Imperial Journal of Interdisciplinary Research (IJIR) Vol-2, Issue-11, 2016 ISSN: 2454-1362, http://www.onlinejournal.in

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Plant-Insect Interactions-Cyanogenic Glucosides

Showket Ahmad Dar1; Amir Bashir Wani2; M. Younus Wani3; Shabber Hussain4 & M. Saleem Majid5

Department of 1Entomology 2 Biotechnology, 3Sericulture; 4Fruit Science; 5Geology Sher-e-Kashmir University of Agricultural Science and Technology, Shalimar, Jammu and

Kashmir-India-190025. University of Kashmir, Hazratbal, Srinager; Department of Geology (KU)

SKUAST-K pin code: 190006

Abstract: An essential driving component in the co-evolution of plants and insects is the ability to produce and handle bioactive compounds. Plants produce bioactive natural products for the defense, but some insects detoxify and/or sequester the compounds, opening up for new niches with fewer competitors. Among the important bioactive component the cyanogenic glucosides are known to be present in more than 2500 plant species and are considered to have an important role in plant defense against herbivores due to bitter taste and release of toxic hydrogen cyanide upon tissue disruption. Some specialized herbivores, especially insects, preferentially feed on cyanogenic plants. Such herbivores have acquired the ability to metabolize cyanogenic glucosides or to sequester them for use in their predator defense. A few species of Arthropoda (Diplopoda, Chilopoda, Insecta) are able to de novo synthesize cyanogenic glucosides and, in addition, some of these species are able to sequester cyanogenic glucosides from their host plant. Moths belonging to the Zygaena family are the only insects known, able to carry out both de novo biosynthesis and sequestration of the same cyanogenic glucosides as those from their feed plants. Evolutionary aspects of these unique plant-insect interactions with focus on the enzyme systems involved in synthesis and degradation of cyanogenic glucosides are discussed. Key words: Insect, Bioactive, Plants, Interactions, Cyanogenic compounds.

Introduction

There are a number of broad categories of toxicologically significant plant constituents. These include alkaloids, amino acids, peptides and proteins, oxalic acids, terpenes, phenolics, tannins, glycosides and essential oils (steam-volatile, primarily lipophilic, organic plant metabolites stored in special plant organs and perceived by man through the stimulation of the sense of smell or taste). Plants are known to produce more than 300,000 different secondary metabolites

including the group of cyanogenic glycosides (CNGs), which are phytoanticipins widely distributed in the plant kingdom (Bolarinwa et al., 2014; Zagrobelny et al., 2013), containing the accompanying intact toxic metabolite β-glucosidase (Abraham et al., 2015). More than 2600 plant including species of ferns, gymnosperms and angiosperms (Zagrobelny et al., 2004) produce myriad of the CNGs, one of the biggest and most studied class of plant secondary metabolites (Vetter, 2000). CNGs have been found in a few arthropod clades and are defensive against most of them (Shlichta et al., 2014). Further, in course of evolution, CNGs in plants have acquired additional roles to improve plant plasticity, i.e, establishment, robustness, and viability in response to the environmental challenges (Gleadow and Moller, 2014). In response to these chemical defense compounds the herbivorous insects have evolved strategies to circumvent the plant chemical defense systems, e.g., by detoxification etc. (Zagrobelny, 2014). Since, cyanogenic glycosides are derivatives of various amino acids. But specifically they are β-glucosides of α-hydroxynitriles (Shlichta et al., 2014) derived from the aliphatic protein amino acids ʟ-valine, ʟ-isoleucine and ʟ-leucine from aromatic amino acids like ʟ-phenylalanine and ʟ-tyrosine and from aliphatic non-protein amino acid cyclopentenyl-glycine (Irmisch et al., 2013). In plants, CNGs are stored in the vacuoles and offers an immediate chemical defense response to insects (Moller, 2010). When plant tissue like lema bean is disrupted e.g. by insect attack, CNGs are brought into contact with β-glucosidases and α-hydroxyntrile lyases that hydrolyze the CNGs and thereby, cause release of toxic hydrogen cyanide (HCN) (Moller, 2010; Shlicha et al., 2014; Blom et al., 2011). Hydrogen cyanide specifically inhibits cytochrome C oxide, a key enzyme in the mitochondrial respiratory pathways (Blom et al., 2011), which causes cell and tissue death within a short time, and strongly deters various non-adapted

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herbivores as fall armyworm larvae quickly die when fed on an artificial diet containing cyanide.

