118], suggesting that these may also be implicated in insecticide resistance in Anopheles spp. Nonetheless, functional studies will be required to directly assess the role of shade and phantom in insecticide resistance in An. gambiae and/or An. funestus. Moreover, it may be relevant to determine whether the overexpression of 20E-related genes in insecticide-resistant An. gambiae affects their longevity, as it could have implications for the parasite EIP, and thus malaria transmission. Finally, it is interesting to note that 20E also plays a role in the extent of nutrient uptake, a feature that has been observed in different insects (e.g. An. freeborni, Helicoverpa armigera, and Bombyx mori) injected with 20E [109, 119, 120]. While it appears that 20E only plays an indirect role in nutrient seeking by blocking dopamine signaling which normally promotes food-seeking behavior [119, 121], this observation is yet to be investigated in mosquitoes.


In An. gambiae males, 20E is exclusively synthesized in the male accessory glands (MAGs), and its production increases from the day of adult emergence until the male become sexually mature and active (i.e. 3–6 days post-emergence) [32]. During copulation, some of the male-synthesized 20E is transferred to the female atrium, as part of a mating plug secreted by MAGs, and it is replenished in the male after copulation [32]. This male-to-female transfer of 20E has been observed in at least four anopheline species, including An. gambiae, An. arabiensis, An. stephensi and An. dirus [122]. Once in the female atrium, the male-derived 20E regulates several processes, such as oviposition, egg fertility and refractoriness to further copulation, and it helps maintain the integrity of the sperm in the spermatheca [25, 123,124,125]. Although little is known about the mechanisms whereby the male-derived 20E regulates these processes, it has been reported that the female atrium-specific MISO protein interacts with the male-derived 20E to regulate egg production [123]. Overall, both male- and female-synthesized 20E contributes towards the reproductive behavior and success of anopheline female mosquitoes (Fig. 2).

The potential of chemical control interventions targeting 20E signaling

20E agonists

20-hydroxyecdysone agonists are insect growth regulators (IGRs) that compete with 20E to bind to its EcR receptor complex, thereby overactivating the 20E signaling pathway. Interestingly, both EcR and USP subunits of the receptor complex are needed for successful 20E agonist activity [24, 126]. The most studied class of 20E agonists are the dibenzoylhydrazine (DBH) compounds [126, 127]. Currently, many IGRs based on DBH compounds are commercially available, such as tebufenozide (RH-5992), methoxyfenozide (RH-2485), halofenozide (RH-0345), fufenozide, chromafenozide (ANS-118) or RH-5849 [128,129,130]. These compounds were initially formulated against lepidopteran and coleopteran crop pests; however there is increasing evidence that they could also be used to control mosquito populations at different developmental stages, including eggs, larvae and adults [23, 26, 27, 131].

Water treatment with methoxyfenozide has been shown to reduce egg hatch rate in Cx. pipiens [131], as well as larval mortality in An. gambiae, Ae. aegypti, and Cx. quinquefasciatus [26, 27]. The effect of 20E agonists on pupae is yet to be tested in mosquitoes, but treating Spodoptera litura pupae with RH-5849 resulted in pupal development abnormalities and a subsequent decrease in adult emergence [132]. In An. gambiae adults, it has been demonstrated that methoxyfenozide and halofenozide reduced P. falciparum and P. berghei transmission, respectively [23, 24]. In addition, fecundity, fertility, mating success and adult longevity were all significantly decreased after DBH exposure [23]. As such, DBH compounds affect both vector abundance and vector competence and have the additional benefit of showing minimal effect on non-target species, as opposed to conventional insecticides which may be toxic to humans and other arthropods (Table 2).

Table 2 Insecticidal properties of the dibenzoylhydrazine compounds which have shown promising results against mosquitoes

Resistance to DBH compounds has been studied in the lepidopterans Plutella xylostella, Cydia pomonella, and Spodoptera exigua [133,134,135,136,137,138], and two mechanisms have been identified. As with classic insecticides, the first mechanism involves an increase in the activity of detoxification enzymes such as carboxylesterase, aryl-acylamidase, cytochrome P450s or glutathione-S-transferases [138,139,140,141]. While an increased expression of cytochrome P450s also constitutes the resistance mechanism of some carbamates, pyrethroids and organochlorines (reviewed in [142]), it is interesting to note that cross-resistance between DBH compounds and these classic insecticides is not always guaranteed (see following paragraph). The second resistance mechanism, identified in P. xylostella, involves the microRNA miR-189942, which decreases the expression of the EcR-B isoform, thereby reducing the susceptibility to fufenozide (because fewer binding sites are available for the 20E agonist) [143]. However, resistance to DBH compounds is unstable due to fitness costs such as higher mortality rate and decreased reproductive capacity that are associated with the DBH-resistant phenotype [137, 138, 144, 145]. As such, most insects revert back to the susceptible phenotype when the 20E agonist is removed [137, 138]. This is a significant advantage over classical insecticides where long-term use has resulted in fixed population-wide genetic changes that confer resistance. Alternatively, the emergence and spread of DBH resistance could be delayed by including available synergists to the DBH formulations [146], such as metyrapone and diethylmaleate, which inhibit the activities of oxidative and glutathione-S-transferase enzymes, respectively [146].

Another important consideration before implementing a chemical control strategy based on 20E agonists is the phenomenon of cross-resistance. This occurs when insects are resistant to multiple insecticides because the insecticides share similar modes of action [147]. Studies in lepidopterans with methoxyfenozide and tebufenozide suggest that cross-resistance between DBH compounds is highly likely (Table 2), while cross-resistance between a DBH compound and the currently WHO-approved classes of insecticides (pyrethroids, organophosphates, carbamates, and organochlorines) is insecticide dependent. For example, while cross-resistance is observed between methoxyfenozide and deltamethrin, no such link is observed between methoxyfenozide and bifenthrin, although both deltamethrin and bifenthrin are pyrethroids (Table 2). In mosquitoes, cross-resistance between methoxyfenozide, pyrethroids, organochlorines (DDT) and carbamates has been characterized [148]. The authors of that study found that Anopheles populations which were resistant to DDT, carbamates and pyrethroids (regardless of the mechanism of pyrethroid resistance) were still susceptible to methoxyfenozide [148]. Collectively, these findings suggest that malaria vectors could be effectively controlled by a rotational strategy between DBHs and conventional insecticides, as part of an insecticide resistance management plan. Such a plan could, for example, involve (i) a rotation between pyrethroids and DBHs (with or without synergists/antimalarials) on LLINs, or (ii) a rotation between DBHs (with or without synergists/antimalarials), pyrethroids, DDT and carbamates for IRS and larvicides.

Another limitation of LLINs and IRS interventions is that they are designed to target mosquitoes indoors. Therefore, exophilic and exophagic vectors, such as An. arabiensis, are poorly controlled by these approaches [149]. To overcome this challenge, attractive toxic sugar baits (ATSB) have been proposed. The components of ATSB include a floral scent, a sugar solution and an oral insecticide, with the aim to attract, feed and kill mosquitoes, respectively [150, 151]. This technique has already been proven successful against An. gambiae and An. arabiensis populations in experimental trials [152, 153], and it would thus be worth investigating if the addition of DBH compounds to ATSB could enhance their efficacy. Additionally, one could also target exophilic/exophagic mosquitoes at the immature aquatic stages using DBH compounds (with or without synergists) as larvicides and ovicides (Table 1).

IGRs that reduce 20E titers and signaling

The 20E signaling pathway is also targeted by IGRs that interfere with its activity or reduce 20E titers. These include cucurbitacins (triterpenoid class of natural products) [154, 155], chlorantraniliprole (CAP; ryanoid class of pesticides) [156, 157] and clothianidin (neonicotinoid class of pesticides) [158]. CAP targets the insect calcium channels to deplete intracellular calcium [156, 157], while clothianidin targets the acetylcholine receptor and affects the insect immune system [158]. With respect to 20E biosynthesis and 20E signaling, a study in Drosophila showed that cucurbitacins are able to either displace a steroid hormone bound to EcR or prevent the formation of the EcR/USP heterodimer complex [154]. Application of CAP on Chilo suppressalis was shown to reduce vitellogenin expression and 20E titers and to increase the expression of three 20E biosynthetic enzymes (phantom, spook and shade), likely in response to the decrease in 20E levels [157, 159]. Finally, a study of the effect of clothianidin on Aphis gossypii revealed that this insecticide reduced vitellogenin and EcR expression [158].

