Molecular targets and their application examples for interrupting chitin biosynthesis

Molecular targets and their application examples for interrupting chitin biosynthesis

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Yanwei Duan^a^,^b, Qing Yang^*^,^a^,^b

Chinese Chemical Letters | 2025, 36(4) : 109905

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Chinese Chemical Letters | 2025, 36(4): 109905

• Review •

Molecular targets and their application examples for interrupting chitin biosynthesis

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Yanwei Duan^a^,^b, Qing Yang^*^,^a^,^b

Affiliations

^aShenzhen Branch, Guangdong Laboratory of Lingnan Modern Agriculture, Genome Analysis Laboratory of the Ministry of Agriculture and Rural Affairs, Agricultural Genomics Institute at Shenzhen, Chinese Academy of Agricultural Sciences, Shenzhen 518120, China

^bState Key Laboratory for Biology of Plant Diseases and Insect Pests, Institute of Plant Protection, Chinese Academy of Agricultural Sciences, Beijing 100193, China

Published: 2025-04-15 doi: 10.1016/j.cclet.2024.109905

Outline

Abstract

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Chitin is an abundant aminopolysaccharide found in insect pests and phytopathogenic microorganisms but absent in higher plants and vertebrates. It is crucial for mitigating threats posed by chitin-containing organisms to human health, food safety, and agriculture. Therefore, targeting the chitin biosynthesis-associated bioprocess holds a promise for developing human-safe and eco-friendly antifungal agents or pesticides. Chitin biosynthesis requires chitin synthase and associated factors, which are involved in the modification, regulation, organization or turnover of chitin during its biosynthesis. A number of enzymes such as chitinases, hexosaminidases, chitin deacetylases are closely related and therefore are promising targets for designing novel agrochemicals that target at chitin biosynthesis. This review summarizes the advances in understanding chitin biology over the past decade by our research group and collaborates, specifically regarding essential proteins linked to chitin biosynthesis that can be exploited as promising pesticide targets. Examples of small bioactive molecules that against the activity of these targets are given.

Key words

Chitin biosynthesis / Chitin deacetylase / Chitinase / Hexosaminidase / Pesticide / Fungicide

Cite this Article

Yanwei Duan, Qing Yang. Molecular targets and their application examples for interrupting chitin biosynthesis[J]. Chinese Chemical Letters, 2025 , 36 (4) : 109905 - . DOI: 10.1016/j.cclet.2024.109905

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1 Introduction

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Chitin, the most abundant aminopolysaccharide in nature, is a linear polymer of β-(1,4)-linked N-acetylglucosamine (GlcNAc) and serves as a crucial component in the exoskeletons of insects and crustaceans, the egg shells of nematodes, the sporangia of oomycetes, and the cell walls of fungi and some protozoa [1-3]. Many chitin-containing organisms pose threats to human health, food safety, and agricultural production. However, chitin is generally absent in higher plants and vertebrates such as mammals, amphibians, birds, and reptiles [4]. Therefore, promising prospects lie in the development of human-safe and eco-friendly antifungal drugs and pesticides directed at chitin.

Chitin biosynthesis involves a key enzyme named chitin synthase, which repeatedly transfers a GlcNAc moiety from an activated sugar donor UDP-N-acetylglucosamine (UDP-N-GlcNAc) to the nonreducing 4‑hydroxyl group of a growing chitin chain [5]. In insects and fungi, several cellular enzymes have been identified to interact with chitin synthases, suggesting accessory factors are required for accomplishing chitin biosynthesis. Chitin preparations derived from natural sources, like crab and shrimp shells or insect exuviae, exhibits varying degrees of deacetylation due to enzymatic processes that convert chitin to chitosan by an enzyme named chitin deacetylase (CDA) [6,7]. During insect periodical molting, the degradation of old cuticle chitin is finely tuned and coupled to the synthesis of new cuticle chitin. Chitin biodegradation typically involves three enzymatic steps: The initial step is commonly catalyzed by lytic polysaccharide monooxygenases (LPMO), while the second step is usually catalyzed by two types of enzymes, namely endo-acting non-processive chitinases, and processive chitinases. Ultimately, β-N-acetyl-D-hexosaminidase (Hex) hydrolyzes oligosaccharides to GlcNAc [8]. Alongside these pivotal enzymes, various other factors, such as cuticle proteins, hormones, and interacting proteins of enzymes, play influential roles in chitin metabolism [9,10].

In the last decade, there has been significant exploration into multiple facets of chitin metabolism, greatly enhancing our understanding of chitin biology. This review focuses on the substantial contributions made by our team and collaborators with respect to essential proteins associated with chitin biosynthesis and relevance as well as small bioactive molecules developed to against these proteins. Some examples that might be used in controlling pest insects and pathogens are given.

2 Chitin synthases

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2.1 Molecular targets for chitin synthase and its auxiliary proteins

2.1.1 Chitin synthases

As early as 2011, Qu et al. identified chitin synthases OfChsA [11] and OfChsB [12] at the genetic level, responsible for chitin synthesis in the epidermis and midgut peritrophic membrane, respectively, of the agricultural pest Ostrinia furnacalis. OfChsA exhibits two alternative-splicing exons, namely exons 2a and 2b, as well as exons 19a and 19b [13]. Studying chitin synthase on a protein level presents considerable challenges due to its status as a membrane-integrated protein with multiple transmembrane domains. However, Chen et al. successfully addressed this issue by analyzing the cryo-electron microscopy structure of PsChs1 (structures of enzymes and their complexes with inhibitors are listed in the same sequence as presented in Table S1 (Supporting information)) [14], a chitin synthase found in Phytophthora sojae. The structural analysis revealed the reaction chamber responsible for governing chitin synthesis, as well as its competitive inhibition by the inhibitor nikkomycin-Z (Fig. 1), providing a structural basis for inhibition of chitin synthesis.

2.1.2 Chitin synthase-interacting proteins

Our studies indicate that chitin synthase requires auxiliary proteins, like chitin synthase-interacting proteins, to fulfill its pivotal role in growth and development. Specifically, the interacting proteins of chitin synthase (krotzkopf verkehrt, kkv) in Drosophila melanogaster were identified, including choline transporter-like protein 2 (Ctl2) [15], sarco/endoplasmic reticulum Ca²⁺-ATPase (Serca) [16] and fatty acid binding protein (Fabp) [17]. Knocking down them individually in the epidermis caused larval and pupal lethality, while in wings, it resulted in smaller, crinkled wings, a significant decrease in chitin deposition, and the loss of chitin lamellar structure.