In insects, CNGs act as feeding and oviposition deterrent and are toxic to enzymatic hydrolysis, for instance mites Tetranychus species (Wybouw et al., 2014). The toxicity is mainly due to its affinity for the terminal cytochrome oxidase in the mitochondrial respiratory pathway (Brattstem et al., 1983). The toxic amino acid β-cyano-L-analine is found to be present in 21 species of the family Zygaenidae (Witthohn and Naumann, 2014).The degradation of the glucosidases yielded the reactive hydroxynitriles, which releases toxic HCN to kill the insects, especially the lepidopterans. Yet many mite and few lepidopteran species can thrive well on the plants defended by cyanogenic glucosides, but the nature of the enzyme known to detoxify different ingested lethal doses of HCN to β-cyanoalanine has remained enigmatic (Wybouw et al., 2014). Since, CNGs are involved in plant defense against herbivores (Ballhorn and Elais, 2014; Bernards et al., 2011) due to release of toxic HCN (Ganjewala et al., 2010; Nahrstedt, 1996; Shlicha et al., 2014). The β-glucosidase is generally very stable enzymes due to their compactly folded structure, and has pH optima in slightly acidic condition in plants as well as insects (Pankoke et al., 2012; Terra and Ferreira, 2012). In most plants and insects β- glucosidase belongs to the glycoside hydrolase (GH) family 1. GHs comprise more than 100 described glycosyl hydrolase families, and these families are classified based on the structure and amino acid sequence similarities. GHs β-glucosidase catalyzes the hydrolysis of a glycosidic bond between two carbohydrates or between a carbohydrates moiety and an aryl or alkyl aglycone moiety (Cantarel et al., 2009). The glucosylated plant defences are activated by plant β- glucosidases to release toxic aglucons upon plant damage e.g, alkaloid, benzoxazinoid, cyanogenic and iridoid alkaloid, benzoxanoid and iridoid glucosidase as well as glucosinolates and salicinoids (Pentzold et al., 2013). The lethal dose of cyanide for vertebrates lies in the range of 35-150 µmol kg-1, if applied in a single dose. Much higher amounts of HCN can be tolerated if consumed or administered over a longer period (Davis and Nahrstedt, 1985). CNGs are however, also known to act as both feeding/oviposition deterrents and phagostimulants for herbivores that are specialists on plants containing CNGs. Therefore, this review is about to explain the current knoweldege on the CNGs and their biosynthesis, hydrolysis and their sequestration and detoxification in arthropods. Degradation and detoxification of cyanogenic glucosides

The CNGs are subjected to degradation and detoxification in plants (Spampinato, 2011). The enzyme cytochrome P450 catalyzes the biosynthesis of CGNs, and the first P450 catalyzed step proceeds via two successive stages including N-hydroxylations of the amino group of the parent amino acid, followed by its decarboxylation and dehydration (Sibbesen et al., 1994; Montellano and Nelson, 2011). During this process the aldoxime formed is subsequently converted to α-hydroxime through the action of a second cytochrome P450 (Irmisch et al., 2013). In western balsam popular (Populous trichocarpa) the two P450 enzymes involved in herbivour-induced aldoxime formation are mostly viz. CYP79D6v3 and CYP79D7v2. The aldoxime are well known precursor of several classes of direct defense compounds and are produced from their corresponding amino acids through the action of multifunctional cytochrome P450 monooxygenase (CYP) of the CYP79 family (Hamberger and Bak, 2013). The first described CYP79 enzyme, CYP79A1, is isolated and characterized from sorghum (Sorghum bicolor), catalyzes the conversion of Try to P-hydroxylphenylaldoxime, the rate limiting step in cyanogenic glucoside biosynthesis. Cytochrome P450 dependent monooxygenenases and DNA molecules encode the monooxygenases are able to catalyze the biosynthetic conversion of aldoximes to nitriles and the later conversion of the same nitriles to the corresponding cyanohydrins, which are the precursors of CNGs (Hamberger and Bak, 2013). The reaction involves an initial dehydration. The final step in CNG synthesis, glycosylation of the cyanohydrin moiety, is catalyzed by a UDPG-glycosyltransferase (Jones et al., 1999). The metabolism involves the two successful N-hydroxylation, dehydration and a final isomerization reaction. Catabolism of CNGs is initiated by enzymatic hydrolysis by a β-glucosidase to afford the corresponding α-hydroxynitrile, which at pH values above 6 spontaneously dissociates into a sugar, a keto compound, and HCN. However, at lower pH values, the dissociation reaction is catalyzed by an α-hydroxynitrile lyase. HCN is detoxified by two main reactions (Bordo and Bork, 2002; Miller and Conn, 1980), involved the formation of β-cyanoalanine from cysteine and is catalyzed by β-cyanoalanine-synthase. The rout which is common in insects and plants, detoxification route involved the β-cyanoalanine pathway; whileas, the thiocyanate pathway occurs mainly in vertebrates but also in some plants and insects. The aldoxime intermediates were not found to accumulate in the plant, but are channeled through a large protein complex called a metabolon (Moller, 2010). This