As expected for insecticides that target 20E signaling, the phenotypes induced by these chemicals include impaired development (P. xylostella, B. mori, C. suppressalis), reduced fecundity (P. xylostella, C. suppressalis, A. gossypii) and mortality (P. xylostella, C. suppressalis, A. gossypii) [156,157,158,159,160]. However, despite these promising phenotypes, IGRs interfering with the 20E pathway may not be suitable for use against Anopheles vectors for multiple reasons. Firstly, unlike 20E agonists, clothianidin may be toxic to humans, rendering it unusable for public health [161]. Second, while 20E agonists have minimal effects on non-target species [162], CAP has shown adverse effects on honeybees, even at sublethal doses, and as such shows off-target effects on other important insects [163]. Third, Werling et al. [105] showed that a reduction in 20E signaling accelerates the P. falciparum sporogonic cycle in such a way that mosquitoes are able to transmit malaria sooner. Therefore, chemical control interventions targeting 20E signaling should rather focus on overactivating the pathway, as do the DBH compounds.

Concluding remarks

The development of novel interventions is urgently needed to counteract insecticide resistance in malaria vectors. In this review, we have summarized the importance of 20E throughout the mosquito life-cycle and consolidated some of the experimental evidence that supports the use of 20E agonists as part of an integrated approach to malaria vector control. Not only are 20E agonists already commercially available, but results from preliminary laboratory experiments suggest that they are effective against all mosquito life-stages (Table 1), with minimal toxicity to non-target species (Table 2). The efficacy of 20E agonists is mainly attributed to their ability to overactivate the 20E signaling pathway, a biological process which regulates vector abundance and competence in mosquitoes.

While the molecular mechanisms by which 20E signaling regulates mosquito reproduction and fecundity have been extensively studied, more research is needed to elucidate how 20E signaling (and 20E agonists) regulates Anopheles’ susceptibility to Plasmodium infection. From an experimental perspective, the timing of 20E injection (either before or after Plasmodium infection) influences whether or not the parasite’s sporogonic cycle is affected. Indeed, injection of 20E in An. gambiae 24 h prior to infection was found to result in a decrease in P. berghei oocyst prevalence and intensity [106], while there was no effect on these parameters when the injection occurred shortly after infection [164]. These divergent outcomes may result from a difference in the timing of 20E-regulated immune priming [106], and this would be worthwhile investigating.

Second, it is likely that DBH compounds (i.e. non-steroid 20E agonists) and 20E regulate Plasmodium development by distinct mechanisms, although they both bind to EcR. For example, even though exposure of An. gambiae to halofenozide and 20E both decrease P. berghei oocyst prevalence and intensity, the authors of these studies observed that only 20E induced the expression of immune genes [24, 106], therefore leaving unanswered the question of what could be the potential non-immune mechanisms by which 20E agonists regulate P. berghei competence. Possibilities include that DBH compounds regulate epigenetic modifications [165, 166], the formation of the peritrophic matrix [167], the expression of specific midgut factors that are essential for Plasmodium invasion [112], metabolism [142], signaling pathways (e.g. c-Jun N-terminal kinase [JNK] pathway), or all of these simultaneously. Clarifying these issues will help researchers to determine the full impact of 20E agonists on Anopheles vector competence.

Third, it is still unclear whether the male-derived 20E contributes to Anopheles susceptibility to P. falciparum (NF54 strain) infection. Dahalan et al. [125] showed that mating increased both P. falciparum oocyst prevalence and intensity in An. coluzzii. This effect was attributed to the male-to-female transfer of 20E, since 20E injection of virgin An. coluzzii produced similar results [125]. On the other hand, Marcenac et al. [168] reported no effect on parasite prevalence or intensity in mated An. gambiae and An. stephensi. Therefore, it is possible that the male-derived 20E is not solely responsible for the phenotypes observed but, rather, it acts in conjunction with other female genes that are affected by post-mating. Consistent with this notion, 13 genes (7 in the lower reproductive tract and 6 in the carcass) were found to be differentially expressed in females after mating in An. coluzzii versus An. gambiae [169]. Further studies should investigate whether one of these genes is responsible for the discrepancy observed between the findings of Dahalan et al. [125] and Marcenacet al. [168].

While we have presented vector competence and vector abundance as two separate entities that are each individually regulated by 20E through distinct mechanisms, in reality, both Plasmodium sporogony and vitellogenesis/egg production occur simultaneously in mosquitoes (reviewed in [170]). Moreover, the two processes are positively correlated in P. falciparum-infected An. gambiae mosquitoes [105]. This implies that the genes regulating anti-Plasmodium responses and those regulating fecundity are coordinately expressed after a mosquito takes an infected blood meal, and possibly co-regulated. Could it be that 20E acts as a “master regulator” which coordinates the timeous expression of genes involved in fecundity or immunity? If so, this further reinforces the premise discussed in this review that 20E is a worth-investigating target for the chemical control of malaria vectors.

Availability of data and materials

Not applicable.



World Health Organization


Ecdysone receptor




Hours post-blood meal


Yolk protein precursor




c-Jun N-terminal kinase pathway


Target of rapamycin


  1. 1.

    World Health Organization. World malaria report. Geneva: World Health Organization; 2020.

  2. 2.

    Hogan AB, Jewell BL, Sherrard-Smith E, Vesga JF, Watson OJ, Whittaker C, et al. Potential impact of the COVID-19 pandemic on HIV, tuberculosis, and malaria in low-income and middle-income countries: a modelling study. Lancet Glob Health. 2020;8(9):e1132–41.

    PubMed  PubMed Central  Article  Google Scholar 

  3. 3.

    Sinclair D, Zani B, Donegan S, Olliaro P, Garner P. Artemisinin‐based combination therapy for treating uncomplicated malaria. Cochranr DB Syst Rev. 2009;3:12.

  4. 4.

    Gutman JR, Chico RM. Evidence for treating malaria with artemisinin-based combination therapy in the first trimester of pregnancy. Lancet Infect Dis. 2020;5:8.

  5. 5.

    World Health Organization. Global plan for insecticide resistance management in malaria vectors. Geneva: World Health Organization; 2012.

  6. 6.

    World Health Organization. Guidelines for malaria vector control. Geneva: World Health Organization; 2019.

  7. 7.

    Utzinger J, Tozan Y, Singer BH. Efficacy and cost-effectiveness of environmental management for malaria control. Trop Med Int Health. 2001;6(9):677–87.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  8. 8.

    Castro MC, Tsuruta A, Kanamori S, Kannady K, Mkude S. Community-based environmental management for malaria control: evidence from a small-scale intervention in Dar es Salaam, Tanzania. Malaria J. 2009;8(1):57.

    Article  Google Scholar 

  9. 9.

    Randell HF, Dickinson KL, Shayo EH, Mboera LE, Kramer RA. Environmental management for malaria control: knowledge and practices in Mvomero, Tanzania. EcoHealth. 2010;7(4):507–16.

    PubMed  Article  PubMed Central  Google Scholar 

  10. 10.

    Manning JE, Oliveira F, Coutinho-Abreu IV, Herbert S, Meneses C, Kamhawi S, et al. Safety and immunogenicity of a mosquito saliva peptide-based vaccine: a randomised, placebo-controlled, double-blind, phase 1 trial. Lancet. 2020;395(10242):1998–2007.

  11. 11.

    Adepoju P. RTS, S malaria vaccine pilots in three African countries. Lancet. 2019;393(10182):1685.

    PubMed  Article  Google Scholar 

  12. 12.

    Laurens MB. RTS, S/AS01 vaccine (Mosquirix™): An overview. Hum Vaccines Immunother. 2020;16(3):480–9.