2.2 Application examples for chitin synthesis

2.2.1 Inhibitors of chitin synthases

Before achieving a significant breakthrough in deciphering the structure of chitin synthase, we have to prompte the exploration of its inhibitors primarily through indirect approaches. Inspired by the diverse biological effects exhibited by many pyridazine derivatives containing oxadiazine heterocycles, as well as the intriguing bioactivity displayed by compounds incorporating hydrophobic and long alkyl chains, Ke et al. synthesized a series of novel 4H-1,3,4-oxadiazin-5(6H)-one derivatives [18]. Compounds 5i (1) (as shown in Fig. 2, inhibitors are numbered according to the order in which they appear in the article) and 5m (2) exhibited significant inhibitory activity on chitin biosynthesis. The oxadiazole heterocycles compound, DOW416, interferes with the formation of insect tissue and epicuticle. Thus, Ke et al. synthesized two series of 1,3,4-oxadiazoline heterocycle derivatives [19]. All synthesized compounds, particularly C1 (3) and C15 (4), potently inhibited chitin synthesis in yeast. Motivated by the aforementioned findings, Ke et al. subsequently developed a series of diacylhydrazine derivatives possessing hydrophobic alkyl chains [20]. Compounds 6b (5), 6g (6), 6j (7), and 6q (8) exhibited moderate to good inhibitory activity. Utilizing the structure of the chitin synthase donor substrate, UDP-GlcNAc, and the modeled structure of bacterial chitin synthase NodC, Chen et al. engineered a series of novel UDP-sugar analogues [21]. These newly synthesized molecules, exemplified by compounds 10 (9), 12 (10), and 13 (11), displayed high inhibitory activities against chitin synthase and demonstrated antifungal effects on plant-parasitic fungi such as Fusarium graminearum, Botrytis cinerea, and Cochliobolus lagenarium. In addition, Zhang et al. synthesized a novel series of 1,2,3,4-tetrahydroquinoline derivatives by incorporating active natural product groups into pesticide molecule structures [22]. Notably, compound 4fh (12) demonstrated slightly stronger inhibitory activity towards chitin synthase compared to polyoxin D.

2.2.2 Nanoscale RNAi carriers

The inherent instability of dsRNA and inadequate uptake through oral administration both pose significant hurdles for its practical implementation in insect pest management. Lu et al. developed core-shell nanoparticles to encapsulate vulnerable dsRNA payloads using block copolymers composed of poly(ethylene glycol)-polylysine(thiol). While oral delivery of RNAi is challenging in Locusta migratoria, our optimized RNAi construct effectively suppressed the expression levels of both LmChs1 and LmChs2, inducing discernible phenotypic changes [23].

3 Chitin modification, regulation, and organization

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3.1 Molecular targets in chitin modification, regulation, and organization

3.1.1 CDAs

Insect CDAs, belonging to the carbohydrate esterase 4 (CE4) family, are crucial for the modification, reorganization, and synthesis of chitin. Liu et al. were the first to report the structural and biochemical features of insect CDAs from Bombyx mori: BmCDA1, possibly functioning in cuticle modification, and BmCDA8, which may modify peritrophic membranes in the midgut [24]. Besides BmCDA8, Liu et al. also characterized two other anterior midgut-expressed CDAs (BmCDA6 and BmCDA7) from Bombyx mori [25]. Of the three enzymes, BmCDA7 and BmCDA8 were observed only at the feeding stage, while BmCDA6, inactive towards PM chitin, was expressed almost exclusively at the mid-molt stage.

Phytopathogenic fungi secrete CDA as a means to evade the host's immunological defense mechanisms during infection. Specifically, Liu et al. demonstrated that the deacetylation activity of CDA is crucial for fungal pathogenicity. They determined the crystal structures of two representative phytopathogenic fungal CDAs: VdPDA1 from Verticillium dahliae and Pst_13661 from Puccinia striiformis f. sp. tritici [26].

3.1.2 Hormones

The nuclear receptor-mediated 20-hydroxyecdysone (20E) signaling pathway plays a vital role in insects. Zhao et al. identified a cDNA encoding a Locusta migratoria hormone receptor 39 (LmHR39) based on transcriptomics data [27]. The transcription of LmHR39 could be induced by 20E in vivo. RNAi experiments implied that the LmHR39 is involved in the regulation of chitinase genes. On another note, the elaborate regulation of tissue- and stage-specific expression of chitin synthase is a prerequisite for insect development. Zhang et al. report a regulatory mechanism for the pupa-specific expression of ChsA-2b in Bombyx mori [28].

3.1.3 Cuticle proteins

Insect cuticle is comprised of a protein matrix and chitin fibers, which exhibit varying degrees of deacetylation. Therefore, specialized proteins are presumed necessary to bind deacetylated chitin chains together. Qu et al. were the first to report a chitin-binding protein, BmCPAP3-D1, exhibited the binding activity toward deacetylated chitin [29]. Qi et al. found cycloxaprid's bioactivity against lepidopteran pests to be comparatively inferior. They revealed that cuticle proteins appear to be crucial for the insensitivity to cycloxaprid by comparing the transcriptomes of cycloxaprid-treated and untreated Ostrinia furnacalis larvae. In line with this conjecture, pretreatment of larvae with methoprene resulted in a 1.12-fold enhancement in cycloxaprid's bioactivity [30]. To comprehend how molecular interactions between cuticle proteins and chitin govern cuticle assembly, Gong et al. discovered a newly identified and highly abundant cuticular protein, hypothetical-1 from Ostrinia furnacalis (OfCPH-1), capable of forming coacervates in the presence of chitosan [31].

3.2 Application examples for chitin modification

The crystal structures of VdPDA1 and Pst_13661 revealed that Zn²⁺, which is necessary for activity, was coordinated by the Asp-His-His-triad (Fig. 3). Thus, metal ion chelating agents could potentially serve as inhibitors of CDAs. In practice, Liu et al. discovered that benzohydroxamic acid (BHA) (13) and its three derivatives showed good inhibitory activities toward VdPDA1 and Pst_13661 [26]. BHA demonstrated high activity against diseases in cotton, wheat, and soybean caused by Verticillium dahliae, as well as four fungal plant pathogens, namely Puccinia striiformis f. sp. tritici, Fusarium oxysporum, Fusarium graminearum, and Rhizoctonia solani.