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mechanism prevents the release of the putative toxic and reactive intermediates (Grootwassink et al., 1990; Moller, 2010). Cytochrome P450 enzyme are heme-thiolate mono-oxygenases that catalyze metabolism of the various endogenous and exogenous compounds, constituting the superfamily of enzymes present in various organisms including the mammals, plants, bacteria and insects. P450 enzymes are diverse and metabolize a wide variety of substrates, but their structures are largely conserved. The irreversible nature of reactions catalyzed by P450s makes these enzymes landmark in the evolution of plant metabolic pathways like degradation and detoxification of harmful substrates (Hamberger and Bak, 2013). Cytochromes P450 are a large group of polyphyletic proteins and is spatially regulated at transcript level showing differential tissue specificity (Rana et al., 2014) with amino acid sequence as low as 20% within species. Unlike microbes, the cytochromes P450 in all plants and animals are type II membrane enzymes, with N-terminally anchored to the endoplasmatic membrane. In prokaryotes, the water soluble cytochromes P450 requires the elicitors for the constant expression (Rana et al., 2014); whileas, in eukaryotes, P450 is membrane bound and in general functions to insert one molecule of oxygen into its substrate, with its heme-prosthetic group playing a role in substrate oxidation. This catalytic reaction requires a pair of electron shuttled from NADPH via the NADPH-cytochrome P450 reductase (CYPOR) enzyme, a P450 redox partners, to target P450 (Ortiz de and De Voss, 2005). In contrast, in bacteria the mitochondria, ferredoxin reductase and iron-Sulphur ferredoxin proteins act as a bridge (Kim et al., 2013) to transfer reducing equivalent from NADP(H) to target P450s are membrane bound enzymes that play key roles in degradation, detoxification and xenobiotic metabolism. Whileas, in plants the proteins viz, glutaredoxin, BOLA and NEET as well as MIP18, MMS19, TAH18, DRE2 for the cytosolic machinery are integrated into a model for the plant Fe-S cluster biogenesis and detoxification systems (Couturier et al., 2013). It becomes evident that P450s are implicated in pyrethroid resistance in insects. In insects and plants, cytochromes P450 are encoded by some of the largest multigene families, with 89 genes in the Drosophila melanogaster (Endopterygota) genome, 100 genes in the Anopheles gambiae (Endopterygota) genome and 272 genes in the Arabidopsis thaliana genome (Werck-Reichhart et al., 2000). Cytochromes P450 catalyze a highly diverse range of chemical reactions (Guengerich and Munro 2013; Heel et al., 2014; Shoji and Watanabe 2014; Mclntosh et al.,

2014) that include C-hydorxylations and epoxidations. In insects, the cytochromes P450 is important for the biosynthesis of various insect metamorphic substances like, ecdysteroids and juvenile hormones (Daimon et al., 2012; Noriega, 2014) as well as degradation of insecticides and their residues. Cytochromes P450 play crucial roles in defense against natural products that insects have to fend off in order to be able to feed on otherwise toxic plants. The ability of an insect cytochrome P450 to degrade a specific natural plant based toxic substance (Mclntosh et al., 2014) is often the key to the adaptation of insect herbivores to their host selection and landing. In plants, cytochromes P450 play main role in the synthesis of a defensive natural products (Mclntosh et al., 2014) involved in plant defense as well as many other biosynthetic pathways (Morant et al., 2003).

The enzyme family 1 glycosyltransferase catalyzed the final step in biosynthesis of CNGs (Gantt et al., 2013; Shlichta et al., 2014; Spampinato, 2011). These enzymes are soluble proteins with a molecular mass of 45-60 kDa, and utilize UDP-activated sugar moieties as the donor molecules to glycosylate the acceptor molecules. Glycosyltransferases generally exhibit a low degree of overall sequence similarity, and are often regioselective or regiospecific rather than highly substrate specific. Further, the families 1 glycosyltransferases are encoded by a multigene family and are ubiquitously found in plants, animals, fungi, bacteria and viruses (Paquette et al., 2003). Lotus japonicas, like several other legumes, biosynthesizes the cyanogenic α-hydroxynitrile glucosides lotaustralin and linamarin. Upon tissue disruption these compounds are hydrolyzed by a specific β-glucosidase, resulting in the release of hydrogen cyanide. Lotus japonicas also produce the non-cyanogenic γ- and β-hydroxynitrile glucosides rhodiocyanoside A and D (Lai et al., 2014) using a biosynthetic pathway that branches off from lotaustralin biosynthesis.

In the biosynthesis of the plant hormone, ethylene cyanides are produced and therefore need to be detoxified. The cyanogenic plants are able to utilize this detoxification pathway when cyanogenic glucosides are degraded into cyanides. In plants, degradation CNGs is catalyzed by β-glucosidases and α-hydroxynitrile lyases (Barmina 2010; Ganjewala et al., 2010). The β-Glucosidases catalyze the hydrolysis of glycosidic linkage in aryl and alkyl B-glucoside in bacteria, fungi, plants and animals; whileas, the well characterized β-glucosidases involved in CNG catabolism is present exclusively in plants. However, only little is known about the insect β-glucosidases and their