    CAS  Article  Google Scholar 

  13. 13.

    Rts S. Efficacy and safety of RTS, S/AS01 malaria vaccine with or without a booster dose in infants and children in Africa: final results of a phase 3, individually randomised, controlled trial. Lancet. 2015;386(9988):31–45.

    Article  CAS  Google Scholar 

  14. 14.

    World Health Organization. World malaria report 2018. Geneva: World Health Organization; 2018.

  15. 15.

    Fillinger U, Lindsay SW. Larval source management for malaria control in Africa: myths and reality. Malaria J. 2011;10(1):353.

    Article  Google Scholar 

  16. 16.

    Knox TB, Juma EO, Ochomo EO, Jamet HP, Ndungo L, Chege P, et al. An online tool for mapping insecticide resistance in major Anopheles vectors of human malaria parasites and review of resistance status for the Afrotropical region. Parasites Vectors. 2014;7(1):76.

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  17. 17.

    World Health Organization. Global report on insecticide resistance in malaria vectors: 2010–2016. Geneva: World Health Organization; 2018.

  18. 18.

    Dadzie SK, Chabi J, Asafu-Adjaye A, Owusu-Akrofi O, Baffoe-Wilmot A, Malm K, et al. Evaluation of piperonyl butoxide in enhancing the efficacy of pyrethroid insecticides against resistant Anopheles gambiae sl in Ghana. Malaria J. 2017;16(1):342.

    Article  Google Scholar 

  19. 19.

    Sahu SS, Dash S, Sonia T, Gunasekaran K. Synergist piperonyl butoxide enhances the efficacy of deltamethrin in deltamethrin-resistant Anopheles culicifacies sensu lato in malaria endemic districts of Odisha State, India. Indian J Med Res. 2019;149(4):554.

    PubMed  PubMed Central  Article  Google Scholar 

  20. 20.

    Gleave K, Lissenden N, Richardson M, Choi L, Ranson H. Piperonyl butoxide (PBO) combined with pyrethroids in insecticide‐treated nets to prevent malaria in Africa. Cochrane Database Syst Rev. 2018;2018:11.

  21. 21.

    Paton DG, Childs LM, Itoe MA, Holmdahl IE, Buckee CO, Catteruccia F. Exposing Anopheles mosquitoes to antimalarials blocks Plasmodium parasite transmission. Nature. 2019;567:239–43.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  22. 22.

    Nakagawa Y, Sonobe H. 20-Hydroxyecdysone. Handbook of hormones. Amsterdam: Elsevier; 2016.

  23. 23.

    Childs LM, Cai FY, Kakani EG, Mitchell SN, Paton D, Gabrieli P, et al. Disrupting mosquito reproduction and parasite development for malaria control. PLoS Pathog. 2016;12(12):e1006060.

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  24. 24.

    Reynolds RA, Smith RC. The 20-hydroxyecdysone agonist, halofenozide, primes anti-Plasmodium immunity in Anopheles gambiae via the ecdysone receptor. Sci Rep. 2020;10:21084.

  25. 25.

    Gabrieli P, Kakani EG, Mitchell SN, Mameli E, Want EJ, Anton AM, et al. Sexual transfer of the steroid hormone 20E induces the postmating switch in Anopheles gambiae. Proc Natl Acad Sci USA. 2014;111(46):16353–8.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  26. 26.

    Beckage NE, Marion KM, Walton WE, Wirth MC, Tan FF. Comparative larvicidal toxicities of three ecdysone agonists on the mosquitoes Aedes aegypti, Culex quinquefasciatus, and Anopheles gambiae. Arch Insect Biochem. 2004;57(3):111–22.

    CAS  Article  Google Scholar 

  27. 27.

    Morou E, Lirakis M, Pavlidi N, Zotti M, Nakagawa Y, Smagghe G, et al. A new dibenzoylhydrazine with insecticidal activity against Anopheles mosquito larvae. Pest Manag Sci. 2013;69(7):827–33.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  28. 28.

    Clayton RB. The utilization of sterols by insects. J Lipid Res. 1964;5(1):3–19.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  29. 29.

    Karlson P. Chemistry and biochemistry of insect hormones. Angew Chem Int Ed. 1963;2(4):175–82.

    Article  Google Scholar 

  30. 30.

    Rees HH. Ecdysteroidogenic pathway. In: Henry HL, Norman AW, editors. Encyclopedia of hormones. New York: Elsevier; 2003. p. 460–1.

    Chapter  Google Scholar 

  31. 31.

    Jenkins SP, Brown MR, Lea AO. Inactive prothoracic glands in larvae and pupae of Aedes aegypti: ecdysteroid release by tissues in the thorax and abdomen. Insect Biochem Mol Biol. 1992;22(6):553–9.

    CAS  Article  Google Scholar 

  32. 32.

    Pondeville E, Maria A, Jacques J-C, Bourgouin C, Dauphin-Villemant C. Anopheles gambiae males produce and transfer the vitellogenic steroid hormone 20-hydroxyecdysone to females during mating. Proc Natl Acad Sci USA. 2008;105(50):19631–6.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  33. 33.

    Raikhel A, Brown M, Belles X. 3.9 Hormonal control of reproductive processes. Comprehens Mol Insect Sci. 2005;3:433–91.

  34. 34.

    Raikhel AS, Kokoza VA, Zhu J, Martin D, Wang S-F, Li C, et al. Molecular biology of mosquito vitellogenesis: from basic studies to genetic engineering of antipathogen immunity. Insect Biochem Mol Biol. 2002;32(10):1275–86.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  35. 35.

    Roy S, Saha TT, Johnson L, Zhao B, Ha J, White KP, et al. Regulation of gene expression patterns in mosquito reproduction. PLoS Genet. 2015;11(8):e1005450.

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  36. 36.

    Yoshiyama-Yanagawa T, Enya S, Shimada-Niwa Y, Yaguchi S, Haramoto Y, Matsuya T, et al. The conserved Rieske oxygenase DAF-36/Neverland is a novel cholesterol-metabolizing enzyme. J Biol Chem. 2011;286(29):25756–62.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  37. 37.

    Gilbert LI, Rewitz KF. The function and evolution of the Halloween genes: the pathway to the arthropod molting hormone. In: Smagghe G, editor. Ecdysone: structures and functions. Dordrecht: Springer Netherlands; 2009. p. 231–69.

  38. 38.

    Vogel KJ, Valzania L, Coon KL, Brown MR, Strand MR. Transcriptome sequencing reveals large-scale changes in axenic Aedes aegypti larvae. PLoS Negl Trop Dis. 2017;11(1):e0005273.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  39. 39.

    Niwa R, Namiki T, Ito K, Shimada-Niwa Y, Kiuchi M, Kawaoka S, et al. Non-molting glossy/shroud encodes a short-chain dehydrogenase/reductase that functions in the ‘Black Box’ of the ecdysteroid biosynthesis pathway. Development. 2010;137(12):1991–9.

    CAS  PubMed  Article  Google Scholar 

  40. 40.

    Niwa R, Sakudoh T, Namiki T, Saida K, Fujimoto Y, Kataoka H. The ecdysteroidogenic P450 Cyp302a1/disembodied from the silkworm, Bombyx mori, is transcriptionally regulated by prothoracicotropic hormone. Insect Mol Biol. 2005;14(5):563–71.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  41. 41.

    Ono H, Rewitz KF, Shinoda T, Itoyama K, Petryk A, Rybczynski R, et al. Spook and Spookier code for stage-specific components of the ecdysone biosynthetic pathway in Diptera. Dev Biol. 2006;298(2):555–70.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  42. 42.

    Ou Q, Magico A, King-Jones K. Nuclear receptor DHR4 controls the timing of steroid hormone pulses during Drosophila development. PLoS Biol. 2011;9(9):e1001160.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  43. 43.

    Rewitz KF, O’Connor MB, Gilbert LI. Molecular evolution of the insect Halloween family of cytochrome P450s: phylogeny, gene organization and functional conservation. Insect Biochem Mol Biol. 2007;37(8):741–53.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  44. 44.