4 Chitin degradation

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4.1 Molecular targets in chitin degradation

In order to perform a comprehensive analysis of the chitinolytic enzymes engaged in chitin degradation, Qu et al. conducted proteomic assessments on molting fluids of Bombyx mori, revealing the prevalence of only four abundant ones in molting fluids. These include two members of the insect glycoside hydrolase family 18 (GH18) endochitinases (group I chitinases and group II chitinases, ChtI and ChtII), a GH18 exochitinase (group h chitinases, Chi-h), which are exclusively found in lepidopterans, and an insect glycoside hydrolase family 20 (GH20) group 1 β-N-acetyl-D-hexosaminidase (Hex1). The specific gene expression patterns in tissues and stages provide compelling evidence for the involvement of these four enzymes in cuticular chitin degradation [32]. In exploring factors influencing enzyme processivity within the chitinolytic enzyme system, Qu et al. scrutinized an insect chitinase cocktail, comprising OfChi-h, OfChtI, and OfChtII, using high-speed atomic force microscopy [33]. Their findings underscore the significant role of the endochitinase in enhancing the processivity of the exochitinase.

4.1.1 GH20 β-N-acetyl-D-hexosaminidases

Prior to the systematic analysis of chitinolytic enzymes, Yang et al. isolated and characterized a Hex1 (OfHex1) from Ostrinia furnacalis, indicating its involvement in insect chitin catabolism [34]. Subsequently, Liu et al. obtained the recombinant OfHex1 [35] and acquired the crystal structure of OfHex1, along with its complex with the substrate analogue N,N,N-trimethyl-D-glucosamine (TMG)-(GlcNAc)₃ [36]. The crystal structures of wild-type OfHex1 and the mutant OfHex1(V327G) in complex with the inhibitor PUGNAc highlighted structural variations that influence diverse sensitivities to PUGNAc [37]. Through molecular docking of OfHex1 with inhibitor allosamidin, showcasing the distinct size and shape of OfHex1’s pocket compared to human Hex (HsHex), indicated allosamidin's potential to selectively inhibit OfHex1 [38]. The comprehensive structural and mutational analyses revealed the essential role of Glu328 in conjunction with Trp490 for substrate binding [39,40]. Apart from OfHex1, insects harbor multiple genes encoding other Hexs that serve various physiological functions. Liu et al. identified two Hexs, OfHex2 and OfHex3, cloned from Ostrinia furnacalis [41]. OfHex2 is presumed to play similar roles as HsHexB in insect [42]. OfHex3 is involved in both molting and fertilization processes [43]. The final one, named FDL (OfFDL), contributes significantly to N-glycan modification [44].

4.1.2 GH18 chitinases

Among the three abundant chitinases in molting fluids, ChtI (OfChtI) from Ostrinia furnacali were firstly expressed, and characterized by Wu et al. [45]. Further studies reported the unliganded and oligosaccharide-complexed crystal structures of OfChtI [46]. Successively, Liu et al. investigated crystal structures of Chi-h from Ostrinia furnacalis (OfChi-h) and its complex with chitoheptaose [47]. Chen et al. presented the crystal structures of catalytically active domains of OfChtII [48]. OfChtII was expressed earlier than OfChtI and OfChi-h during the molting transition and was responsible for completing the pretreatment of crystal chitin substrates [49]. Liu et al. investigated crystal structures of the group III chitinase (OfChtIII) from Ostrinia furnacalis [50]. The gene-expression pattern and subcellular localization of OfChtIII mirrored that of OfChsA, suggesting involvement in the chitin-synthesis pathway. The group IV chitinase of Ostrinia furnacalis (OfChtIV) is specifically expressed in the midgut. Liu et al. showed that OfChtIV exhibited high stability and mycelial hydrolytic activity in the extreme midgut environment through hyper-N-glycosylation [51]. Furthermore, Chen et al. presented the first crystal structure of a representative nematode chitinas CeCht1 from Caenorhabditis elegans [52]. These chitinase crystal structures mentioned above provide a solid basis for the development of inhibitors.

4.1.3 LPMOs

LPMOs are abundantly present in various insect species and belong to the auxiliary activities family 15 (AA15) LPMOs (LPMO15). Qu et al. selected Tribolium castaneum and Locusta migratoria to investigate the functions of insect LPMO15 subgroup I-like proteins (LPMO15–1s) [53]. In both species, LPMO15–1-deficient animals were unable to undergo exuviation and exhibited mortality due to impaired turnover of the chitinous cuticle. Qu et al. also investigated the function of midgut-specific LmLPMO15–3 during development in Locusta migratoria [54]. Knockdown of LmLPMO15–3 at instar nymph stage resulted in lethal phenotypes, characterized by incomplete digestion of the peritrophic matrix.

4.2 Application examples for chitin degradation

4.2.1 Inhibitors of OfHex1

The expression and crystallization of OfHex1 fifteen years ago marked a significant milestone in the study of chitinolytic enzymes. Since then, substantial progress has been made in the study of its inhibitors. Screening and optimizing known inhibitors based on structural insights represent a promising strategy for inhibitor development. TMG-(GlcNAc)₃ (14) (Fig. 4) is a natural inhibitor of insect Hexs. Our structural analysis indicated that the two GlcNAc residues at the reducing end might be unnecessary. Therefore, Yang et al. designed a significantly simplified skeleton molecule, TMG-(GlcNAc)₂ (15), which exhibits activity equivalent to TMG-(GlcNAc)₃ [55]. The symmetrical bis-naphthalimide M-31850 (16) was previously obtained by screening for specificity against human Hex (HsHex). Liu et al. designed an unsymmetrical dyad of naphthalimide and thiadiazole, Q2 (17), which shifted naphthalimide specificity from HsHex to insect and bacterial Hexs [56]. NAG-thiazoline (NGT) (18) is a well-established inhibitor against most beta-GlcNAcases. Using co-crystallization, Liu et al. designed a compound NMAGT (19) with a bulky substituent on the thiazoline ring of NGT, exhibiting a K_i value of 0.13 µmol/L, which is 600-fold lower than the K_i value observed for NGT [57]. Through screening a library of microbial secondary metabolites, phlegmacin B₁ (20) was identified as an inhibitor of OfHex1 with K_i values of 26 µmol/L. Injection and feeding experiments demonstrated that phlegmacin B₁ has an insecticidal effect on Ostrinia furnacalis larvae [58]. Inspired by the work above, Duan et al. noted that compounds with a large conjugated plane were highly potent inhibitors of GH20 Hex. They first revealed that berberine (21), a typical compound with a large conjugated plane, acted as competitive inhibitors of OfHex1 [59].

