Jump to ContentJump to Main Navigation
Show Summary Details
More options …

Zeitschrift für Naturforschung C

A Journal of Biosciences

Editor-in-Chief: Seibel, Jürgen

Editorial Board: Aigner , Achim / Boland, Wilhelm / Bornscheuer, Uwe / Hoffmann, Klaus

12 Issues per year


IMPACT FACTOR 2017: 0.882
5-year IMPACT FACTOR: 0.912

CiteScore 2017: 0.92

SCImago Journal Rank (SJR) 2017: 0.288
Source Normalized Impact per Paper (SNIP) 2017: 0.448

Online
ISSN
1865-7125
See all formats and pricing
More options …
Volume 72, Issue 9-10

Issues

Biotechnological potential of insect fatty acid-modifying enzymes

Michal Tupec
  • Institute of Organic Chemistry and Biochemistry of the Czech Academy of Sciences, Flemingovo n. 2, 166 10 Prague 6, Czech Republic
  • Other articles by this author:
  • De Gruyter OnlineGoogle Scholar
/ Aleš Buček
  • Institute of Organic Chemistry and Biochemistry of the Czech Academy of Sciences, Flemingovo n. 2, 166 10 Prague 6, Czech Republic
  • Other articles by this author:
  • De Gruyter OnlineGoogle Scholar
/ Irena Valterová
  • Institute of Organic Chemistry and Biochemistry of the Czech Academy of Sciences, Flemingovo n. 2, 166 10 Prague 6, Czech Republic
  • Other articles by this author:
  • De Gruyter OnlineGoogle Scholar
/ Iva PichováORCID iD: http://orcid.org/0000-0001-5723-6143
Published Online: 2017-07-25 | DOI: https://doi.org/10.1515/znc-2017-0031

Abstract

There are more than one million described insect species. This species richness is reflected in the diversity of insect metabolic processes. In particular, biosynthesis of secondary metabolites, such as defensive compounds and chemical signals, encompasses an extraordinarily wide range of chemicals that are generally unparalleled among natural products from other organisms. Insect genomes, transcriptomes and proteomes thus offer a valuable resource for discovery of novel enzymes with potential for biotechnological applications. Here, we focus on fatty acid (FA) metabolism-related enzymes, notably the fatty acyl desaturases and fatty acyl reductases involved in the biosynthesis of FA-derived pheromones. Research on insect pheromone-biosynthetic enzymes, which exhibit diverse enzymatic properties, has the potential to broaden the understanding of enzyme specificity determinants and contribute to engineering of enzymes with desired properties for biotechnological production of FA derivatives. Additionally, the application of such pheromone-biosynthetic enzymes represents an environmentally friendly and economic alternative to the chemical synthesis of pheromones that are used in insect pest management strategies.

Keywords: fatty acyl desaturases; fatty acyl reductases; lipases; pheromones

1 Introduction

The fatty acids (FAs) and FA derivatives are a diverse group of compounds with a range of applications. They may be used as food supplements, cosmetics, adhesives, industrial lubricants, polymer plasticizers and stabilizers [1], and raw materials for further chemical processing (reviewed by Metzger and Bornscheuer [2]). Economic production of these compounds from affordable raw materials, such as hydrocarbons from petroleum refining and vegetable or animal oils, is well established at an industrial scale [3]. However, traditional procedures are not applicable to the production of unconventional FAs and FA derivatives, e.g. with double bonds in unusual positions, that might be useful for specific applications [2]. The utilization of metabolically engineered organisms or heterologous production of engineered enzymatic catalysts is promising tools for the production of such FA derivatives. Plants and microorganisms metabolically engineered to produce polyunsaturated FAs (PUFAs) [4], [5], [6] and fatty alcohols [7] have been tested. Here, we focus on the potential application of insect enzymes to the biotechnological production of unusual FA derivatives.