    Pondeville E, David J-P, Guittard E, Maria A, Jacques J-C, Ranson H, et al. Microarray and RNAi Analysis of P450s in Anopheles gambiae male and female steroidogenic tissues: CYP307A1 is required for ecdysteroid synthesis. PLoS One. 2013;8(12):e79861–6.

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  45. 45.

    Niwa R, Matsuda T, Yoshiyama T, Namiki T, Mita K, Fujimoto Y, et al. CYP306A1, a cytochrome P450 enzyme, is essential for ecdysteroid biosynthesis in the prothoracic glands of Bombyx and Drosophila. J Biol Chem. 2004;279(34):35942–9.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  46. 46.

    Warren JT, Petryk A, Marqués G, Parvy J-P, Shinoda T, Itoyama K, et al. Phantom encodes the 25-hydroxylase of Drosophila melanogaster and Bombyx mori: a P450 enzyme critical in ecdysone biosynthesis. Insect Biochem Mol Biol. 2004;34(9):991–1010.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  47. 47.

    Chávez VM, Marqués G, Delbecque JP, Kobayashi K, Hollingsworth M, Burr J, et al. The Drosophila disembodied gene controls late embryonic morphogenesis and codes for a cytochrome P450 enzyme that regulates embryonic ecdysone levels. Development. 2000;127(19):4115–26.

    PubMed  Article  PubMed Central  Google Scholar 

  48. 48.

    Warren JT, Petryk A, Marqués G, Jarcho M, Parvy J-P, Dauphin-Villemant C, et al. Molecular and biochemical characterization of two P450 enzymes in the ecdysteroidogenic pathway of Drosophila melanogaster. Proc Natl Acad Sci USA. 2002;99(17):11043–8.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  49. 49.

    Petryk A, Warren JT, Marqués G, Jarcho MP, Gilbert LI, Kahler J, et al. Shade is the Drosophila P450 enzyme that mediates the hydroxylation of ecdysone to the steroid insect molting hormone 20-hydroxyecdysone. Proc Natl Acad Sci USA. 2003;100(24):13773–8.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  50. 50.

    Oro AE, McKeown M, Evans RM. Relationship between the product of the Drosophila ultraspiracle locus and the vertebrate retinoid X receptor. Nature. 1990;347(6290):298–301.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  51. 51.

    Koelle MR, Talbot WS, Segraves WA, Bender MT, Cherbas P, Hogness DS. The Drosophila EcR gene encodes an ecdysone receptor, a new member of the steroid receptor superfamily. Cell. 1991;67(1):59–77.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  52. 52.

    Cho W-L, Kapitskaya MZ, Raikhel AS. Mosquito ecdysteroid receptor: analysis of the cDNA and expression during vitellogenesis. Insect Biochem Molec. 1995;25(1):19–27.

    CAS  Article  Google Scholar 

  53. 53.

    Kapitskaya M, Wang S, Cress DE, Dhadialla TS, Raikhel AS. The mosquito ultraspiracle homologue, a partner of ecdysteroid receptor heterodimer: cloning and characterization of isoforms expressed during vitellogenesis. Mol Cell Endocrinol. 1996;121(2):119–32.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  54. 54.

    Rowinska-Zyrek M, Wiȩch A, Wa̧tły J, Wieczorek R, Witkowska D, Ożyhar A, et al. Copper (II)-binding induces a unique polyproline type II helical structure within the ion-binding segment in the intrinsically disordered F-domain of ecdysteroid receptor from Aedes aegypti. Inorg Chem. 2019;58(17):11782–92.

  55. 55.

    Więch A, Rowińska-Żyrek M, Wątły J, Czarnota A, Hołubowicz R, Szewczuk Z, et al. The intrinsically disordered C-terminal F domain of the ecdysteroid receptor from Aedes aegypti exhibits metal ion-binding ability. J Steroid Biochem Mol Biol. 2019;186:42–55.

    PubMed  Article  CAS  PubMed Central  Google Scholar 

  56. 56.

    Yao T-P, Segraves WA, Oro AE, McKeown M, Evans RM. Drosophila ultraspiracle modulates ecdysone receptor function via heterodimer formation. Cell. 1992;71(1):63–72.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  57. 57.

    Kumar MB, Fujimoto T, Potter DW, Deng Q, Palli SR. A single point mutation in ecdysone receptor leads to increased ligand specificity: implications for gene switch applications. Proc Natl Acad Sci USA. 2002;99(23):14710–5.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  58. 58.

    Hu X, Cherbas L, Cherbas P. Transcription activation by the ecdysone receptor (EcR/USP): identification of activation functions. Mol Endocrinol. 2003;17(4):716–31.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  59. 59.

    Cherbas L, Lee K, Cherbas P. Identification of ecdysone response elements by analysis of the Drosophila Eip28/29 gene. Gene Dev. 1991;5(1):120–31.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  60. 60.

    Wang S-F, Miura K, Miksicek RJ, Segraves WA, Raikhel AS. DNA binding and transactivation characteristics of the mosquito ecdysone receptor-Ultraspiracle complex. J Biol Chem. 1998;273(42):27531–40.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  61. 61.

    Sun G, Zhu J, Li C, Tu Z, Raikhel AS. Two isoforms of the early E74 gene, an Ets transcription factor homologue, are implicated in the ecdysteroid hierarchy governing vitellogenesis of the mosquito, Aedes aegypti. Mol Cell Endocrinol. 2002;190(1–2):147–57.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  62. 62.

    Sun G, Zhu J, Raikhel AS. The early gene E74B isoform is a transcriptional activator of the ecdysteroid regulatory hierarchy in mosquito vitellogenesis. Mol Cell Endocrinol. 2004;218(1–2):95–105.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  63. 63.

    Margam VM, Gelman DB, Palli SR. Ecdysteroid titers and developmental expression of ecdysteroid-regulated genes during metamorphosis of the yellow fever mosquito, Aedes aegypti (Diptera: Culicidae). J Insect Physiol. 2006;52(6):558–68.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  64. 64.

    Chen L, Zhu J, Sun G, Raikhel A. The early gene Broad is involved in the ecdysteroid hierarchy governing vitellogenesis of the mosquito Aedes aegypti. J Mol Endocrinol. 2004;33(3):743–61.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  65. 65.

    Zhu J, Chen L, Raikhel AS. Distinct roles of broad isoforms in regulation of the 20-hydroxyecdysone effector gene, Vitellogenin, in the mosquito Aedes aegypti. Mol Cell Endocrinol. 2007;267(1):97–105.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  66. 66.

    Mane-Padros D, Cruz J, Cheng A, Raikhel AS. A critical role of the nuclear receptor HR3 in regulation of gonadotrophic cycles of the mosquito Aedes aegypti. PLoS One. 2012;7(9):e45019.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  67. 67.

    Kapitskaya MZ, Li C, Miura K, Segraves W, Raikhel AS. Expression of the early-late gene encoding the nuclear receptor HR3 suggests its involvement in regulating the vitellogenic response to ecdysone in the adult mosquito. Mol Cell Endocrinol. 2000;160(1–2):25–37.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  68. 68.

    Johnson LK. Regulation of gene expression patterns during reproduction in the female mosquito. Aedes aegypti: UC Riverside; 2018.

    Google Scholar 

  69. 69.

    Martı́n D, Wang S-F, Raikhel AS. The vitellogenin gene of the mosquito Aedes aegypti is a direct target of ecdysteroid receptor. Mol Cell Endocrinol. 2001;173(1–2):75–86.

  70. 70.

    Cruz J, Sieglaff DH, Arensburger P, Atkinson PW, Raikhel AS. Nuclear receptors in the mosquito Aedes aegypti. FEBS J. 2009;276(5):1233–54.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  71. 71.

    Parthasarathy R, Palli SR. Stage-and cell-specific expression of ecdysone receptors and ecdysone-induced transcription factors during midgut remodeling in the yellow fever mosquito, Aedes aegypti. J Insect Physiol. 2007;53(3):216–29.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  72. 72.