The synthesis of novel inhibitors through leveraging highly active groups obtained previously presents an efficient approach for inhibitor development. In a study by Yang et al., a series of thiazolylhydrazone derivatives were designed, amalgamating a thiazoline group from NGT and a naphthalimide group from Q2. Among these derivatives, 3k (22) exhibited increased potency with a K_i of 10.2 µmol/L. Subsequent optimization led to the discovery of an even more potent inhibitor, derivative 7 (23), boasting a K_i value of 2.1 µmol/L [60]. Using a similar approach, Yang et al. synthesized a range of Q2 derivatives. Among these derivatives, compound 3m (24) displayed the most promising inhibitory activity, while compound 6a (25) demonstrated superior activity among the quinoline-containing derivatives [61]. They also simultaneously designed and synthesized two additional series of thiazolylhydrazones. Derivatives I-3d (26) and II-3d (27) exhibited remarkable inhibitory activities against OfHex1 [62]. In efforts to enhance the inhibitory efficiency of naphthalimide derivatives, Shen et al. undertook rational molecular design and optimization, culminating in the synthesis of compounds 15r (28) and 15y (29). These compounds showcased superior activity and selectivity toward OfHex1 compared to lead compounds [63]. Furthermore, the triazole moiety demonstrates optimal structural attributes for forming hydrogen bonds and π-π stacking interactions with biological targets. Dong et al. designed a series of novel glycosyl triazoles. Among these, compound 17c (30) displayed suitable activity and selectivity against OfHex1 [64]. C-Glycosides are frequently employed as analogs of native O-glycosides. Liang et al. synthesized a series of novel C-glycosidic oximino carbamate derivatives. Compound 7k (31) demonstrated the most potent inhibitory activity against OfHex1, in addition to exhibiting remarkable larvicidal activity against Plutella xylostella [65].

The utilization of structure-based virtual screening for identifying inhibitors stands as an efficient strategy in inhibitor development. Building upon the crystal structure of OfHex1, a virtual screening of approximately 200,000 small molecules from the SPECS database was conducted. Twenty-eight compounds were chosen for subsequent bioactivity evaluation, and compound 3 (32) exhibited potent inhibitory activity against OfHex1 [66]. In a parallel study, Dong et al. carried out sequential virtual screenings of the ZINC library, housing 8 million compounds. They ultimately selecteed 15 molecules for further enzymatic assays. Among these, compound 5 (33) displayed promising inhibitory activity against OfHex1, demonstrating significant selectivity [67]. More recently, a comprehensive approach utilized the SPECS and Maybridge databases, alongside our in-house library of 2393 molecules, for structure-based virtual screening. After simplification of structures, compounds featuring a biphenyl–sulfonamide skeleton emerged as potential OfHex1 inhibitors. Compounds 10k (34), 10u (35), and 10v (36) exhibited K_i values comparable to the most active nonglycosyl-based inhibitor, Q2 [68]. What distinguishes these newly discovered compound skeletons are their simple chemical structures, ease of synthesis, and potent activity.

4.2.2 Inhibitors of other Hexs

In addition to OfHex1, we have utilized the aforementioned approaches to investigate inhibitors targeting Hex across various subgroups, species, or GH families. Guo et al. synthesized a novel and efficient skeleton comprising naphthalimide and methoxyphenyl moieties. The most potent inhibitor, compound 7a (37), exhibits high inhibitory activity with K_i values of 0.63 µmol/L against HsHex [69]. Similarly, a series of naphthalimide-scaffold were designed based on molecular modeling analysis of OfHex2. The most potent inhibitor, compound 20 (38), displayed a K_i value of 0.37 µmol/L [70]. The GH84 β-N-acetyl-D-hexosaminidase (OGA) is responsible for removing GlcNAc from serine or threonine residues of glycoproteins. Kong et al. generated a total of 24 new NGT derivatives across two separate batches. Among the compounds screened, compounds 5a (39) (The half maximal inhibitory concentration (IC₅₀) = 12.6 µmol/L, hOGA) and 5e (40) (IC₅₀ = 12.5 µmol/L, OfOGA) [71], as well as compounds 7d (41) (IC₅₀ = 6.4 µmol/L, hOGA) and 7f (42) (IC₅₀ = 11.9 µmol/L, hOGA) [72], demonstrated their high potency as inhibitors. Moreover, Chen et al., using the complex structure of hOGA-PUGNAc as a basis, synthesized a series of novel thioglycosyl–naphthalimide hybrid inhibitors. Notably, compounds 5c (43) (hOGA, K_i = 3.46 µmol/L; OfHex1, K_i > 200 µmol/L) and 6f (44) (hOGA, K_i > 200 µmol/L; OfHex1, K_i = 21.81 µmol/L) exhibited high selectivity [73]. Additionally, different linkers were introduced into the naphthalimide-bearing thioglycoside derivatives to enhance interactions. The most potent compounds were 15j (45) (K_i = 0.91 µmol/L, HsHexB; K_i>100 µmol/L, hOGA) and 15b (46) (K_i = 3.76 µmol/L, hOGA; K_i = 30.42 µmol/L, HsHexB), which displayed significant selectivity between these two enzymes [74].

4.2.3 Inhibitors of OfChtI

Based on the catalytic mechanism, Chen et al. discovered that the inhibitory effects of fully deacetylated chitooligosaccharides (GlcN)_2–7 (47) (Fig. 5) against OfChtI are dependent on the degree of polymerization. The injection of mixed (GlcN)_2–7 into fifth instar larvae of Ostrinia furnacalis resulted in 85% larval arrest and subsequent death within 10 days [75]. Utilizing virtual screening from the ZINC database, Jiang et al. developed two chemical series of chitinase inhibitors, FQ (48), displaying specific inhibition against OfChtI, and TP (49), exhibiting broad inhibitory activity against chitinases derived from insects, humans, fungi, and bacteria [76]. Revolving around the identical concept, Dong et al. found a chemical fragment and five variant compounds as inhibitors of OfChtI from SPECS chemical database. Compound 3 (50) showed preferential inhibitory activity with a K_i value of 1.5 µmol/L against OfChtI [77]. It possessed a large modification space at 6-position of compound 3 based on the predicted binding mode. Its analogue compound 8 (51) with 6‑tert-pentyl showed preferential inhibitory activity with a K_i value of 0.71 µmol/L [78]. Expanding on the active lead compound 6i, Jiang et al. synthesized a series of novel heptacyclic pyrazolamide derivatives. Notably, compound D-27 (52) exhibited promising activities against Plutella xylostella and Mythimna separata [79]. Additionally, compound D-08 (53) induced aberrant molting in Plutella xylostell, while its derivative III-27 (54) exhibited superior activity against Plutella xylostella [80]. Han et al. initially discovered that piperine, a natural product, exhibited partial inhibition of OfChtI. Subsequently, they synthesized compounds 5a–f by incorporating a butenolide scaffold into piperine. Compounds 5e (55) and 5f (56) demonstrated moderate insecticidal activity against Ostrinia furnacalis and displayed approximately 80-fold higher inhibitory activity against OfChtI compared to piperine [81]. Recently, Han et al. designed a series of piperonyl-rhodanine derivatives. The optimized compound IIIAe (57) exhibited significant inhibitory activity against OfChtI (K_i = 2.4 µmol/L) and demonstrated potent insecticidal efficacy with a mortality rate of 63.33% [82].