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substrate specificity. β-Glucosidases generally have a subunit molecular mass of 55-65 kDa, acidic pH optima (pH 5-6) and an absolute specificity towards β-glucosidases involved in cleavage of CNGs, exhibit a high specificity towards the aglycone moiety of CNGs present in the same plant species. Cyanogenic glucosides, the precursor of cyanide in many plants, arthropods and some bacteria are amino acid derivative β-glycosides of α-hydroxynitriles. Since the enzyme α-Hydroxynitrile lyases have been characterized in plants and insect. In plants, they appear to degrade β-glucosidases though their activity in protein bodies, is well known (Swain et al., 1992), instead of in chloroplasts or apoplastic space as typically reported for β-glucosidases (Hickel et al., 1996). The α-Hydroxynitrile lyases constitute two broad phylogenetically distinct groups. One homogeneous group comprises monomeric FAD-containing glycosylated enzymes. These enzymes have only been found in two subfamilies within the rosaceae and utilize the aromatic cyanohydrins mandelonitrile as a substrate. The second group comprises dimeric or oligomeric non-FAD containing enzymes that typically are not glycosylated, found in di-and monocotyledonous plant families, and their natural substrates may be p-hydroxymandelonitrile as well as acetone cyanohydrins, the latter being the most common substrate. This group of non-FAD containing α-hydroxynitrile lyases may be divided into subgroups depending on highest sequence similarity to serine carboxypeptidases and alcohol dehydrogenases. In plants, β-cyanoalanine synthase has pyridoxal phosphate as a cofactor and is a member of an ancestral family of β-substituted alanine synthases that also includes cysteine synthase (Ikegami and Murakoshi, 1994). The β-cyanoalanine is a potent neurotoxin and its accumulation in plants may serve to deter predators. The activity of the β-Cyanoalanine synthase plays a pivotal role in detoxification of HCN released as a result of cleavage of cyanogenic glucosides, or formed in stoichiometric amounts by the action of an enzyme 1-aminocyclo-propane-1-carboxylic acid oxidase (Yip and Yang, 1988). The β-cyanoalanine synthase activity in plants and insects is primarily located in mitochondria, the organelle that is most vulnerable to HCN toxification. In contrast to B-cyanoalanine synthase, rhodanese is not ubiquitously present in plants. In those species of higher animals, plants and insects in which rhodanese is present, it is thought to play a role in cyanide detoxification (Muinat et al., 2014). Rhodanese is a mitochondrial enzyme that detoxifies cyanide (CN-) by converting it to thiocyanate (SCN-). In plants, this assignment

is supported by high levels of rhodanese activity in Sorghum bicolor seedlings. In these seedlings, the cyanide potential is exceptionally high and also serve a variety of other functions, the most important of which is to donate sulfur to proteins (Bordo and Bork, 2002).

Cyanogenic glucosides and plant-herbivore interactions

In pollen feeding Heliconius (Papilionoidea), eggs are produced mainly from adult acquired resources, leaving somatic development and maintenance to the larval efforts. This has spurred the evolution of the chemical defenses via amino acid derived cyanogenic glycosides (Cardoso and Gilbert 2013). Plant cyanogenesis release gaseous hydrogen cyanide (HCN) in response to the cell damage and is considered as an effective defense against generalist herbivores, however the specialists are generally believed not to be affected negatively by this trait and the quantative data on long-term effects of cyanogenesis on specialists are rear. E.g, lima bean accessions (Fabaceae: Phaseolus linatus L.) possess high quantative variability of cyanogenic potential (HCNp: concentration of the cyanogenic precursors) and cyanogenic capacities (HCNc release of gaseous HCN per unit time). The herbivour Mexican bean beetle (coleoptera: Coccinellidae: Epilachna varivestis Muslnat) feed on the selected lines characterized by high (HC-plants) and low HCNp (LC-pants) (Ballhorn et al., 2007). Larval and adult stages of this herbivore feed on a narrow range of legumes and prefer cyanogenic lima bean as a host plant. Nevertheless, the performance of beetles was significantly affected by lima bean HCNp: body weight decreased and developmental period of larvae and pupae increased on HC-plants during the first generation of beetles and then remained constant for four consecutive generations and low HCNp (LC-plants) (Ballhorn et al 2011). Larval and adult stages of this herbivore feed on a narrow range of legumes and prefer cyanogenic lima bean as host plant. Effects can only be detected when considering more than one generation. Thus, cyanide-containing precursors can have negative effects even on herbivores adapted to feed on cyanogenic plants.