    Wang S-F, Li C, Sun G, Zhu J, Raikhel AS. Differential expression and regulation by 20-hydroxyecdysone of mosquito ecdysteroid receptor isoforms A and B. Mol Cell Endocrinol. 2002;196(1–2):29–42.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  73. 73.

    Wang S-F, Li C, Zhu J, Miura K, Miksicek RJ, Raikhel AS. Differential expression and regulation by 20-hydroxyecdysone of mosquito ultraspiracle isoforms. Dev Biol. 2000;218(1):99–113.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  74. 74.

    Wu Y, Parthasarathy R, Bai H, Palli SR. Mechanisms of midgut remodeling: juvenile hormone analog methoprene blocks midgut metamorphosis by modulating ecdysone action. Mech Dev. 2006;123(7):530–47.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  75. 75.

    Abrieux A, Debernard S, Maria A, Gaertner C, Anton S, Gadenne C, et al. Involvement of the G-protein-coupled dopamine/ecdysteroid receptor DopEcR in the behavioral response to sex pheromone in an insect. PLoS One. 2013;8(9):e72785.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  76. 76.

    Takeuchi H, Paul RK, Matsuzaka E, Kubo T. EcR-A expression in the brain and ovary of the honeybee (Apis mellifera L.). Zool Sci. 2007;24(6):596–603.

  77. 77.

    Hossain M, Shimizu S, Fujiwara H, Sakurai S, Iwami M. EcR expression in the prothoracicotropic hormone-producing neurosecretory cells of the Bombyx mori brain: an indication of the master cells of insect metamorphosis. FEBS J. 2006;273(16):3861–8.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  78. 78.

    Zhu J, Miura K, Chen L, Raikhel AS. AHR38, a homolog of NGFI-B, inhibits formation of the functional ecdysteroid receptor in the mosquito Aedes aegypti. EMBO J. 2000;19(2):253–62.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  79. 79.

    Cruz J, Mane-Padros D, Zou Z, Raikhel AS. Distinct roles of isoforms of the heme-liganded nuclear receptor E75, an insect ortholog of the vertebrate Rev-erb, in mosquito reproduction. Mol Cell Endocrinol. 2012;349(2):262–71.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  80. 80.

    Zhu J, Chen L, Sun G, Raikhel AS. The competence factor βFtz-F1 potentiates ecdysone receptor activity via recruiting a p160/SRC coactivator. Mol Cell Biol. 2006;26(24):9402–12.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  81. 81.

    Gaddelapati SC, Dhandapani RK, Palli SR. CREB-binding protein regulates metamorphosis and compound eye development in the yellow fever mosquito, Aedes aegypti. Biochim Biophys Acta Gene Regul Mech. 2020:194:576.

  82. 82.

    Williams J, Pinto J. Training manual on malaria entomology for entomology and vector control technicians (basic level). Washington DC: U.S. Agency for International Development; 2012.

    Google Scholar 

  83. 83.

    Clements A, Stanley-Samuelson DW. The biology of mosquitoes, volume 1: development, nutrition, and reproduction. J Med Entomol. 1994;31(1):181.

    Article  Google Scholar 

  84. 84.

    Scaraffia PY. Disruption of mosquito blood meal protein metabolism. In: Adelman ZN, editor. Genetic control of malaria and dengue. New York: Elsevier; 2016. p. 253–75.

  85. 85.

    Hou Y, Wang X-L, Saha TT, Roy S, Zhao B, Raikhel AS, et al. Temporal coordination of carbohydrate metabolism during mosquito reproduction. PLoS Genet. 2015;11(7):e1005309.

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  86. 86.

    Dong D, Zhang Y, Smykal V, Ling L, Raikhel AS. HR38, an ortholog of NR4A family nuclear receptors, mediates 20-hydroxyecdysone regulation of carbohydrate metabolism during mosquito reproduction. Insect Biochem Mol Biol. 2018;96:19–26.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  87. 87.

    Wang X, Hou Y, Saha TT, Pei G, Raikhel AS, Zou Z. Hormone and receptor interplay in the regulation of mosquito lipid metabolism. Proc Natl Acad Sci USA. 2017:2016:19326–26.

  88. 88.

    Ziegler R, Ibrahim MM. Formation of lipid reserves in fat body and eggs of the yellow fever mosquito, Aedes aegypti. J Insect Physiol. 2001;47(6):623–7.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  89. 89.

    Brown MR, Clark KD, Gulia M, Zhao Z, Garczynski SF, Crim JW, et al. An insulin-like peptide regulates egg maturation and metabolism in the mosquito Aedes aegypti. Proc Natl Acad Sci USA. 2008;105(15):5716–21.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  90. 90.

    Brown MR, Graf R, Swiderek KM, Fendley D, Stracker TH, Champagne DE, et al. Identification of a steroidogenic neurohormone in female mosquitoes. J Biol Chem. 1998;273(7):3967–71.

    CAS  PubMed  Article  Google Scholar 

  91. 91.

    Riehle MA, Brown MR. Insulin receptor expression during development and a reproductive cycle in the ovary of the mosquito Aedes aegypti. Cell Tissue Res. 2002;308(3):409–20.

    CAS  PubMed  Article  Google Scholar 

  92. 92.

    Vogel KJ, Brown MR, Strand MR. Ovary ecdysteroidogenic hormone requires a receptor tyrosine kinase to activate egg formation in the mosquito Aedes aegypti. Proc Natl Acad Sci USA. 2015;112(16):5057–62.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  93. 93.

    Valzania L, Mattee MT, Strand MR, Brown MR. Blood feeding activates the vitellogenic stage of oogenesis in the mosquito Aedes aegypti through inhibition of glycogen synthase kinase 3 by the insulin and TOR pathways. Dev Biol. 2019;454(1):85–95.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  94. 94.

    Capurro MdL, De Bianchi A, Marinotti O. Aedes aegypti lipophorin. Comp Biochem Physiol B Comp Biochem. 1994;108(1):35–39.

  95. 95.

    Van Heusden MC, Erickson BA, Pennington JE. Lipophorin levels in the yellow fever mosquito, Aedes aegypti, and the effect of feeding. Arch Insect Biochem Physiol. 1997;34(3):301–12.

    PubMed  Article  Google Scholar 

  96. 96.

    Carpenter VK, Drake LL, Aguirre SE, Price DP, Rodriguez SD, Hansen IA. SLC7 amino acid transporters of the yellow fever mosquito Aedes aegypti and their role in fat body TOR signaling and reproduction. J Insect Physiol. 2012;58(4):513–22.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  97. 97.

    Attardo GM, Hansen IA, Shiao S-H, Raikhel AS. Identification of two cationic amino acid transporters required for nutritional signaling during mosquito reproduction. J Exp Biol. 2006;209(16):3071.

    CAS  PubMed  Article  Google Scholar 

  98. 98.

    Boudko DY, Tsujimoto H, Rodriguez SD, Meleshkevitch EA, Price DP, Drake LL, et al. Substrate specificity and transport mechanism of amino-acid transceptor Slimfast from Aedes aegypti. Nat Commun. 2015;6(1):8546.

    CAS  PubMed  Article  Google Scholar 

  99. 99.

    Carney GE, Bender M. The Drosophila ecdysone receptor (EcR) gene is required maternally for normal oogenesis. Genetics. 2000;154(3):1203–11.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  100. 100.

    Fournet F, Sannier C, Moriniere M, Porcheron P, Monteny N. Effects of two insect growth regulators on ecdysteroid production in Aedes aegypti (Diptera: Culicidae). J Med Entomol. 1995;32(5):588–93.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  101. 101.

    Lan Q, Grier CA. Critical period for pupal commitment in the yellow fever mosquito, Aedes aegypti. J Insect Physiol. 2004;50(7):667–76.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  102. 102.

    Whisenton LR, Warren JT, Manning MK, Bollenbacher WE. Ecdysteroid titres during pupal-adult development of Aedes aegypti: basis for a sexual dimorphism in the rate of development. J Insect Physiol. 1989;35(1):67–73.

    CAS  Article  Google Scholar 

  103. 103.