4.2.4 Inhibitors of OfChtI

The exploration of small molecules as inhibitors of ChtII had been limited until our report on two constructs of OfChtII. Subsequently, Chen et al. conducted a screening of the compound library in our laboratory and identified four potent inhibitors: (GlcN)₈ (47), dipyrido-pyrimidine derivative (DP) (58), piperidine-thienopyridine derivative (PT) (59), and naphthalimide derivative (NI) (60). Injection of the inhibitors into 4^th instar Ostrinia furnacalis larvae resulted in defects in development and pupation [83].

4.2.5 Inhibitors of OfChi-h

When Chen et al. first investigated OfChi-h and obtained its crystal structure, they simultaneously discovered that OfChi-h was inhibited by TMG-(GlcNAc)₄ (61), an inhibitor of OfHex1 [47]. Similarlly, Chen et al. identified phlegmacin B₁ [58] and two thiazolylhydrazones derivatives I-3d and II-3d [62], which demonstrated inhibitory activities against OfHex1, as also effective inhibitors of OfChi-h. Through hierarchical virtual screening of the ZINC database, Jiang et al. isolated dipyrido-pyrimidine derivatives compound 53 (62) as an insect-selective OfChi-h inhibitor with a K_i value of 9 nmol/L, and compound 40 (63) as a novel, human Chit1-selective inhibitor with a K_i value of 49 nmol/L [84]. Subsequently, Dong et al. designed a series of azo-aminopyrimidine derivatives. Among these derivatives, compound 8f (64) stood out as the most potent with a K_i value of 64.7 nmol/L. Notably, this compound exhibited superior insecticidal activity when compared to the control pesticide hexaflumuron [85].

With a meticulous analysis of existing inhibitor-chitinase interactions, Yuan et al. proposed that planar polycyclic compounds might establish hydrophobic stacking interactions with aromatic residues, mimicking the GlcNAc unit's stacking interactions to inhibit chitinase activity. They constructed a series of dipyridopyrimidine-3-carboxamide derivatives. Among these derivatives, compound 6t (65) exhibited the most potent activity against OfChi-h, with K_i values of 5.6 nmol/L. The strong stacking interaction of compound 6p (66) with conserved residues found in the co-crystal structure validated the feasibility of their design [86]. Following a similar principle, Zhu et al. identified berberine as a moderate inhibitor of OfChi-h with a K_i of 16.1 µmol/L. Their strategy to enhance its efficacy involved introducing aromatic and heterocyclic rings to the intact berberine framework. As a result, compound 19e (67) emerged as a potent OfChi-h inhibitor with a K_i of 0.093 µmol/L. Treatment with structurally stable compound 19c (68) hindered the growth and metamorphosis of Ostrinia furnacalis larvae [87]. Lu et al. discovered lynamicin B (69), a natural product, as a selective inhibitor of OfChi-h with a K_i value of 8.76 µmol/L. Lynamicin B demonstrated substantial insecticidal activity against various lepidopteran pests while exerting minimal effects on the ahymenopteran natural enemies of these pests [88]. Another natural product, argifin, exhibited potent inhibitory activity against chitinase. To address the challenge of its limited synthetic accessibility, Zhao et al. devised a strategy involving the replacement of the cyclic peptide backbone skeleton found in 12- and 16-azamacrolides. Compounds 19c (70) emerged as the most potent, displaying IC₅₀ values of 56 nmol/L against OfChi-h and inducing a 76% mortality rate for Plutella xylostella at a concentration of 50 mg/L [89].

4.2.6 Inhibitors of multiple chitinases

In recent years, the multitarget approach has gained traction in drug development. Qi et al. discovered in their initial screening of a natural products library that kasugamycin (71) inhibits OfChtI with a K_i value of 0.47 µmol/L and OfChi-h with a K_i value of 2.7 µmol/L [90]. On the basis of the same multitarget strategy, Li et al. conducted a high-throughput screen of the TargetMol library (1680 natural products) for inhibitors targeting OfChtI, OfChtII, OfChi-h, and OfHex1. Shikonin (72) and wogonin (73) emerged as potent inhibitors against all four enzymes, displaying significant insecticidal effects on lepidopteran agricultural pests. Additionally, 10-hydroxycamptothecin (10-HCPT) exhibited inhibitory activity against all four enzymes [91]. Later, camptothecin and its derivatives, such as SN-38 (74), were identified as competitive inhibitors of OfChtII and OfChi-h with micromolar K_i values. Camptothecin also showed substantial insecticidal activity against Locusta migratoria [92]. Taking advantage of above data, Ding et al. devised a strategy that integrates machine learning to sift through a large natural product library from Topscience (17,600 compounds) in search of novel multitarget inhibitors. They identified 3,5-di-O-caffeoylquinic acid (75) and γ-mangostin (76) as inhibitors for all four enzymes, with K_i values at the µmol/L level. These compounds exhibited notable biological activities against lepidopteran pests [93]. Jiang et al. designed 19 novel piperine derivatives based on the interaction modes between piperine and OfChtI, OfChtII, and OfChi-h. Among these derivatives, compound 5k (77) (K_i = 11.78–22.82 µmol/L) was identified as the most potent multichitinase inhibitor and demonstrated superior insecticidal activity against Ostrinia furnacalis compared to dual- or single-chitinase inhibitors [94]. Han et al. synthesized a series of piperonyl-tethered rhodanine derivatives. In comparison to piperine, compounds 7a–f (78) and 7g–j (79) exhibited approximately 100- to 400-fold or 110- to 210-fold higher inhibitory capacity against OfChtI and OfChi-h, respectively. Furthermore, compounds 7a–c demonstrated significant inhibition of the growth and development of Ostrinia furnacalis larvae [95]. Building upon a12 (80), Jin et al. rationally designed a series of benzo[d][1,3]dioxole-6-benzamide derivatives. Among them, compound d29 (81) acted simultaneously on OfChtI, OfChtII, and OfChi-h with K_i values of 0.8, 11.9, and 2.3 µmol/L, and exhibited significant activity against two lepidopteran pests [96]. To increase the ability of the target compounds to penetrate the insect cuticle in vivo by increasing their lipophilicity, Zhao et al. synthesized a series of potent insecticides targeting OfChtI and OfChi-h using lipophilic piperidine as a bridge connecting the planar structures and the N-methylcarbamoylguanidino. Compounds 6e (82), 6g (83), 6j (84), and 6o (85) significantly affected the growth and development of Plutella xylostell [97].

