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Ebihara, K.; Niwa, R. Compounds Inhibiting Noppera-bo as Insect Growth Regulators. Encyclopedia. Available online: https://encyclopedia.pub/entry/44000 (accessed on 21 December 2024).
Ebihara K, Niwa R. Compounds Inhibiting Noppera-bo as Insect Growth Regulators. Encyclopedia. Available at: https://encyclopedia.pub/entry/44000. Accessed December 21, 2024.
Ebihara, Kana, Ryusuke Niwa. "Compounds Inhibiting Noppera-bo as Insect Growth Regulators" Encyclopedia, https://encyclopedia.pub/entry/44000 (accessed December 21, 2024).
Ebihara, K., & Niwa, R. (2023, May 09). Compounds Inhibiting Noppera-bo as Insect Growth Regulators. In Encyclopedia. https://encyclopedia.pub/entry/44000
Ebihara, Kana and Ryusuke Niwa. "Compounds Inhibiting Noppera-bo as Insect Growth Regulators." Encyclopedia. Web. 09 May, 2023.
Compounds Inhibiting Noppera-bo as Insect Growth Regulators
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This entry is adapted from the peer-reviewed paper 10.3390/biom13030461

Glutathione S-transferases (GSTs) are conserved in a wide range of organisms, including insects. In 2014, an epsilon GST, known as Noppera-bo (Nobo), was shown to regulate the biosynthesis of ecdysteroid, the principal steroid hormone in insects. Studies on fruit flies, Drosophila melanogaster, and silkworms, Bombyx mori, demonstrated that loss-of-function mutants of nobo fail to synthesize ecdysteroid and die during development, consistent with the essential function of ecdysteroids in insect molting and metamorphosis. This genetic evidence suggests that chemical compounds that inhibit activity of Nobo could be insect growth regulators (IGRs) that kill insects by disrupting their molting and metamorphosis. In addition, because nobo is conserved only in Diptera and Lepidoptera, a Nobo inhibitor could be used to target IGRs in a narrow spectrum of insect taxa. Dipterans include mosquitoes, some of which are vectors of diseases such as malaria and dengue fever. Given that mosquito control is essential to reduce mosquito-borne diseases, new IGRs that specifically kill mosquito vectors are always in demand. 

glutathione S-transferases (GSTs) insecticide insect growth regulator (IGR) mosquito ecdysone

1. Introduction

Mosquitoes are responsible for the largest number of human deaths. They act as vectors for many pathogenic or parasitic infections such as malaria and dengue fever, which kill more than 700,000 people annually [1][2][3]. Therefore, it is essential to develop effective ways to control mosquito populations and prevent disease transmission. One method of mosquito control is to use insecticides that specifically act on cholinergic neurotransmission in insects and inhibit their motor functions. Organophosphates [4] and pyrethroids [5][6] are commonly used for mosquito control. However, some mosquitoes are developing resistance to them [7][8].