    Fujiwara H, Ogai S. Ecdysteroid-induced programmed cell death and cell proliferation during pupal wing development of the silkworm, Bombyx mori. Dev Genes Evol. 2001;211:3.

  104. 104.

    Haouari-Abderrahim J, Rehimi N. Biological and reproduction activities of mosquito larvae of Culiseta morsitans (Theobald) after treatment by ecdysone agonist methoxyfenozide. Annu Res Rev Biol. 2014:4152–65.

  105. 105.

    Werling K, Shaw WR, Itoe MA, Westervelt KA, Marcenac P, Paton DG, et al. Steroid hormone function controls non-competitive Plasmodium development in anopheles. Cell. 2019;177(2):315–25.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  106. 106.

    Reynolds RA, Kwon H, Smith RC. 20-hydroxyecdysone (20E) primes innate immune responses that limit bacteria and malaria parasite survival in Anopheles gambiae. mSphere. 2020;5(2):e00983–19.

  107. 107.

    Ye W, Liu X, Guo J, Sun X, Sun Y, Shen B, et al. piRNA-3878 targets P450 (CpCYP307B1) to regulate pyrethroid resistance in Culex pipiens pallens. Parasitol Res. 2017;116(9):2489–97.

    PubMed  Article  PubMed Central  Google Scholar 

  108. 108.

    Sun X, Xu N, Xu Y, Zhou D, Sun Y, Wang W, et al. A novel miRNA, miR-13664, targets CpCYP314A1 to regulate deltamethrin resistance in Culex pipiens pallens. Parasitology. 2019;146(2):197.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  109. 109.

    Beach R. Mosquitoes: biting behavior inhibited by ecdysone. Science. 1979;205(4408):829–31.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  110. 110.

    Simonetti AB. The biology of malarial parasite in the mosquito: a review. Mem I Oswaldo Cruz. 1996;91(5):519–41.

    CAS  Article  Google Scholar 

  111. 111.

    Vlachou D, Schlegelmilch T, Runn E, Mendes A, Kafatos FC. The developmental migration of Plasmodium in mosquitoes. Curr Opin Genet Dev. 2006;16(4):384–91.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  112. 112.

    Bennink S, Kiesow MJ, Pradel G. The development of malaria parasites in the mosquito midgut. Cell Microbiol. 2016;18(7):905–18.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  113. 113.

    Stone WJ, Eldering M, van Gemert G-J, Lanke KH, Grignard L, van de Vegte-Bolmer MG, et al. The relevance and applicability of oocyst prevalence as a read-out for mosquito feeding assays. Sci Rep. 2013;3:3418.

    PubMed  PubMed Central  Article  Google Scholar 

  114. 114.

    Miura K, Swihart BJ, Deng B, Zhou L, Pham TP, Diouf A, et al. Transmission-blocking activity is determined by transmission-reducing activity and number of control oocysts in Plasmodium falciparum standard membrane-feeding assay. Vaccine. 2016;34(35):4145–51.

    PubMed  PubMed Central  Article  Google Scholar 

  115. 115.

    Ohm JR, Baldini F, Barreaux P, Lefevre T, Lynch PA, Suh E, et al. Rethinking the extrinsic incubation period of malaria parasites. Parasites Vectors. 2018;11(1):1–9.

    Article  Google Scholar 

  116. 116.

    Shaw WR, Catteruccia f. Vector biology meets disease control: using basic research to fight vector-borne diseases. Nat Microbiol. 2018;1:1.

    Google Scholar 

  117. 117.

    Vontas J, Blass C, Koutsos A, David JP, Kafatos F, Louis C, et al. Gene expression in insecticide resistant and susceptible Anopheles gambiae strains constitutively or after insecticide exposure. Insect Mol Biol. 2005;14(5):509–21.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  118. 118.

    Riveron JM, Ibrahim SS, Mulamba C, Djouaka R, Irving H, Wondji MJ, et al. Genome-wide transcription and functional analyses reveal heterogeneous molecular mechanisms driving pyrethroids resistance in the major malaria vector Anopheles funestus across Africa. G3 (Bethesda). 2017;7(6):1819–32.

  119. 119.

    Kang X-L, Zhang J-Y, Di Wang Y-MZ, Han X-L, Wang J-X, Zhao X-F. The steroid hormone 20-hydroxyecdysone binds to dopamine receptor to repress lepidopteran insect feeding and promote pupation. PLoS Genet. 2019;15:8.

  120. 120.

    Wang S, Liu S, Liu H, Wang J, Zhou S, Jiang R-J, et al. 20-Hydroxyecdysone reduces insect food consumption resulting in fat body lipolysis during molting and pupation. J Mol Cell Biol. 2010;2(3):128–38.

    PubMed  Article  CAS  Google Scholar 

  121. 121.

    Landayan D, Feldman DS, Wolf FW. Satiation state-dependent dopaminergic control of foraging in Drosophila. Sci Rep. 2018;8(1):1–9.

    CAS  Article  Google Scholar 

  122. 122.

    Mitchell SN, Kakani EG, South A, Howell PI, Waterhouse RM, Catteruccia F. Evolution of sexual traits influencing vectorial capacity in anopheline mosquitoes. Science. 2015;347(6225):985–8.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  123. 123.

    Baldini F, Gabrieli P, South A, Valim C, Mancini F, Catteruccia F. The interaction between a sexually transferred steroid hormone and a female protein regulates oogenesis in the malaria mosquito Anopheles gambiae. PLoS Biol. 2013;11(10):e1001695.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  124. 124.

    Shaw WR, Teodori E, Mitchell SN, Baldini F, Gabrieli P, Rogers DW, et al. Mating activates the heme peroxidase HPX15 in the sperm storage organ to ensure fertility in Anopheles gambiae. Proc Natl Acad Sci USA. 2014;111(16):5854–9.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  125. 125.

    Dahalan FA, Churcher TS, Windbichler N, Lawniczak MKN. The male mosquito contribution towards malaria transmission: mating influences the Anopheles female midgut transcriptome and increases female susceptibility to human malaria parasites. PLoS Pathog. 2019;15:11.

  126. 126.

    Dhadialla TS, Carlson GR, Le DP. New insecticides with ecdysteroidal and juvenile hormone activity. Annu Rev Entomol. 1998;43(1):545–69.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  127. 127.

    Smagghe G, Zotti M, Retnakaran A. Targeting female reproduction in insects with biorational insecticides for pest management: a critical review with suggestions for future research. Curr Opin Insect Sci. 2019;31:65–9.

    PubMed  Article  PubMed Central  Google Scholar 

  128. 128.

    Nakagawa Y. Nonsteroidal ecdysone agonists. Vitam Horm. 2005;73:131–73.

    CAS  PubMed  Article  Google Scholar 

  129. 129.

    Wing KD, Slawecki RA, Carlson GR. RH 5849, a nonsteroidal ecdysone agonist: effects on larval Lepidoptera. Science. 1988;241(4864):470–2.

    CAS  PubMed  Article  Google Scholar 

  130. 130.

    Carlson GR, Dhadialla TS, Hunter R, Jansson RK, Jany CS, Lidert Z, et al. The chemical and biological properties of methoxyfenozide, a new insecticidal ecdysteroid agonist. Pest Manag Sci. 2001;57(2):115–9.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  131. 131.

    Hamaidia K, Soltani N. Ovicidal activity of an insect growth disruptor (methoxyfenozide) against Culex pipiens L. and delayed effect on development. J Entomol Zool. 2016;4:4.

  132. 132.

    Tateishi K, Kiuchi M, Takeda S. New cuticle formation and molt inhibition by RH-5849 in the common cutworm, Spodoptera litura (Lepidoptera: Noctuidae). Appl Entomol Zool. 1993;28(2):177–84.

    CAS  Article  Google Scholar 

  133. 133.

    Cao G, Han Z. Tebufenozide resistance selected in Plutella xylostella and its cross-resistance and fitness cost. Pest Manag Sci. 2006;62(8):746–51.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  134. 134.

    Qian L, Cao G, Song J, Yin Q, Han Z. Biochemical mechanisms conferring cross-resistance between tebufenozide and abamectin in Plutella xylostella. Pestic Biochem Physiol. 2008;91(3):175–9.