Many insect species use a diverse range of FA-modifying enzymes to synthesize pheromones that mediate communication among individuals of the same species. The FA-derived pheromones are the most common [8], encompassing thousands of compounds and mixtures of compounds [9]. Insect pheromone-biosynthetic enzymes presumably evolved via a divergence of the original functions of the FA-biosynthetic and FA-modifying enzymes participating in insect primary metabolism [10]. The diversification of the pheromone-biosynthetic enzymes likely has been driven by evolutionarily imposed requirements on sex pheromone signal specificity [11], [12]. A broad spectrum of insect pheromone-biosynthetic enzymes has already been functionally characterized, and transcriptomic sequencing (RNA-seq) of pheromone glands using next-generation sequencing has identified new candidates for characterization [13], [14], [15], [16], [17], [18], [19]. RNA-seq of other insect glands and tissues may open the door to the discovery of many additional enzymes with remarkable biosynthetic capabilities [8].

Among insects, moths (Lepidoptera) have received the greatest scientific attention focused on pheromone biosynthesis. The moth female sex pheromones, which attract conspecific males, are generally FA-derived alcohols, acetates, aldehydes, esters, hydrocarbons, or epoxides of various hydrocarbon chain lengths and contain zero, one, or multiple double bonds (or triple bonds) placed apart from each other, methylene interrupted, or conjugated. The number, position, and stereochemistry of double bond(s) in the chains of the FAs influence their biophysical properties and the properties of the resulting lipids [20] and are critical for the biological activity of the FA-derived pheromones.

Moths include numerous pest species that cause billions of Euros of damage annually in forestry and agriculture. For example, the diamondback moth (Plutella xylostella), a pest of rapeseed (Brassica napus), is estimated to cost almost 4 billion Euros annually on combined pest management and residual losses in crop production [21]. Pest management strategies that employ insect pheromones to trap and kill, monitor insect abundance, or confuse insect pheromone communication channels are environmentally friendly alternatives to traditional, widespread strategies of fighting insect pests, such as the use of insecticides. Synthetic pheromones have been used to fight pest moth species [22]; however, new biotechnological approaches in which pheromones are produced with the help of genetically modified plants or microorganisms are currently being developed [23], [24], [25]. Here, we not only review this application of insect pheromone-biosynthetic enzymes but also propose as-yet unexplored or unexploited applications that may benefit from the catalytic potential of these enzymes.

2 Insect enzymes catalyzing biosynthesis of FA-derived pheromones

Efforts to understand the basis of insect pheromone biosynthesis have generated insights into the enzymatic specificities of several classes of FA-modifying enzymes. The majority of information is available for one insect group – the moths [18]. FA-modifying enzymes from other insect orders, such as beetles (Coleoptera) [26], flies (Diptera) [27], [28], [29], crickets (Orthoptera) [30], [31] and bees (Hymenoptera) [32], [33], [34], are substantially less well explored and usually connected to primary FA metabolism rather than pheromone biosynthesis.

Two enzyme groups that are encoded by large multigene families in insects – the membrane fatty acyl desaturases (mFADs) [35], [36] and the fatty acyl reductases (FARs) [37] – have attracted the most research attention.

2.1 mFADs

The mFADs (EC 1.14.19.–) belong to a superfamily of oxygen-dependent membrane di-iron-containing enzymes that share common features including a conserved tripartite histidine-rich motif coordinating two iron ions in the active center. These enzymes catalyze the highly energy-demanding removal of hydrogen from an unactivated fatty acyl at a precise position along the hydrocarbon chain. The process involves a reactive oxo-intermediate formed by the activation of molecular oxygen. The net result of the desaturation reaction is the introduction of a double bond into the fatty acyl chain (and reduction of molecular oxygen to water) [38] (Figure 1). The evolutionarily unrelated class of apparently convergent soluble FADs [39] is expressed exclusively in the stroma of plant plastids, and these enzymes desaturate fatty acyls bound to an acyl carrier protein.

Schematic depiction of the reactions catalyzed by membrane fatty acyl desaturases (mFADs) and fatty acyl reductases (FARs).
Figure 1:

Schematic depiction of the reactions catalyzed by membrane fatty acyl desaturases (mFADs) and fatty acyl reductases (FARs).