Herbivores react very differently to the presence of CNGs in their diet. Nearly all of the variability in the effectiveness of CNGs in plant defense against herbivory is explained by four confounding factors (Gleadow and Woodrow, 2002), like the concentration of CNGs in a host plant may be below threshold toxicity, the herbivore may be a specialist that has evolved mechanisms to cope

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with high levels of HCN in the diet, the cyanogenic plant may be consumed as part of a mixed diet and the toxicity of CNGs in this way diluted to below threshold value and the mode of herbivore feeding may be adapted to minimize tissue damage to leaves (e.g. aphids, which are phloem feeders) to limit exposure of CNGs to degradative β-glucosidases. It appears that the prime deterrent effect of CNGs is linked to the keto compound released in equimolar amounts to HCN during CNG degradation, rather than to the CNG or HCN (Jones, 1988). The biosynthetic pathway for CNGs is highly channeled, preventing intermediates to dissociate from the enzyme complex (Moller and Conn, 1980). Accordingly, the biosynthetic intermediates are not likely to act as deterrents to herbivores. In contrast, degradation of CNGs may result in accumulation of cyanohydrins, keto compounds or aldehydes, HCN, β-cyanoalanine, thiocyanate and sulfite. Each of these compounds may be envisioned to possess defensive properties: CNGs have a bitter taste and have been shown to act as feeding deterrents; aldehydes and ketones possess cytotoxic activities; HCN is a powerful inhibitor of respiration and of enzymes that contain heavy metals; β-cyanoalanine is a neurotoxin (Nahrstedt, 1985) and thiocyanate and sulfite are enzyme inhibitors. Z. filipendulae belongs to the only insect family known to both de novo biosynthesize and sequester the same compounds directly from its food-pant. Z.filipendulae and L.Corniculatus both contain the two cyanogenic glucosides linamarin and lotaustralin, which are defense compounds that can be hydrolyzed to liberate toxic hydrogen cyanide (Hagg et al., 2014). The amount and the ratios of the linamarin and lotaustralin in Z. filipendulae are tightly regulated and only to a low extent reflect the ratio in the ingested food-plant. Z. filipendulae adjusts the de novo biosynthesis of CNglcs by regulation at both the transcriptional and protein level depending on the food plant composition. Ultimately, larva saves energy and nitrogen while maintaining an effective defense system to fend off predators. By using in situ PCR and immunolocalization, the biosynthetic pathway was resolved to the larval fat body and integument, which infers rapid replenishment of defense compounds following an encounter with predator. The de novo biosynthesis of CNglcs in Z.filipendulae preceded the ability to sequester, and facilitated a food plant switch to cyanogenic plants, after which sequestration could evolve.

Lotus japonicus contains the two cyanogenic glucosides, linamarin and lotaustralin, and the non-cyanogenic hydroxynitriles, rhodiocyanoside A and D, with rhodiocyanoside A as the major rhodiocyanoside. The plants are acyanogenic either because they do not synthesize

CNGs, or because they lack the β-glucosidase required for degradation and HCN release (Jones, 1988). Rhodiocyanosides are structurally related to cyanogenic glucosides but are not cyanogenic (Saito et al., 2012). Between different insects, response to the presence of CNGs in leaves varies from total indifference to evident distaste. After starvation, insects are generally more willing to feed on cyanogenic L.corniculatus leaves. This indicates that for each insect, the deterrent capabilities of CNGs are dependent on the immediate demand for food calories (Compton and Jones, 1985). Among those species that rely on plants containing the CNGs as a major food source, there was a lack of selectivity against CNGs, which probably reflects specialized adaptations for a cyanogenic diet. High tolerance to CNGs may be characteristic of many polyphagous lepidoptera species, and accordingly, the role of CNGs in protection of plants from herbivores must be assessed on a species to species basis. The cyanogenic glucosides and rhodiocyanosides pathways share CYP79Ds to obtain (Z)-2-methylbutanalxime from I-isoleucine, whereas the subsequent conversion are catalyzed by different P450s. The aglycone of rhodiocyanoside A forma the cyclic product 3-methyl-2(5H)-furanone. Furanones are known to possess antimicrobial properties indicating that rhodiocyanoside A may have evolved to serve as a phytoanticipins that following β-glucosidase activation and cyclization of the formed give rise to a potent defense compound (Saito et al., 2012). CNGs in many plant species may serve as a warning to generalist insect herbivores that the plant is unpalatable. CNGs are recognized by a wide variety of herbivores, and is therefore a relatively cheap type of plant defense, and because HCN liberation only occurs after tissue damage, hence conserving materials and reducing the risks that adapted herbivores gain the ability to use HCN as an attractant (Compton and Jones, 1985).The most likely overall function of CNGs appears to be to deter herbivores that would casually try to feed on cyanogenic plants (Jones, 1988).

Cyanogenic glucosides in Arthropoda Cyanogenic glucosides are among the most widespread defense chemicals of plants (Jensen et al., 2011; Simon et al., 2010). Upon plant tissue disruption, these glucosides are hydrolyzed to a reactive hydroxynitrile that release toxic hydrogen cyanide (Agerbirk and Olsen, 2012). Many mite and lepidopteran species can thrive on plants defended by cyanogenic glucosides, but the nature of enzyme known to detoxify HCN to β-Cyanoalanine remained enigmatic. Phylogenetic analysis showed that the gene is a member of the cysteine synthesis family horizontally transferred