    CAS  Article  Google Scholar 

  135. 135.

    Mota‐Sanchez D, Wise JC, Poppen RV, Gut LJ, Hollingworth RM. Resistance of codling moth, Cydia pomonella (L.) (Lepidoptera: Tortricidae), larvae in Michigan to insecticides with different modes of action and the impact on field residual activity. Pest Manag Sci. 2008;64(9):881–90.

  136. 136.

    Jia B, Liu Y, Zhu YC, Liu X, Gao C, Shen J. Inheritance, fitness cost and mechanism of resistance to tebufenozide in Spodoptera exigua (Hübner) (Lepidoptera: Noctuidae). Pest Manag Sci. 2009;65(9):996–1002.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  137. 137.

    Sun J, Liang P, Gao X. Cross-resistance patterns and fitness in fufenozide-resistant diamondback moth, Plutella xylostella (Lepidoptera: Plutellidae). Pest Manag Sci. 2012;68(2):285–9.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  138. 138.

    Tang B, Sun J, Zhou X, Gao X, Liang P. The stability and biochemical basis of fufenozide resistance in a laboratory-selected strain of Plutella xylostella. Pestic Biochem Physiol. 2011;101(2):80–5.

    CAS  Article  Google Scholar 

  139. 139.

    Mosallanejad H, Badisco L, Swevers L, Soin T, Knapen D, Broeck JV, et al. Ecdysone signaling and transcript signature in Drosophila cells resistant against methoxyfenozide. J Insect Physiol. 2010;56(12):1973–85.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  140. 140.

    Mosallanejad H, Smagghe G. Biochemical mechanisms of methoxyfenozide resistance in the cotton leafworm Spodoptera littoralis. Pest Manag Sci. 2009;65(7):732–6.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  141. 141.

    Yin Q, Qian L, Song P, Jian T, Han Z. Molecular mechanisms conferring asymmetrical cross-resistance between tebufenozide and abamectin in Plutella xylostella. J Asia Pac Entomol. 2019;22(1):189–93.

    Article  Google Scholar 

  142. 142.

    Adedeji EO, Ogunlana OO, Fatumo S, Beder T, Ajamma Y, Koenig R, et al. Anopheles metabolic proteins in malaria transmission, prevention and control: a review. Parasites Vector. 2020;13(1):1–30.

    Article  CAS  Google Scholar 

  143. 143.

    Li X, Ren X, Liu Y, Smagghe G, Liang P, Gao X. MiR-189942 regulates fufenozide susceptibility by modulating ecdysone receptor isoform B in Plutella xylostella (L.). Pestic Biochem Physiol. 2020;163:235–40.

  144. 144.

    Rehan A, Freed S. Fitness cost of methoxyfenozide and the effects of its sublethal doses on development, reproduction, and survival of Spodoptera litura (Fabricius)(Lepidoptera: Noctuidae). Neotrop Entomol. 2015;44(5):513–20.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  145. 145.

    Shah RM, Shad SA, Abbas N. Methoxyfenozide resistance of the housefly, Musca domestica L.(Diptera: Muscidae): cross‐resistance patterns, stability and associated fitness costs. Pest Manag Sci. 2017;73(1):254–61.

  146. 146.

    Smagghe G. Synergism of diacylhydrazine insecticides with metyrapone and diethylmaleate. J Appl Entomol. 2004;128(7):465–8.

    CAS  Article  Google Scholar 

  147. 147.

    Rogier C, Henry M, Rowland M, Carnevale P, Chandre F, Corbel V, et al. Guidelines for phase III evaluation of vector control methods against malaria. Med Trop. 2009;69(2):173–84.

    CAS  Google Scholar 

  148. 148.

    Brown F, Paton DG, Catteruccia F, Ranson H, Ingham VA. A steroid hormone agonist reduces female fitness in insecticide-resistant Anopheles populations. Insect Biochem Mol Biol. 2020;121:103372.

  149. 149.

    Sougoufara S, Ottih EC, Tripet F. The need for new vector control approaches targeting outdoor biting Anopheline malaria vector communities. Parasites Vectors. 2020;13(1):1–15.

    Article  Google Scholar 

  150. 150.

    Fiorenzano JM, Koehler PG, Xue R-D. Attractive toxic sugar bait (ATSB) for control of mosquitoes and its impact on non-target organisms: a review. Int J Environ Res Public Health. 2017;14(4):398.

    PubMed Central  Article  CAS  Google Scholar 

  151. 151.

    Kline DL, Muller GC, Junnila A, Xue R-d. Attractive toxic sugar baits (ATSB): a novel vector management tool. In: Norris EJ, Coats JR, Gross AD, Clark JM, editors. Advances in the biorational control of medical and veterinary pests. Washington, DC: ACS Publications; 2018. p. 63–73.

    Chapter  Google Scholar 

  152. 152.

    Tenywa FC, Kambagha A, Saddler A, Maia MF. The development of an ivermectin-based attractive toxic sugar bait (ATSB) to target Anopheles arabiensis. Malaria J. 2017;16(1):1–10.

    Article  CAS  Google Scholar 

  153. 153.

    Müller GC, Beier JC, Traore SF, Toure MB, Traore MM, Bah S, et al. Successful field trial of attractive toxic sugar bait (ATSB) plant-spraying methods against malaria vectors in the Anopheles gambiae complex in Mali, West Africa. Malaria J. 2010;9(1):210.

    Article  CAS  Google Scholar 

  154. 154.

    Dinan L, Whiting P, Girault J-P, Lafont R, Dhadialla ST, Cress ED, et al. Cucurbitacins are insect steroid hormone antagonists acting at the ecdysteroid receptor. Biochem J. 1997;327(3):643–50.

  155. 155.

    Zou C, Liu G, Liu S, Liu S, Song Q, Wang J, et al. Cucurbitacin B acts a potential insect growth regulator by antagonizing 20-hydroxyecdysone activity. Pest Manag Sci. 2018;74(6):1394–403.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  156. 156.

    Chen J, Lu Z, Li M, Mao T, Wang H, Li F, et al. The mechanism of sublethal chlorantraniliprole exposure causing silkworm pupation metamorphosis defects. Pest Manag Sci. 2020;11:851.

  157. 157.

    Meng X, Zhang N, Yang X, Miao L, Jiang H, Ji C, et al. Sublethal effects of chlorantraniliprole on molting hormone levels and mRNA expressions of three Halloween genes in the rice stem borer, Chilo suppressalis. Chemosphere. 2020;238:124676.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  158. 158.

    Ullah F, Gul H, Desneux N, Tariq K, Ali A, Gao X, et al. Clothianidin-induced sublethal effects and expression changes of vitellogenin and ecdysone receptors genes in the melon aphid, Aphis gossypii. Entomol Gen. 2019;39(2):137–49.

    Article  Google Scholar 

  159. 159.

    Huang L, Lu M, Han G, Du Y, Wang J. Sublethal effects of chlorantraniliprole on development, reproduction and vitellogenin gene (CsVg) expression in the rice stem borer, Chilo suppressalis. Pest Manag Sci. 2016;72(12):2280–6.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  160. 160.

    Han W, Zhang S, Shen F, Liu M, Ren C, Gao X. Residual toxicity and sublethal effects of chlorantraniliprole on Plutella xylostella (Lepidoptera: Plutellidae). Pest Manag Sci. 2012;68(8):1184–90.

    CAS  PubMed  Article  Google Scholar 

  161. 161.

    Di Prisco G, Iannaccone M, Ianniello F, Ferrara R, Caprio E, Pennacchio F, et al. The neonicotinoid insecticide clothianidin adversely affects immune signaling in a human cell line. Sci Rep. 2017;7(1):1–8.

    Article  CAS  Google Scholar 

  162. 162.

    Loetti V, Bellocq I. Effects of the insecticides methoxyfenozide and cypermethrin on non-target arthropods: a field experiment. Austral Entomol. 2017;56(3):255–60.

    Article  Google Scholar 

  163. 163.

