Journal Pre-proof Weaponisation ‘on the fly’: convergent recruitment of knottin and defensin peptide scaffolds into the venom of predatory assassin flies Jiayi Jin, Akello J. Agwa, Tibor G. Szanto, Agota Csóti, Gyorgy Panyi, Christina I. Schroeder, Andrew A. Walker, Glenn F. King PII: S0965-1748(19)30425-4 DOI: https://doi.org/10.1016/j.ibmb.2019.103310 Reference: IB 103310 To appear in: Insect Biochemistry and Molecular Biology Received Date: 8 October 2019 Revised Date: 12 December 2019 Accepted Date: 16 December 2019 Please cite this article as: Jin, J., Agwa, A.J., Szanto, T.G., Csóti, A., Panyi, G., Schroeder, C.I, Walker, A.A., King, G.F., Weaponisation ‘on the fly’: convergent recruitment of knottin and defensin peptide scaffolds into the venom of predatory assassin flies Insect Biochemistry and Molecular Biology, https:// doi.org/10.1016/j.ibmb.2019.103310. This is a PDF file of an article that has undergone enhancements after acceptance, such as the addition of a cover page and metadata, and formatting for readability, but it is not yet the definitive version of record. This version will undergo additional copyediting, typesetting and review before it is published in its final form, but we are providing this version to give early visibility of the article. Please note that, during the production process, errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain. © 2019 Published by Elsevier Ltd. Intended for submission to Insect Biochemistry and Molecular Biology Special Issue on Active Peptides in Insects Weaponisation ‘on the fly’: convergent recruitment of knottin and defensin peptide scaffolds into the venom of predatory assassin flies Jiayi Jin 1, Akello J. Agwa 1,# , Tibor G. Szanto 2, Agota Csóti 2, Gyorgy Panyi 2, Christina I. Schroeder 1, Andrew A. Walker 1,@ , and Glenn F. King 1,@ 1 Institute for Molecular Bioscience, The University of Queensland, St Lucia, QLD 4072, Australia. 2Department of Biophysics and Cell Biology, Faculty of Medicine, University of Debrecen, Debrecen H-4032, Hungary. @Please address correspondence to [email protected] (AAW) or [email protected] (GFK) #Current address: Department of Biology and Biochemistry, University of Bath, BA2 7AY, Bath, United Kingdom Keywords: Diptera, toxin, ion channel, K V11.1 (hERG), Dolopus genitalis 1 1 Abstract 2 Many arthropod venom peptides have potential as bioinsecticides, drug leads, and pharmacological 3 tools due to their specific neuromodulatory functions. Assassin flies (Asilidae) are a family of 4 predaceous dipterans that produce a unique and complex peptide-rich venom for killing insect prey 5 and deterring predators. However, very little is known about the structure and function of their 6 venom peptides. We therefore used an E. coli periplasmic expression system to express four 7 disulfide-rich peptides that we previously reported to exist in venom of the giant assassin fly 8 Dolopus genitalis . After purification, each recombinant peptide eluted from a C18 column at a 9 position closely matching its natural counterpart, strongly suggesting adoption of the native tertiary 10 fold. Injection of purified recombinant peptides into blowflies ( Lucilia cuprina ) and crickets 11 (Acheta domestica ) revealed that two of the four recombinant peptides, named rDg3b and rDg12, 12 inhibited escape behaviour in a manner that was rapid in onset (<1 min) and reversible. 13 Homonuclear NMR solution structures revealed that rDg3b and rDg12 adopt cystine-stabilised α/ß 14 defensin and inhibitor cystine knot folds, respectively. Although the closest known homologues of 15 rDg3b at the level of primary structure are dipteran antimicrobial peptides such as sapecin and 16 lucifensin, a DALI search showed that the tertiary structure of rDg3b most closely resembles the 17 KV11.1-specific α-potassium channel toxin CnErg1 from venom of the scorpion Centruroides 18 noxius . This is mainly due to the deletion of a large, unstructured loop between the first and second 19 cysteine residues present in Dg3b homologues from non-asiloid, but not existing in asiloid species. 20 Patch-clamp electrophysiology experiments revealed that rDg3b shifts the voltage-dependence of 21 KV11.1 channel activation to more depolarised potentials, but has no effect on KV1.3, K V2.1, 22 KV10.1, K Ca 1.1, or the Drosophila Shaker channel. Although rDg12 shares the inhibitor cystine 23 knot structure of many gating modifier toxins, rDg12 did not affect any of these KV channel 24 subtypes. Our results demonstrate that multiple disulfide-rich peptide scaffolds have been 25 convergently recruited into asilid and other animal venoms, and they provide insight into the 26 molecular evolution accompanying their weaponisation. 