2. Insect Growth Regulators (IGRs)

Insect growth regulators (IGRs) are drugs that exhibit high insecticidal or growth-inhibitory activity against insect pests, but have no effect on other organisms. IGRs cause death by disrupting insect molting and metamorphosis [9]. Some of these target insect-specific structural materials, such as chitins, which are essential for formation of the cuticular layer, and insect hormones, which are required for molting and metamorphosis [10][11].
The Insect Resistance Action Committee (IRAC) classifies 34 groups of insecticides based on mode-of-action, identifying the following groups as IGRs.
(1) ”Juvenile hormone mimics” (Group 7) regulate insect molting and metamorphosis by agonizing the action of juvenile hormones (JHs) and indispensable sesquiterpenoid hormones. Commercially used mimics include Methoprene [12][13][14][15], Fenoxycarb [16], and Pyriproxyfen [17][18][19]. These insecticides are effective against a variety of insect species, such as those belonging to the orders Diptera, Lepidoptera, Coleoptera, Orthoptera, and Hymenoptera [20].
(2) “Mite growth inhibitors affecting CHS1” (Group 10) [21], “Inhibitors of chitin biosynthesis affecting CHS1” (Group 15) [14], and “Inhibitors of chitin biosynthesis, type1” (Group 16) [22] inhibit chitin synthase 1 (CHS1) in the epidermis and prevent insects from molting, which leads to death. The CHS1 gene, which encodes enzymes that produce chitin in the cuticle, is found in the exoskeleton, and these insecticides target agricultural pests, such as mosquitoes, and mites.
(3) “Inhibitors of acetyl CoA carboxylase” (Group 23) disrupt the first reaction of lipid biosynthesis by inhibiting the enzyme, acetyl CoA carboxylase, resulting in death. The representative IGR, Spirodiclofen [23][24], developed as an acaricide, effectively controls a broad spectrum of sucking pests such as mosquitoes and mites.
(4) “Molting disruptor, Dipteran” (Group 17) causes irregular melanization and sclerotization of the cuticle, resulting in necrotic lesions, rupture of the insect body, and death in dipteran larvae. In this class, Cyromazine [25] is a widely used IGR, for which the target molecule is unknown.
(5) “Ecdysone receptor agonists” (Group 18) bind to the Ecdysone receptor, the nuclear receptor for active ecdysteroids that are indispensable in arthropods, including insects, but do not have significant physiological effects on other organisms [26]. Dibenzoylhydrazines are well-known commercially available ecdysone receptor agonists [11][27][28][29]. Dibenzoyl hydrazines include Tebufenozide, Methoxyfenozide, and Chromafenozide, which exhibit selective insecticidal activity against lepidopterans [29].
The target molecule of IGRs is specific to insects, resulting in lower toxicity to non-target other animals, including humans, compared to other insecticides, thereby minimizing their impact on the environment [30]. IGRs classified in (2), (3), and (4) inhibit in vivo biosynthetic reactions through several mechanisms, while IGRs classified in (1) and (5) impact the binding of insect hormones, JHs and ecdysteroids by disrupting their receptors. However, no IGRs have yet been developed that target the biosynthesis of these hormones. Fortunately, in the past two decades, the biosynthetic pathways of JH and ecdysteroids and the enzymes involved have been identified and well characterized [31][32][33]. Therefore, IGR strategies that inhibit biosynthetic pathways rather than receptors are currently being considered [34].

3. Noppera-bo, the Ecdysteroidogenic GST

Ecdysteroids are arthropod steroid hormones, such as ecdysone and the more active 20-hydroxyecdysone, which regulate developmental and physiological processes in insects. Previous studies have shown that the amounts of active ecdysteroids, particularly 20E, in hemolymph fluctuate temporally during development [35]. Typically, surges of active ecdysteroid amounts are observed multiple times during development, which trigger temporal transitions in development, such as hatching, molting, metamorphosis, and eclosion.
During development, ecdysteroids are synthesized in the prothoracic gland (PG) from dietary cholesterol and plant sterol through multiple steps with various enzymes [32][36]. Ecdysteroid biosynthetic enzymes, including Nobo, collectively called Halloween enzymes [32], have been identified in Drosophila melanogaster and other insects over the past two decades [37][38][39]. Others include Neverland [40][41], non-molting glossy/Shroud [42], Spook/CYP307A1 [43][44], Spookier/CYP307A2 [44], Phantom/CYP306A1 [45][46], Disembodied/CYP302A1 [47], Shadow/CYP315A1 [47], and Shade/CYP314A1 [48]. Spook, spookier, phantom, disembodied, shadow, and shade encode cytochrome P450 monooxygenases, while others encode non-P450 type enzymes. All Halloween genes, except shadow, are specifically expressed in ecdysteroidgenic biosynthetic organs, including the PG and adult ovary, while shade is expressed in many peripheral tissues [32]. Genetic analyses of Halloween genes have been extensively conducted in D. melanogaster, and have revealed that the loss of Halloween genes causes embryonic or larval lethality due to defective ecdysteroid biosynthesis [49].
Among Halloween enzymes, Nobo is unique because it is the only ecdysteroidogenic enzyme belonging to the GST family, which are enzymes conserved in a wide range of organisms, from plants to animals [50][51][52]. GSTs generally catalyze conjugation of reduced glutathione (GSH) to substrates. Of the insect GSTs classified into six groups (delta, epsilon, sigma, theta, pi, and zeta) [53][54][55][56], Nobo belongs to the epsilon class. in vitro biochemical analysis has demonstrated that D. melanogaster Nobo (DmNobo; also known as GSTe14) exhibits steroid double-bond isomerase activity; however, corresponding steroid isomerization in vivo is unknown [57]. Although an endogenous substrate of Nobo has not yet been identified, genetic evidence strongly suggests that Nobo participates in cholesterol transport and/or metabolism [37][38][39]. Moreover, the requirement for GSH in ecdysteoid biosynthesis has also been confirmed; this is because a loss-of-function mutant of glutamate-cysteine ligase catalytic subunit (gclc), the critical enzyme for GSH biosynthesis, results in developmental lethality, partly due to a loss of ecdysteroid biosynthesis in D. melanogaster [58]. Thus, it is clear that GSH-dependent biochemical reactions mediated by Nobo are crucial for ecdysteroid biosynthesis in both dipteran and lepidopteran species. Of note, the developmental arrest phenotype observed in the loss-of-function mutant of nobo can be rescued by overexpression of nobo orthologs but not by the overexpression of other GST genes that do not belong to the Nobo family [38]. 