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from bacteria to phytophagous mites and lepidoptera. The recombinant mite enzyme had both β- cyanoalanine synthesis and cysteine synthase activity but enzyme kinetics showed that cyanide detoxification activity was strongly favored. An ancient horizontal transfer of the gene originally involved in Sulphur amino acid biosynthesis in bacteria was copied by herbivour arthropods to detoxify plant produced cyanide (Wybouw et al., 2014) In contrast to the taxonomically widespread distribution of CNGs within the plant kingdom presence of CNGs in animals appears to be restricted to a single phylum out of the currently known 31 Arthropod species (Nahrstedt, 1996). Within arthropods, presence of CNGs seems to the restricted to members of Chilopoda (centipedes), Diplopoda (millipedes) and in particular to Insecta, CNGs have hitherto only been found in Coleoptera (beetles), Heteroptera (true bugs) and in Lepidoptera (butterflies and moths) (Nahrstedt, 1988). Chilopoda, Diplopoda and some Coleoptera (Paropsis atomaria, Chrysophtharta variicollis and C. amoena) contain aromatic CNGs in their defensive secretions. Three beetle species appear to de novo synthesize CNGs as these are not present in their diet. Two species of diplopods (Oxidus gracilis and Harpaphe ahydeniana) have evolved biochemical pathways for cyanogenic glucoside biosynthesis and degradation that involve very similar or identical intermediates compared to those known to be used by higher plants (Duffey, 1981). Species H. haydeniana has cyanogenic glands that contain β-glucosidase and α-hydroxynitrile lyase activity, physically separated from the part containing CNGs. This prevents untimely the release of HCN, but offers the possibility of immediate and combined ejection and thereby mimics the phytoanticipin defense effect in plants. In contrast to other arthropod groups, many members of the Lepidoptera are able to de novo synthesize CNGs as well as to sequester it from their host plants. Only a single species of another arthropod group, the Heteroptera, has been proposed to be able to sequester CNGs from its host plant. Furthermore, members of the lepidoptera contain mainly aliphatic CNGs as opposed to other arthropod groups that are only known to contain aromatic CNGs. Insect evolved at least 390 Million years ago, so Diplopoda and Chilopoda have evolved from a common ancestor, they shared with Hexapoda at an earlier time point. This time span combined with the fact that many hexapod groups do not contain CNGs, and that those groups that do, contain different types of CNGs, points to convergent evolution of CNGs in these clades rather than to homology. The presence of CNGs has only been examined in a few species

of arthropods, so the distribution of CNGs may in fact be broader than currently recognized.

Cyanogenic glucosides in Zygaenidae

The toxic amino acid β-cyano-L-alanine is shown to be present in 21 species of the Zygaenidae, which are known to produce and store the cyanoglucosides linamarin and lotaustralin. β-cyanoalanine is also reported for the first time from other cyanogenic lepidoptera, i.e. the Acraeinae and the Heliconiinae of the nymphalid family and other species of the Nymphalidae, have not been known to be cyanogenic before. Cyanogenesis, including the presence of β-cyano-alanine and the cyanoglucosifes linamarin and lotaustralin is demonestrated for the first time from Heterogynidae (Witthohn and Naumann 2014). Thirty-nine species of the Zygaenidae have been investigated qualitatively and quantitatively for the cyanogenic glucosides linamarin and lotaustralin. Both glucosides are widely distributed within the genus Zygaena and related genera in different ratios and different quantities, suggesting that this is a common marker of the Zygaenidae. During the whole life cycle of the Z. filipendulae both glucosides are present in every stage of the development within an increasing linamarin: lotaustralin ratio from post-diapause larvae over all stages to newly emerged larvae. Resistance of Zygaena species to HCN is well known from very beginning and species can for example remain in a concentrated atmosphere of HCN for an hour and still revive quickly when removed to clean air (Naumann et al., 1999).

Linamarin and lotaustralin distribution in Zygaenidae

Since cyanogenic glucosides are restricted in animals. Among arthropods the six spotted burnet moth, Zygaena filipendulae is a model system to study cyanogenic glucosides in moths. Recently in post man butterfly, Heliconius melpomene, which also contains the cyanogenic compounds but the knowledge about the underlying mechanism is scare (Zagrobelny, 2014). Z.filipendulae are known for their high content of cyanogenic glucosides which they use as a defense chemicals (Zagrobelny and Moller, 2011), because they liberate toxic hydrogen cyanide upon breakdown. Since Z.filipendulae contains linamarin and lotaustralin which are sequestered from the host Lotus corniculatus as well as de novo biosynthesis from the amino acids valine and isoleucine, respectively (Jensen et al., 2011).The biosynthetic pathway of cyanogenic glucosides in these insects consists of the three enzymes CYP405A2, CYP332A3 and UGT33A1, and it turns out that the pathway is

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remarkably similar to the pathway in the plants (Takos et al.,2011), although clearly convergently evolved (Jensen et al., 2011). The Zygaena larvae prefer to feed on the highly cyanogenic lotus plants over lowly cyanogenic or acyanogenic lotus plants, probably to optimize the amount of cyanogenic glucosides available for sequestering (Zagrobelny et al., 2007b). In addition to serving as defense compounds, the two cyanogenic glucosides have previously been shown to play an intimate role in the mating process of the moth, in the course of which the male transfers the two compounds to the female (Zagrobelny et al., 2013, Zagrobelny et al., 2014). The cyanogenic glucosides are taken up during sequestration by Z.filipendulae larvae and are rapidly distributed to all tissues (Zagrobelny et al., 2014). Z.filipendulae have evolved adaptations to facilitate sequestration of the intact cyanogenic glucosides namely a high PH in the gut, fast feeding and the use of a leaf snipping feeding mode (Pentzold et al., 2014).