4.2.7 Inhibitors of other chitinases

In addition to the previously mentioned chitinolytic enzymes derived from insect molting fluids, Liu et al. employed these approaches to investigate inhibitors targeting chitinases across different subgroups and species. Following our initial investigation of OfChtIV, Liu et al. concurrently discovered that OfChtIV exhibited inhibition by allosamidin (86) with a K_i value of 1.35 µmol/L [51]. Chen et al. synthesized a variety of berberine analogues using a hydrophobic cavity-based optimization strategy. The compound 4c (87) showed an 80-fold elevated inhibitory activity against SmChiB and the human hAMCas [98]. Through hierarchical virtual screening from the ZINC database, two series of novel inhibitors against CeCht1 were successfully identified. These inhibitors featured a BP (88) scaffold [52] and a PP (89) scaffold [99]. Meanwhile, compounds HAU-4 (90) and HAU-7 (91) were also identified as CeCht1 inhibitors with IC₅₀ values of 4.2 µmol/L and 10.0 µmol/L, respectively [100]. Recently, Jin et al. conducted a virtual screening of a library comprising over 16,000 natural products and successfully identified lunidonine as an inhibitor of CeCht1. Subsequently, Jin et al. employed a pocket-based lead optimization strategy, and new compound a12 (80) exhibited a strong interaction with CeCht1 and demonstrated remarkable in vitro nematicidal activity against Caenorhabditis elegans [101].

5 Conclusion and future work

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Researchers have long recognized the potential of chitin biosynthesis as a target for insecticides and have maintained a dedicated focus in this area [102-104]. Correspondingly, investigations into chitin biosynthesis primarily center around insects and fungi [6,105-107]. A comprehensive book, published in 2019, delved into various aspects of chitin biology, with a specific focus on chitin remodeling enzymes and inhibitors [108]. As supplements and updates, our research advances in chitin biosynthesis have highlighted chitin-biosynthesis relevant targets for developing highly selective pesticides and anti-pathogen agents [110,111]. A variety of lead compounds with high inhibitory abilities have been reported, paving a path for a new era of agrochemicals to interrupt chitin biosynthesis [7,109-114]. The future research priorities may encompass: (1) Further in-depth biological investigations into co-factors that assist the cellular regulation and transportation of chitin synthases; (2) structural investigation into the mechanism of the co-factors; and (3) chemical modification and optimization of lead compounds to improve their biological activities.

Declaration of competing interest

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The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

CRediT authorship contribution statement

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Yanwei Duan: Writing – original draft, Methodology. Qing Yang: Writing – review & editing, Supervision, Project administration, Funding acquisition, Conceptualization.

Acknowledgments

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This work was supported by the National Key Research and Development Program of China (No. 2022YFD1700200), the National Natural Science Foundation of China (Nos. 32161133010, 3230170969), the Innovation Program of Chinese Academy of Agricultural Sciences, the Shenzhen Science and Technology Program (No. KQTD20180411143628272), and the Special Funds for Science Technology Innovation and Industrial Development of Shenzhen Dapeng New District (No. PT202101–02).

Supplementary materials

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Supplementary material associated with this article can be found, in the online version, at doi:10.1016/j.cclet.2024.109905.

References

Less

W. Cheng, M. Lin, M. Qiu, et al., Environ. Microbiol. 21 (2019) 4537–4547.

B. Moussian, Chitin: structure, chemistry and biology, in: Q. Yang, T. Fukamizo (Eds. ), Targeting Chitin-containing Organisms, Springer, Singapore, 2019, pp. 5–18.

A. Schwelm, J. Fogelqvist, A. Knaust, et al., Sci. Rep. 5 (2015) 11153.

E. Cohen, Chitin biochemistry: synthesis, hydrolysis and inhibition, in: J. Casas, S. Simpson (Eds. ), Advances in Insect Physiology, Academic Press, San Diego, 2010, pp. 5–74.

H. Merzendorfer, J. Comp. Physiol. B 176 (2006) 1–15.

K. Zhu, H. Merzendorfer, W. Zhang, J. Zhang, S. Muthukrishnan, Annu. Rev. Entomol. 61 (2016) 177–196.

Y. Li, L. Liu, J. Yang, Q. Yang, J. Pestic. Sci. 46 (2021) 43–52.

G. Courtade, F.L. Aachmann, Chitin-active lytic polysaccharide monooxygenases, in: Q. Yang, T. Fukamizo (Eds. ), Targeting Chitin-containing Organisms, Springer, Singapore, 2019, pp. 115–129.