    Kadala A, Charreton M, Charnet P, Collet C. Honey bees long-lasting locomotor deficits after exposure to the diamide chlorantraniliprole are accompanied by brain and muscular calcium channels alterations. Sci Rep. 2019;9(1):1–9.

    Google Scholar 

  164. 164.

    Yang J, Schleicher TR, Dong Y, Park HB, Lan J, Cresswell P, et al. Disruption of mosGILT in Anopheles gambiae impairs ovarian development and Plasmodium infection. J Exp Med. 2020;217:1.

  165. 165.

    Claudio-Piedras F, Recio-Tótoro B, Condé R, Hernández-Tablas JM, Hurtado-Sil G, Lanz-Mendoza H. DNA methylation in Anopheles albimanus modulates the midgut immune response against Plasmodium berghei. Front Immunol. 2020;10:3025.

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  166. 166.

    Kirilly D, Wong JJL, Lim EKH, Wang Y, Zhang H, Wang C, et al. Intrinsic epigenetic factors cooperate with the steroid hormone ecdysone to govern dendrite pruning in Drosophila. Neuron. 2011;72(1):86–100.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  167. 167.

    Rodgers FH, Gendrin M, Wyer CA, Christophides GK. Microbiota-induced peritrophic matrix regulates midgut homeostasis and prevents systemic infection of malaria vector mosquitoes. PLoS Pathog. 2017;13(5):e1006391.

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  168. 168.

    Marcenac P, Shaw WR, Kakani EG, Mitchell SN, South A, Werling K, et al. A mating-induced reproductive gene promotes Anopheles tolerance to Plasmodium falciparum infection. PLoS Pathog. 2020;16(12):e1008908.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  169. 169.

    Thailayil J, Gabrieli P, Caputo B, Bascuñán P, South A, Diabate A, et al. Analysis of natural female post-mating responses of Anopheles gambiae and Anopheles coluzzii unravels similarities and differences in their reproductive ecology. Sci Rep. 2018;8(1):1–10.

    CAS  Article  Google Scholar 

  170. 170.

    Mitchell SN, Catteruccia F. Anopheline reproductive biology: impacts on vectorial capacity and potential avenues for malaria control. Cold Spring Harb Perspect Med. 2016;25593(1):15.

    Google Scholar 

  171. 171.

    Rosenberg R. Ovarian control of blood meal retention the the mosquito Anopheles freeborni. J Insect Physiol. 1980;26(7):477–80.

    Article  Google Scholar 

  172. 172.

    Bryant B, Raikhel AS. Programmed autophagy in the fat body of Aedes aegypti is required to maintain egg maturation cycles. PloS One. 2011;6(11):e25502.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  173. 173.

    Bouaziz A, Amira K, Djeghader N, Aïssaoui L, Boudjelida H. Impact of an insect growth regulator on the development and the reproduction potency of mosquito. J Entomol Zool. 2017;5(3):1662–7.

    Google Scholar 

  174. 174.

    Amira K, Boudjelida H, Farine J-P. Effect of an insect growth regulator (halofenozide) on the cuticular hydrocarbons of Culex pipiens larvae. Afr Entomol. 2013;21(2):343–8.

    Article  Google Scholar 

  175. 175.

    Boudjelida H, Bouaziz A, Soin T, Smagghe G, Soltani N. Effects of ecdysone agonist halofenozide against Culex pipiens. Pestic Biochem Phys. 2005;83(2–3):115–23.

    CAS  Article  Google Scholar 

  176. 176.

    Fei X, Zhang Y, Ding L, Li Y, Deng X. Controlling the development of the dengue vector Aedes aegypti using HR3 RNAi transgenic Chlamydomonas. PLoS One. 2020;15(10):e0240223.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  177. 177.

    Kamimura M, Saito H, Niwa R, Niimi T, Toyoda K, Ueno C, et al. Fungal ecdysteroid-22-oxidase, a new tool for manipulating ecdysteroid signaling and insect development. J Biol Chem. 2012;287(20):16488–98.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  178. 178.

    Dunley JE, Brunner JF, Doerr MD, Beers E. Resistance and cross-resistance in populations of the leafrollers, Choristoneura rosaceana and Pandemis pyrusana, in Washington apples. J Insect Sci. 2006;6(1):14.

    PubMed Central  PubMed  Google Scholar 

  179. 179.

    Smirle MJ, Thomas Lowery D, Zurowski CL. Resistance and cross-resistance to four insecticides in populations of obliquebanded leafroller (Lepidoptera: Tortricidae). J Econ Entomol. 2002;95(4):820–5.

    CAS  PubMed  Article  Google Scholar 

  180. 180.

    Rehan A, Freed S. Resistance selection, mechanism and stability of Spodoptera litura (Lepidoptera: Noctuidae) to methoxyfenozide. Pestic Biochem Physiol. 2014;110:7–12.

    CAS  PubMed  Article  Google Scholar 

  181. 181.

    Stacke RF, Godoy DN, Pretto VE, Führ FM, Gubiani PdS, Hettwer BL, et al. Field-evolved resistance to chitin synthesis inhibitor insecticides by soybean looper, Chrysodeixis includens (Lepidoptera: Noctuidae), in Brazil. Chemosphere. 2020:127499.

  182. 182.

    Cui L, Wang Q, Qi H, Wang Q, Yuan H, Rui C. Resistance selection of indoxacarb in Helicoverpa armigera (Hübner) (Lepidoptera: Noctuidae): cross-resistance, biochemical mechanisms and associated fitness costs. Pest Manag Sci. 2018;74(11):2636–44.

    CAS  PubMed  Article  Google Scholar 

  183. 183.

    Meikle WG, Corby-Harris V, Carroll MJ, Weiss M, Snyder LA, Meador CA, et al. Exposure to sublethal concentrations of methoxyfenozide disrupts honey bee colony activity and thermoregulation. PLoS One. 2019;14(3):e0204635.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  184. 184.

    Soltani N, Chouahda S, Smagghe G. Evaluation of halofenozide against prey mosquito larvae Culex pipiens and the predator fish Gambusia affinis: impact on growth and enzymatic activities. Comm Appl Biol Ghent Univ. 2008;73(3):659–66.

    CAS  Google Scholar 

  185. 185.

    Feng S, Kong Z, Wang X, Peng P, Zeng EY. Assessing the genotoxicity of imidacloprid and RH-5849 in human peripheral blood lymphocytes in vitro with comet assay and cytogenetic tests. Ecotoxicol Environ Saf. 2005;61(2):239–46.

    CAS  PubMed  Article  Google Scholar 

  186. 186.

    Jiang J, Shan Z, Wang X, Zhu Y, Zhou J. Ecotoxicity of the nonsteroidal ecdysone mimic RH-5849 to Daphnia magna. Environ Sci Pollut Res. 2018;25(11):10730–9.

    CAS  Article  Google Scholar 

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The degree (PhD) program from which this study emanated was funded by the South African Medical Research Council through its Division of Research Capacity Development under the Internship Scholarship Programme from funding received from the South African National Treasury. The content hereof is the sole responsibility of the authors and does not necessarily represent the official views of the SAMRC or the funders. The authors would also like to thank Ms. Marilyn Ekoka (Graphic Designer) for drawing the mosquitoes and larvae that appear in Fig. 2.


This study was funded by the South African National Research Foundation (DST/NRF) Chairs Initiative Grant (Grant No: 171215294399) and partial funding Communities of Practice grant from SARCHI (110666) awarded to LLK. In addition, funding was also received from the University of the Witwatersrand Medical Faculty Research Endowment Fund (Grant No: 0012548469201512110500000000000000005254) awarded to EE.

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EE conceived the review. EE and SM wrote the first draft. EE designed and drew the figures. EE, LN, YDM and LLK revised and contributed to the subsequent versions. All authors read and approved the final manuscript.

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Correspondence to Elodie Ekoka.

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Ekoka, E., Maharaj, S., Nardini, L. et al. 20-Hydroxyecdysone (20E) signaling as a promising target for the chemical control of malaria vectors. Parasites Vectors 14, 86 (2021).

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