Defensive secretion and cuticular cavities

As a defensive reaction against predators (shrews, hedgehogs, starlings and carabid beetles), larvae of Zygaenini species may release highly viscous, colorless fluid droplets from cuticular cavities placed on their dorsal side. The toxic defensive droplets appear on the cuticular surface upon contraction of irritated segments (Franzl and Naumann, 1985). The droplets from Z. trifolii has been shown to be composed of various constuients like linamarin and lotaustralin (7% CNGs), β-cyanoalanine (0.3%), proteins (8%, including β-glucosidase) and water (Witthohn and Naumann, 1984). A linamarin: lotaustralin ratio of 1:1 was measured in the defensive secretion whereas that of the haemolymph of the Zygaena larvae has 19:1 (Franzl et al., 1986), which indicates that lotaustralin is transported more effectively than linamarin, maybe as a result of increased lipid solubility caused by its longer aliphatic side chain (Franzl et al., 1986). Alternatively, the change in ratios could reflect a slower turnover rate of lotaustralin compared to linamarin in larvae of Zygaena. In the Z. trifolii two morphologically different types of cavities have been found; the larger cavities release their contents as a response to a slight irritation, whereas the smaller cavities react following severe irritation and therefore, release much smaller droplets. Defense droplets may be reabsorbed a few seconds after irritation has stopped. In contrast to most diplopods and chilopods that have specialized cyanogenic glands, there are no gland cells or cuticular ducts leading through the cuticle into the cavities in Zygaena larva, and no special morphological adaptation for

secretion has been developed in the epidermis (Franzl and Naumann, 1985).

Metabolism and detoxification of cyanogenic glucosides

For the synthesis of linamarin and lotaustralin in plants, the same enzyme system uses valine and isoleucine as precursors, respectively, but with different catalytic efficiencies toward the substrate amino acids. This is reflected in the relative amounts of linamarin and lotaustralin in cassava (Andersen et al., 2000) and in L. japonicas, further the biosynthesis may involve the same set of enzymes for both amino acid precursors (Takos et al., 2010; Olsen et al., 2014). Lotus japonicus also produce the non-cyanogenic γ- and β-hydroxynitrile glucosides rhodiocyanoside A and D using a biosynthetic pathway and the branches off from lotaustralin biosynthesis (Lai et al., 2014). Further in plants, exogenously administered N-hydroxyamino acids, aldoximes and nitriles can be incorporated into CNGs (Jones et al., 2000) and same results are obtained with Zygaena species, and suggesting that the biosynthetic pathway for CNGs in Zygaena is identical to the pathway in plants.

The β-Glucosidase dependent HCN release has been observed from different life stages of many Zygaenidae species (Hagg et al., 2014; Pentzold et al., 2014). β-Glucosidase from Z. trifolii is a dimer consisting of two supposedly identical 66 kDa subunits. The β-glucosidase exhibits a strong activity towards the endogenous substrates linamarin and lotaustralin, with lotaustralin being a better substrate. β-Glucosidase activity was found exclusively in haemolymph of Bombyx mori, which has a pH of 6.2 (Franzl et al., 1989). At pH 6.2, the β-glucosidase is present in an almost inactive state but the enzyme becomes active when pH decreases (Nahrstedt and Muller, 1993). This may point to a situation where stomach acid from a predator will activate the β-glucosidase leading to a rapid and strong release the HCN. Mg++ ions are inhibitors of the Zygaena β-glucosidase; this is the only example of a β-glucosidase which is inhibited by alkaline earth metal ions (Nahrstedt and Muller, 1993).

In vitro experiments showed a noncompetitive or mixed type inhibition of the linamarase by the alkaline earth metal ions Mg++ and Ca++.When fully activated by chelating agents the linamarase cause strong cyanogenesis and liberation of the HCN in the haemolymph. Lowering the PH from its physiological valve of 6.2 in a non-chelating buffer to 3.6 cause full cyanogenesis. Bothe the ions are the natural inhibitors of the linamarase in the intact

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insect (Nahrstedt and Muller, 2009). The α-Hydroxynitrile lyase was found from the haemolymph of Z. trifolii and characterized as a FAD-containing enzyme (Lechtenberg, 2011; Nahrstedt, 1996).The β-Cyanoalanine synthase activity is found in the various parts in different proportions e.g, integument (22%), fat body (27%) and gut (12%) of Zygaena larvae, with the highest activity of the enzyme in the gut. β-Cyanoalanine synthase detoxifies HCN to β-cyanoalanine, which accumulates in haemolymph (75%) and integument (16%). Suggesting, that these insects can easily detoxify HCN and the members of the Zygaenidae seem to use the β-Cyanoalanine pathway as the sole pathway for detoxification of HCN, since rhodanese has not been found in any species containing β-cyanoalanine.