The mFADs are present in the cell membranes of some bacteria [40], [41], thylakoid and cytoplasmic membranes of cyanobacteria [42], thylakoid and cytoplasmic membranes of plants, and ubiquitously in eukaryotic endoplasmic reticulum (ER) membranes [35]. The ER mFADs, on which we focus throughout this review, use an electron pair supplied by nicotinamide adenine dinucleotide (NADH) via an electron transport system consisting of NADH:cytochrome b5 reductase and cytochrome b5 [38]. A slight modification of this electron-supply chain is seen in some mFADs, in which a cytochrome b5 domain is fused to the N- or C-terminus of the desaturase [43], [44].

The research conducted on mFADs primarily aims to identify the determinants of desaturase specificity, enabling engineering of mFADs that produce economically or industrially relevant FAs, such as the PUFAs that serve as nutritional supplements or starting materials in the chemical industry [45], [46]. Another major research goal is to uncover the mechanistic details of FA desaturation, which might enable rational design of specific inhibitors targeting either the mFADs involved in human metabolic diseases, such as diabetes or obesity [47], or the mFADs essential for pathogenic microorganisms, such as pathogenic yeasts [48], [49], [50] and trypanosomatids [51], [52]. Basic research also aims to uncover the molecular basis of insect pheromone evolution by studying the pheromone-biosynthetic mFADs [18].

2.1.1 mFAD properties

The mFADs display several enzymatic activities. They may be (i) stereospecific, i.e. introduce a double bond into fatty acyl chains in either an E or Z configuration, or (ii) regiospecific, i.e. they prefer a particular position along the fatty acyl hydrocarbon chain for double bond introduction, usually marked as ΔX or ZX-, EX- [53]. The mFADs also display diverse substrate specificities; they may prefer a particular chain length, the presence of pre-existing double bond(s) at particular position(s), stereochemical configuration and a head group of fatty acyl substrate (e.g. acyl-CoA or acyl-lipid substrates) [54]. Moreover, hydroxylated (and less commonly acetylenated, i.e. bearing a triple bond) products may accompany the desaturated products as a result of the mechanistic similarity between the reaction mechanisms [55], [56]. Several mFADs exhibit a fatty acyl conjugase activity – the ability to produce a system of conjugated double bonds by a reaction mechanism involving a shift in the position of a pre-existing double bond [57], [58]. The knowledge gained from mFAD characterization indicates that although numerous mFADs are highly specific and produce only a limited set of unsaturated products, they can follow more than one specificity mode under particular conditions, such as the presence of different substrates or during sequential biosynthesis of FAs with multiple double bonds [59], [60]. Most likely, the insect mFADs evolved an excessively wide range of specificities in connection with pheromone biosynthesis [35].

The mFADs producing Z9-monounsaturated FAs are the most widespread eukaryotic desaturases, followed by mFADs with Z5-, Z6-, Z12- and Z15-specificities. The majority of animal and insect mFADs desaturate fatty acyl-CoA groups, except some animal mFADs involved in PUFA biosynthesis that prefer fatty acyl lipids [61], [62]. The experimental evidence on the identity of the mFAD substrate head group is, however, scarce [54].

The first functionally characterized insect mFAD was isolated from the cabbage looper moth (Trichoplusia ni) and exhibited Z11-desaturase specificity. An FA-derived pheromone component containing a double bond at position Δ11 is present in many moth species [63]. Subsequently, more than 50 distinct insect FAD genes have been identified, cloned and functionally characterized [18], [36] (Table 1).

The moth mFADs reflect the diverse desaturase specificities in insects. In addition to the ability to introduce Z-double bonds, moth mFADs can catalyze the introduction of rather uncommon E-double bond in nature [64], [65], [66], [67], [68] or produce a mixture of E- and Z-unsaturated FAs [69], [70]. The preferred fatty acyl chain length may be C14 [27], [66], [67], [68], [70], [71], [72], [73], C16 [74], or C18 [63], [75], [76], but some mFADs can desaturate a broad range of fatty acyl chain lengths, such as C14-C20 [67], [77]. The positions of the introduced double bond include ∆9, ∆4 [33], ∆5 [72], ∆6 [64], [78], ∆8 [65], ∆10 [69], [78], ∆11 [10], [63], [65], [66], [67], [74], [75], [76], [79], [80], ∆13 [81], and ∆14 [70], [82]. Δ12 mFADs have also been identified, but they are involved in primary metabolism during the production of methylene-interrupted PUFAs [31] rather than pheromone biosynthesis. The moth mFADs can also introduce a double bond at the terminal position between the penultimate and ultimate carbon atoms [78], [83] and can produce FAs with a system of isolated double bonds [64] or a system of conjugated double bonds [10], [65], [66], [76], [79], [80], [82]. Among the more bizarre desaturation reactions, an mFAD identified in the processionary moth, Thaumetopoea pityocampa, can introduce triple bonds into the FA chains [81], resembling in activity an mFAD described in the plant of Crepis genus [84]. Two moth Δ11-mFADs have been shown to exhibit minor Δ11-hydroxylation activity [85]. Table 1 shows functionally characterized insect mFADS.