4. High-Throughput In Vitro Screening of Nobo Catalytic Activity

In general, several colorimetric and fluorometric methods have been used to detect GST activity in vitro. Conventional and typical substrates for colorimetric and fluorometric methods are 1-Chloro-2,4-dinitrobenzene (CDNB) [59][60] and monochlorobimane [61][62], respectively. However, both methods have low sensitivity and are unsuitable for high-throughput screening [34]. To overcome this low sensitivity, a novel fluorescent substrate, 3,4-DNADCF, was developed [63], which fluoresces approximately 54-fold upon GSH conjugation in the presence of GST. The enzymatic assay with 3,4-DNADCF was feasible even at a 1000-fold lower concentration than that of CDNB. Moreover, the high kcat/KM for 3,4-DNADCF is advantageous for the sensitive detection of GST enzymatic activity [63].
A high-throughput screening was conducted using 3,4-DNADCF with DmNobo recombinant proteins and a library of 9600 small-molecule compounds to identify inhibitors of DmNobo [64], and several inhibitors were identified [64][65][66]. Interestingly, these inhibitors included three steroid compounds, one of which was the mammalian steroid hormone 17 β-Estradiol (EST) [63]. EST is a strong inhibitor of DmNobo with 50% inhibitory concentration (IC50) of 1.2 ± 0.1 μM; however, it does not inhibit human GST P1-1 [63]. Nonetheless, EST is an endocrine-disrupting chemical, which renders it difficult to use as an insecticide [67]. Therefore, EST serves as a model compound to clarify the general mode of action of DmNobo inhibitors.

5. 17β-Estradiol Inhibits Drosophila melanogaster Nobo Enzymatic Activity

The Mannervik group solved the co-crystal structure of DmNobo with GSH and 2-methyl-2,4-pentanediol, originating from the crystallization mother liquor (Protein Data Bank (PDB):6T2T) [57]. Meanwhile, the Niwa group solved crystal structures of DmNobo: the apo form of DmNobo (PDB:6KEL), co-crystals with GSH (PDB:6KEN), EST (DmNobo-EST; PDB:6KEO), and both GSH and EST (PDB:6KEP) [65]. For any of these structures, like other GSTs generally, DmNobo forms a homodimer, conserving the GSH-binding pocket (G-site) and hydrophobic substrate-binding pocket (H-site). As expected, GSH is intercalated into the G-site, and EST and 2-methyl-2,4-pentanediol were bound to the H-sites.
Several amino acids in DmNobo interact with EST and GSH, via hydrophobic interaction with Phe39. However, the most important of these is the aspartate residue located at position 113 (Asp113), which is situated in the innermost region of the H-site. Specifically, the Oδ atom of Asp113 forms a hydrogen bond with the O3 atom in the hydroxyl group of EST [65]. The importance of the hydrogen bond between Asp113 and EST is also indicated by the fragment molecular orbital (FMO) method, which evaluates inter-fragment interaction energy [68][69][70]. Moreover, a point mutant DmNobo protein, in which Asp113 is replaced by alanine, has no loss of enzymatic activity even in the presence of 25 μM EST, providing bio-chemical evidence for the importance of hydrogen bonding between Asp113 and EST.