6. Distribution, detoxification, synthesis and sequestration of Cyanogenic glucosides in lepidopteran insects

Linamarin and lotaustralin distribution

Cyanogenesis describes the ability of living organisms to liberate hydrogen cyanide (Ballhorn et al., 2007) from the stored cyanogenic glucosides, cyanogenic lipids or cyanohydrins on tissue damage by hydrolysis and/or decomposition (Lechtenberg, 2011). Additionally, some species of Heliconius genera are able to sequester cyclic cyanogenic glucosides from their host plants (Zagrobelny et al., 2009; Engler et al., 2000). Nevertheless, during the evolution of these butterflies, the ability to sequester this compound seems to be negatively correlated with the ability to produce aliphatic cyanogenic glucosides by the de novo pathway (Engler-Chaoust and Gilbert 2007). Furthermore, in Heliconius butterflies it has not been shown whether the ability to sequester cyanogenic glucosides is restricted to cyclic compounds or whether it also occurs with aliphatic, such as linamarin and lotaustralin. The linamarin and often the lotaustralin is distributed in various families of the lepidopterans (Hagg et al., 2014), and is accumulated in all life stages of butterflies from the Heliconiinae, Acraeinae, Nymphalinae and Polyommatinae (Papilionoidea) groups. The butterflies de novo synthesize linamarin and lotaustralin from valine and isoleucine, respectively, and because linamarin and lotaustralin are not present in their larval host plants (Passifloraceae) therefore the sequestration cannot occur (Engler et al., 2000). The amount of linamarin is higher than that of lotaustralin as also observed in imagines of Zygaenidae, and the CNGs have the same bodily distribution as observed in Z. trifolii. The butterflies also contain monoglycoside cyclopentenyl cyanogens, probably sequestered

from their host plants, at the larval stage (Engler et al., 2000).

Detoxification and sequestration

The α –hydroxynitriles (cyanohydrins) are stabilized by glycosylation (Gleadow and Moller, 2014). However, the cyclopentenyl cyanogens are detoxified and sequestered by lepidopterans (Hagg et al., 2014; Lechtenberg, 2011) but the toxic metabolite monoglycoside cyclopentenyl cyanogens, obtained and sequestered from the host plant are detoxified by heliconius sara (Papilionoidea) using a unique enzymatic mechanism not found in the host plants. This mechanism involves substitution of the nitrile group of the cyanohydrins function with a mercapturic group. The reaction mechanism involved has not been elucidated and it remains to be shown whether free cyanide is released during the reaction or whether the nitrile group is transferred to a specific acceptor molecule, like cysteine, as a part of the reaction sequence. The cyanogenic glucoside-derived mercaptic compound was not found in the host plant (Engler et al., 2000). This is in contrast to the mechanism proposed for H. melpomene (Papilionoidea) and Z. trifolii. The presence of β-cyanoalanine synthase was demonstrated in Clossiana euphrosyne (Papilionoidea) which confirms that some Papilionoidea species detoxify HCN using the same mechanism as Zygaenidae species. However, in the model plant Arabidopsis, the cyanide degradation pathway is present through which degradation of the cyanide to ammonia and carbon dioxide occurs (Kebeish et al., 2015).

The cyclopentenyl glycine derived CNGs from its host plant Turnera ulmifolia (Szewczyk and Zidorn 2014) were hypothesized and sequestered by Eutoptia hegesia (Heliconiinae, Papilionoidea). The sequestering was proposed because of significantly higher CNG levels in larvae reared on cyanogenic plants compared to siblings reared on acyanogenic plants (Jensen et al., 2011). Accordingly, the ability to both de novo synthesize and sequester the same CNGs appears not to be an exclusive feature for Zygaenidae species

Summary and Conclusions

In nature the cyanogenic glucosides serve as good chemotaxonomic markers for plant relatedness. The result of natural mutations have caused the loss of the ability to produce one or more of the enzymes involved in cyanogenic glucoside biosynthesis and/or in the degradation of CNGs. The insects only need cyanogenic host plants to minimize their own biosynthesis of CNGs. The

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transfer of genes across species using genetic engineering enables the design of plants with an altered qualitative and quantitative content of natural products thereby bypassing millions of year of co-evolution of plants and their herbivores.

ACKNOWLEDGEMENT

Authors are highly thankful to the Department of Science and Technology, New Delhi, India for providing the financial assistance to compile this paper. Author is also thankful to SKUAST-K, srinager for providing the internet facilities.

Contradiction of authors: No contradictions of authors at all.

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