B. Moussian, Insect Biochem. Mol. Biol. 40 (2010) 363–375.

H. Merzendorfer, Eur. J. Cell. Biol. 90 (2011) 759–769.

M. Qu, Q. Yang, Insect Biochem. Mol. Biol. 41 (2011) 923–931.

M. Qu, T. Liu, J. Yang, Q. Yang, Biochem. Biophys. Res. Commun. 404 (2011) 302–307.

M. Qu, Q. Yang, Insect. Mol. Biol. 21 (2012) 395–404.

W. Chen, P. Cao, Y. Liu, et al., Nature 610 (2022) 402–408.

Y. Duan, W. Zhu, X. Zhao, et al., Insect Biochem. Mol. Biol. 141 (2022) 103718.

W. Zhu, Y. Duan, J. Chen, et al., Insect Biochem. Mol. Biol. 145 (2022) 103783.

J. Chen, X. Zou, W. Zhu, et al., Insect Biochem. Mol. Biol. 149 (2022) 103845.

S. Ke, X. Qian, F. Liu, et al., Eur. J. Med. Chem. 44 (2009) 2113–2121.

S. Ke, F. Liu, N. Wang, Q. Yang, X. Qian, Bioorg. Med. Chem. Lett. 19 (2009) 332–335.

S. Ke, X. Qian, F. Liu, et al., Eur. J. Med. Chem. 44 (2009) 2985–2993.

Q. Chen, J. Zhang, L. Chen, et al., Chin. Chem. Lett. 28 (2017) 1232–1237.

X. Zhang, Z. Yang, H. Xu, et al., J. Agric. Food Chem. 70 (2022) 9262–9275.

Q. Lu, H. Cui, W. Li, et al., J. Agric. Food Chem. 70 (2022) 10762–10770.

L. Liu, Y. Zhou, M. Qu, et al., J. Biol. Chem. 294 (2019) 5774–5783.

L. Liu, M. Qu, T. Liu, et al., J. Insect. Physiol. 113 (2019) 42–48.

L. Liu, Y. Xia, Y. Li, et al., Nat. Commun. 14 (2023) 3857.

X. Zhao, Z. Qin, J. Zhang, et al., Insect Mol. Biol. 28 (2019) 537–549.

J. Zhang, G. Xu, B. Qiu, et al., Insect Biochem. Mol. Biol. 116 (2020) 103264.

M. Qu, Y. Ren, Y. Liu, Q. Yang, Insect Mol. Biol. 26 (2017) 432–439.

H. Qi, T. Liu, Q. Lu, Q. Yang, J. Agric. Food Chem. 68 (2020) 982–988.

Q. Gong, L. Chen, J. Wang, et al., Biomacromolecules 23 (2022) 2562–2571.

M. Qu, L. Ma, P. Chen, Q. Yang, J. Proteome Res. 13 (2014) 2931–2940.

M. Qu, T. Watanabe-Nakayama, S. Sun, et al., ACS Catal. 10 (2020) 13606–13615.

Q. Yang, T. Liu, F. Liu, M. Qu, X. Qian, FEBS J 275 (2008) 5690–5702.

T. Liu, F. Liu, Q. Yang, J. Yang, Protein Expr. Purif. 68 (2009) 99–103.

T. Liu, H. Zhang, F. Liu, et al., J. Biol. Chem. 286 (2011) 4049–4058.

T. Liu, H. Zhang, F. Liu, et al., Biochem. J. 438 (2011) 467–474.

Y. Wang, T. Liu, Q. Yang, Z. Li, X. Qian, Chem. Biol. Drug Des. 79 (2012) 572–582.

T. Liu, Y. Zhou, L. Chen, et al., PLoS One 7 (2012) e52225.

T. Liu, Q. Wu, L. Liu, Q. Yang, Process. Biochem. 48 (2013) 103–108.

T. Liu, M. Qu, Q. Yang, J. Yang, X. Qian, J. Biotechnol. 136 (2008) S109.

F. Liu, T. Liu, M. Qu, Q. Yang, Int. J. Biol. Sci. 8 (2012) 1085–1096.

M. Qu, T. Liu, P. Chen, Q. Yang, PLoS One 8 (2013) e71738.

Y. Huo, L. Chen, M. Qu, Q. Chen, Q. Yang, Arch. Insect Biochem. Physiol. 83 (2013) 115–126.

Q. Wu, T. Liu, Q. Yang, Insect Sci. 20 (2013) 147–157.

L. Chen, T. Liu, Y. Zhou, et al., Acta Crystallogr. D: Biol. Crystallogr. 70 (2014) 932–942.

T. Liu, L. Chen, Y. Zhou, et al., J. Biol. Chem. 292 (2017) 2080–2088.

W. Chen, M. Qu, Y. Zhou, Q. Yang, J. Biol. Chem. 293 (2018) 2652–2660.

M. Qu, S. Sun, Y. Liu, et al., Insect Sci. 28 (2021) 692–704.

T. Liu, W. Zhu, J. Wang, et al., Acta Crystallogr. D: Struct. Biol. 74 (2018) 30–40.

T. Liu, X. Guo, Y. Bu, et al., Insect Biochem. Mol. Biol. 119 (2020) 103326.

Q. Chen, W. Chen, A. Kumar, et al., J. Agric. Food Chem. 69 (2021) 3519–3526.

M. Qu, X. Guo, S. Tian, et al., Commun. Biol. 5 (2022) 518.

M. Qu, X. Guo, L. Kong, L. Hou, Q. Yang, Insect Sci. 29 (2022) 1287–1298.

Y. Yang, T. Liu, Y. Yang, et al., ChemBioChem 12 (2011) 457–467.

T. Liu, P. Guo, Y. Zhou, et al., Sci. Rep. 4 (2014) 6188.

T. Liu, M. Xia, H. Zhang, et al., FEBS Lett. 589 (2015) 110–116.

L. Chen, T. Liu, Y. Duan, X. Lu, Q. Yang, J. Agric. Food Chem. 65 (2017) 3851–3857.

Y. Duan, T. Liu, Y. Zhou, T. Dou, Q. Yang, J. Biol. Chem. 293 (2018) 15429–15438.

H. Yang, T. Liu, H. Qi, et al., Bioorg. Med. Chem. 26 (2018) 5420–5426.

H. Yang, H. Qi, T. Liu, et al., Chin. Chem. Lett. 30 (2019) 977–980.

H. Yang, H. Qi, Z. Hao, et al., Chin. Chem. Lett. 31 (2020) 1271–1275.

S. Shen, L. Dong, W. Chen, et al., J. Agric. Food Chem. 67 (2019) 6387–6396.

L. Dong, S. Shen, W. Chen, et al., Bioorg. Med. Chem. 27 (2019) 2315–2322.

P. Liang, Q. Xu, R. Chen, et al., Carbohydr. Res. 520 (2022) 108629.

Y. Dong, S. Hu, X. Zhao, et al., Pest Manag. Sci. 76 (2020) 3030–3037.

L. Dong, S. Shen, Y. Xu, et al., J. Biomol. Struct. Dyn. 39 (2021) 1735–1743.

T. Chen, W. Li, Z. Liu, et al., J. Agric. Food Chem. 69 (2021) 12039–12047.

P. Guo, Q. Chen, T. Liu, et al., ACS Med. Chem. Lett. 4 (2013) 527–531.

Q. Chen, P. Guo, L. Xu, et al., Biochimie 97 (2014) 152–162.

H. Kong, W. Chen, H. Lu, et al., Carbohydr. Res. 413 (2015) 135–144.

H. Kong, W. Chen, T. Liu, et al., Carbohydr. Res. 429 (2016) 54–61.

W. Chen, S. Shen, L. Dong, J. Zhang, Q. Yang, Bioorg. Med. Chem. 26 (2018) 394–400.

S. Shen, W. Chen, L. Dong, et al., J. Enzyme Inhib. Med. Chem. 33 (2018) 445–452.

L. Chen, Y. Zhou, M. Qu, Y. Zhao, Q. Yang, J. Biol. Chem. 289 (2014) 17932–17940.

X. Jiang, A. Kumar, T. Liu, K. Zhang, Q. Yang, J. Chem. Inf. Model. 56 (2016) 2413–2420.

Y. Dong, X. Jiang, T. Liu, et al., J. Agric. Food Chem. 66 (2018) 3351–3357.