6. Flavonoidal Compounds, Such as Desmethylglycitein, Inhibit Aedes aegypti Nobo Enzymatic Activity

After identifying EST as a DmNobo inhibitor, it was initially speculated that EST would also inhibit the enzymatic activity of Nobo derived from the yellow fever mosquito, A. aegypti (AeNobo; also known as GSTe8). However, this is not the case. The IC50 of EST against AeNobo is 21.3 μM, which is approximately 10-fold higher than against DmNobo [64]. Therefore, to identify inhibitors of AeNobo, a high-throughput screening against AeNobo recombinant protein was performed using a library of 9600 small molecule compounds [64]. The compound, 2′-hydroxyflavanone, was identified with an IC50 of 4.76 μM. The inhibitory activities of other subclasses of flavonoids, including flavanone, flavone, isoflavone, flavanol, isoflavan, and anthocyanidin, were also tested against AeNobo. More than half of the tested flavonoid compounds exhibited inhibitory activity against AeNobo with IC50s less than 10 μM.
Three complex structures with three flavonoid inhibitors, daidzein (PDB:7EBU), luteolin (PDB:7EBV), and desmethylglycitein (DMG; IUPAC name:4′,6,7-trihy-droxyisoflavone; PDB:7EBW), were determined by X-ray crystallography. All three compounds formed hydrogen bonds with Glu113 in AeNobo. Moreover, as in the case of DmNobo and EST, the enzymatic activity of a point mutation of the AeNobo protein substituting Glu113 with alanine was not inhibited by any of these flavonoids, even at a concentration of 25 μM. These results suggest that AeNobo Glu113 is essential for the inhibitory activity of flavonoids, as with Asp113.
Of note, DMG-treated animals (2.5 ppm) exhibited growth delays and suppressed expression of the ecdysteroid-inducible gene, E74B, consistent with the fact that DMG inhibits AeNobo enzymatic activity. 

7. Glutathionylation of DmNobo Inhibitors

Inhibitors of DmNobo include not only EST, but also five non-steroidal compounds [66]: TDP011 (IUPAC name:4-bromo-2-[4-(3-methoxyphenyl)-2,2-dimethyl-5,6-dihydro-1H-pyrimidin-6-yl]phenol), TDP012 (IUPAC name:1-(4-fluorobenzyl)-1H-thieno [3,2-c][1,2]thiazin-4(3H)-one 2,2-dioxide), TDP013 (IUPAC name:2-(5-tert-butyl-2-methyl-benzenesulfonylamino)-benzoic acid), TDP015 (IUPAC name:6-(6,7-dihydrothieno [3,2-c]pyridin-5(4H)-yl)-3-pyridinamine), and TDP044 (IUPAC name:2,2′-(1,1-ethanediyl)bis(3-hydroxy-5,5-dimethyl-2-cyclohexen-1-one). Interactions between DmNobo and these five inhibitory compounds were investigated by X-ray crystallography, and three-dimensional structures of co-crystals of DmNobo with GSH and TDP011 (PDB:7BD3), TDP012 (PDB:7DB4), TDP013 (PDB:7DAY), and TDP015 (PDB:7DAZ) have been determined.
All four chemicals insert into the H-sites of DmNobo. Surprisingly, unlike EST, none of the four compounds form a hydrogen bond with Asp113. Moreover, none of the four compounds interacted with Phe39. Instead, the position of Phe39 changed between the apo-form and the complex forms with these non-steroidal compounds. When they are present in the H-sites, the aromatic ring of Phe39 moves away from the compounds, preventing their interaction with Phe39. These data imply that the inhibitory activity of these non-steroidal compounds is achieved by another mechanism.

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Subjects: Biochemistry & Molecular Biology
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Kana Ebihara
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Ryusuke Niwa
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Update Date: 09 May 2023
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