2 27 1. Introduction 28 Venoms are potent and highly adapted biochemical weapons that are used for diverse ecological 29 roles, including predation, parasitism, and the deterrence of predators and/or competitors (Casewell 30 et al., 2013; Fry et al., 2009). Understanding the biology of venom use allows one to relate 31 ecological interactions, animal behaviour and life history, and the evolution and pharmacology of 32 venom toxins (Barlow et al., 2009; Fry et al., 2015; Walker et al., 2017). Venoms are complex 33 cocktails of proteins, peptides, salts, and organic molecules. However, venom peptides in particular 34 are regarded as prime candidates for the discovery of environmentally-friendly insecticides (King, 35 2019; King and Hardy, 2013; Schwartz et al., 2012; Smith et al., 2013), medicines (Chassagnon et 36 al., 2017; Chi et al., 2012; King, 2015), and scientific tools (Dutertre and Lewis, 2010; Klint et al., 37 2012) because they typically display high levels of potency, selectivity, and chemically stability. 38 Cysteine-rich peptides are found in the venoms of diverse animal groups. Many are ion channel 39 modulators that underlie the ability of venom to induce paralysis or pain in prey and predators, 40 respectively (Bohlen and Julius, 2012; King et al., 2008; Lewis et al., 2012). While the primary 41 structures of individual members of a peptide family are often highly variable, their tertiary 42 structures are relatively conserved through shared secondary structural elements and disulfide-bond 43 connectivities. Comparative venom studies show an emerging pattern: although each venom is 44 unique and idiosyncratic, some ‘usual suspect’ peptide scaffolds occur in venoms of multiple 45 animal groups that have evolved venom use independently. The most likely explanation for this 46 phenomenon is that these peptide scaffolds are widespread, inherited from a common ancestor, and 47 adapted for non-venom physiological functions, but have intrinsically toxin-like pharmacokinetic 48 properties. Because of these properties, they have been convergently recruited as venom peptides 49 and further weaponised through evolution (Fry et al., 2009; Undheim et al., 2015a). These ‘usual 50 suspect’ scaffolds include cysteine-rich peptides such as inhibitor cystine knots (ICKs) (Pallaghy et 51 al., 1994; Undheim et al., 2016), cis- and trans- defensins (Shafee et al., 2016), Kazal domain 52 peptides (Walker et al., 2018b), and von Willebrand factor (vWF) domain peptides (Sunagar et al., 53 2013). Previous studies have sought to understand the molecular evolutionary events that underlie 54 the process of weaponisation, in which peptides adapted for other purposes become adapted as 55 venom toxins (Fry, 2005; Kini and Doley, 2010; Meng et al., 2016; Undheim et al., 2015a; Zhang et 56 al., 2018; Zhu et al., 2014). Further insights into weaponisation will undoubtedly be discovered not 57 only through peptide engineering and mutagenesis, but via structural and functional characterisation 58 of venom peptides made by additional taxa that evolved venom use independently. 59 Insects are the most speciose class of animals, with ~5.5 million extant species (Stork, 2018). Many 3 60 predaceous insect species that practice extra-oral digestion (EOD) are likely to possess venom 61 (Cohen, 1995), and venom use has evolved independently at least 13 times in the insects (Beard, 62 1963; Schmidt, 1982; Walker et al., 2018c; Zlotkin, 1984). Four of these instances occur in Diptera 63 (true flies), one of the largest and most diverse insect orders (Yeates et al., 2007). Diptera contains 64 many species that in the larval stage use venoms for capturing prey, deterring predators or both, 65 including the larval stages of snail-killing flies (Sciomyzidae), march and horse flies and allies 66 (Tabanomorpha), and the aphid midge Aphidoletes aphidimyza (Cecidomyiidae ) (von Reumont et 67 al., 2014; Walker et al., 2018c). These venoms have neurotoxic effects such as paralysis and 68 blocking of neuronal action potentials (Foote, 1961; Trelka and Berg, 1977), and they cause rapid 69 tissue liquefaction (Jackman et al., 1983; Nowicki and Eisner, 1983). The only dipteran group that 70 is known to use venom for prey capture in the adult stage are the assassin flies (also known as 71 robber flies; family Asilidae) (Kahan, 1964; Skevington and Dang, 2002; Walker et al., 2018c). 72 Assassin flies are a brachyceran group comprising > 7,000 species worldwide that originated at 73 least 112 million years ago during the Cretaceous Period (Dikow and Grimaldi, 2014). Adult 74 assassin flies have an enlarged syringe-like hypopharynx that is connected to two pairs of salivary 75 glands – named thoracic and labial glands due to the position of the secretory tissue – via a muscle 76 ring valve (Skevington and Dang, 2002; Whitfield, 1925). The paralysing function of asilid venom 77 was first reported almost a century ago (Melin, 1923). Early tests demonstrated that the toxins 78 delivered by assassin flies cause fast paralysis and death of insect prey (Whitfield, 1925). The 79 neurotoxic and lethal effects of assassin fly venom were further demonstrated in studies in which 80 thoracic gland extracts were injected into prey insects, causing paralysis and death (Kahan, 1964; 81 Musso et al., 1978).