Y. Dong, S. Hu, X. Jiang, et al., J. Agric. Food Chem. 67 (2019) 3575–3582.

B. Jiang, X. Jin, Y. Dong, et al., J. Agric. Food Chem. 68 (2020) 6347–6354.

B. Jiang, B. Guo, J. Cui, et al., Bioorg. Med. Chem. Lett. 30 (2020) 127500.

Q. Han, N. Wu, H. Li, et al., J. Agric. Food Chem. 69 (2021) 7534–7544.

Q. Han, N. Wu, J. Zhang, et al., J. Agric. Food Chem. 71 (2023) 18685–18695.

W. Chen, Y. Zhou, Q. Yang, J. Biol. Chem. 294 (2019) 9358–9364.

X. Jiang, A. Kumar, Y. Motomura, et al., J. Med. Chem. 63 (2020) 987–1001.

L. Dong, S. Shen, X. Jiang, et al., J. Agric. Food Chem. 70 (2022) 12203–12210.

P. Yuan, X. Jiang, S. Wang, et al., J. Agric. Food Chem. 68 (2020) 13584–13593.

L. Zhu, L. Chen, X. Shao, et al., J. Agric. Food Chem. 69 (2021) 7526–7533.

Q. Lu, L. Xu, L. Liu, et al., J. Agric. Food Chem. 69 (2021) 14086–14091.

Z. Zhao, Q. Xu, W. Chen, et al., J. Agric. Food Chem. 70 (2022) 4889–4898.

H. Qi, X. Jiang, Y. Ding, T. Liu, Q. Yang, Front Mol. Biosci. 8 (2021) 640356.

W. Li, Y. Ding, H. Qi, T. Liu, Q. Yang, J. Agric. Food Chem. 69 (2021) 10830–10837.

Y. Ding, S. Chen, F. Zhang, et al., J. Agric. Food Chem. 71 (2023) 1845–1851.

Y. Ding, S. Chen, H. Liu, T. Liu, Q. Yang, J. Agric. Food Chem. 71 (2023) 8769–8777.

Z. Jiang, D. Shi, H. Li, et al., J. Agric. Food Chem. 70 (2022) 10326–10336.

Q. Han, N. Wu, Y. Liu, et al., J. Agric. Food Chem. 70 (2022) 7387–7399.

X. Jin, T. Sun, B. Guo, et al., J. Agric. Food Chem. 71 (2023) 8345–8355.

Z. Zhao, W. Chen, Y. Dong, et al., J. Agric. Food Chem. 71 (2023) 12431–12439.

L. Chen, L. Zhu, J. Chen, et al., J. Enzyme Inhib. Med. Chem. 35 (2020) 1937–1943.

W. Chen, Q. Chen, A. Kumar, et al., J. Enzyme Inhib. Med. Chem. 36 (2021) 1198–1204.

100

S. Shen, B. Ding, X. Jiang, et al., Front Chem. 10 (2022) 1021295.

101

X. Jin, T. Sun, X. Zhang, et al., J. Agric. Food Chem. 71 (2023) 244–254.

102

E. Cohen, Pest Manag. Sci. 57 (2001) 946–950.

103

F. Matsumura, Pestic. Biochem. Phys. 97 (2010) 133–139.

104

H. Merzendorfer, Insect Sci. 20 (2013) 121–138.

105

H. Merzendorfer, L. Zimoch, J. Exp. Biol. 206 (2003) 4393–4412.

106

S. Muthukrishnan, H. Merzendorfer, Y. Arakane, K.J. Kramer, Chitin metabolism in insects, in: L. Gilbert (Ed. ), Insect Molecular Biology and Biochemistry, Academic Press, San Diego, 2012, pp. 193–235.

107

S. Muthukrishnan, H. Merzendorfer, Y. Arakane, Q. Yang, Chitin metabolic pathways in insects and their regulation, in: E. Cohen, B. Moussian (Eds. ), Extracellular Composite Matrices in Arthropods, Springer, Cham, 2016, pp. 31–65.

108

Q. Yang, T. Fukamizo, Targeting Chitin-containing Organisms, 1st. ed, Springer, Singapore, 2019.

109

W. Chen, X. Jiang, Q. Yang, Biotechnol. Adv. 43 (2020) 107553.

110

A. Yu, M. Beck, H. Merzendorfer, Q. Yang, Insect Biochem. Mol. Biol. 164 (2023) 104058.

111

T. Liu, Y. Duan, Q. Yang, Biotechnol. Adv. 36 (2018) 1127–1138.

112

W. Chen, Q. Yang, J. Agric. Food Chem. 68 (2020) 4559–4565.

113

Y. Ding, Q. Lu, T. Liu, Q. Yang, Adv. Agrochem. 2 (2023) 306–312.

114

Q. Lu, H. Xie, M. Qu, T. Liu, Q. Yang, J. Agric. Food Chem. 71 (2023) 5944–5952.

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Year 2025 volume 36 Issue 4

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doi: 10.1016/j.cclet.2024.109905

Receive Date：2024-01-20
Online Date：2025-11-19
Published：2025-04-15

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Received：2024-01-20
Revised：2024-04-12
Accepted：2024-04-16

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^bState Key Laboratory for Biology of Plant Diseases and Insect Pests, Institute of Plant Protection, Chinese Academy of Agricultural Sciences, Beijing 100193, China

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表12种不同金属材料的力学参数

科 Family	属数 Number of genus	种数 Number of species	占总种数比例 Percentage of total species (%)	属 Genus	种数 Number of species	占总种数比例 Percentage of total species (%)
鹅膏菌科Amanitaceae	2	11	5.26	鹅膏菌属 Amanita	10	4.78
小菇科 Mycenaceae	2	12	5.74	丝盖伞属 Inocybe	5	2.39
多孔菌科 Polyporaceae	8	14	6.70	蜡蘑属 Laccaria	5	2.39
红菇科 Russulaceae	3	23	11.00	小皮伞属 Marasmius	6	2.87
				小菇属 Mycena	11	5.26
				光柄菇属 Pluteus	5	2.39
				红菇属 Russula	17	8.13
				栓菌属 Trametes	5	2.39

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Fig. 1 Inhibition of PsChs1 by nikkomycin-Z. Sliced-surface view (left) of the nikkomycin-Z-binding site and detailed interactions between nikkomycin-Z and PsChs1 (right).

Fig. 2 Chemical structures of chitin synthases inhibitors and CDAs inhibitor. Inhibitors were numbered according to the order in which they appeared in the article.

Fig. 3 Inhibition of Pst_13661 by BHA. Surface view of the BHA bound in Pst_13661 (left) and interactions between BHA and Pst_13661 (right).

Fig. 4 Chemical structures of Hex inhibitors. Inhibitors were numbered according to the order in which they appeared in the article.

Fig. 5 Chemical structures of chitinase inhibitors. Inhibitors were numbered according to the order in which they appeared in the article.

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