Table 1:

Overview of functionally characterized insect fatty acyl desaturases (mFADs).

2.2 FARs

The alcohol-forming FARs [EC 1.2.1.84, systematically long-chain acyl-CoA: nicotinamide adenine dinucleotide phosphate (NADPH) reductases] belong to a family of oxidoreductases and catalyze reduction of activated FAs to the corresponding fatty alcohols by means of a four-electron process employing a reduced dinucleotide (either NADPH or NADH) as a reductant [91]. The reaction takes place on the thioester moiety through a putative aldehyde intermediate, which is usually not released from the enzyme-substrate complex [92]. In addition to the alcohol-forming FARs, there are also reductase enzymes that produce aldehydes from fatty acyl-CoAs (aldehyde-forming reductases) and fatty alcohols from aldehydes (aldehyde reductases) [91], [93]. The FARs are most probably localized to ER membranes [94], [95].

The fatty alcohols, which are usually defined as primary alcohols having more than 12 carbon atoms in the chain, are naturally abundant FA derivatives that play a variety of biological roles. The fatty alcohols are precursors of waxes that serve as surface-protective compounds in plant pollen [96], preventing excessive water loss in insects [29], and are secreted as skin-, eye-, or feather-protective compounds in mammals and birds [97], [98]. The fatty alcohols are also components of ether lipids, which are abundant in the cell membranes of cardiac, nervous and lymphoid tissues [99]. In some plants such as jojoba (Simmondsia chinensis), marine crustaceans (e.g. Calanus finmarchius), and protists (e.g. Euglena gracilis), fatty alcohols serve as precursors of energy-storing waxes [100], [101], [102]. Notably, the higher volatility of fatty alcohols and acetates, as compared to the respective FAs, may have led to their utilization as airborne signals – pheromones and scents. These compounds may serve as pheromone components in reptiles [103] and some mammals, such as deer [104], but above all, they are among the main pheromone components in insects, including moths [105], termites [106], bees [107], [108], [109], and wasps (reviewed by Tillman et al. [110]).

2.2.1 FAR properties

Virtually all organisms produce one or multiple types of fatty alcohols of various chain lengths and degrees of unsaturation. The majority of the findings available to date suggest that the FARs in general exhibit diverse substrate specificities with respect to the fatty acyl chain length and unsaturation state of the substrate. The pool of fatty acyls available for reduction in the host organism can also influence the apparent FAR specificity [111], [112], [113].

The first FARs studied by genetic methods were plant FARs from jojoba and wheat [100], [114]. Heterologous expression of plant FARs in Escherichia coli, rapeseed (B. napus), and Saccharomyces cerevisiae led to a range of saturated and unsaturated alcohols of C14 to C26 chain length, depending on the host organism [100], [114], [115]. Besides plant FARs, microbial FAR genes [102], [116], [117] and FARs from vertebrates have also been isolated and functionally described [97], [98].

The insect reductases first received attention not long after the enzymatic characterization of their plant orthologs. They have been studied extensively, primarily in moths. In silk moth (Bombyx mori), researchers discovered a pheromone gland-specific FAR [118] that is able to convert E10,Z12-16:2 FA precursor to the corresponding alcohol bombykol, the main component of the female sex pheromone. Since then, a range of moth pheromone-biosynthetic FARs have been isolated and functionally characterized [89], [111], [119], [120], [121], [122], [123] (Table 2). Although moth FARs usually exhibit a broad substrate preference [77], [89], [111], [120], [122], [123], some of them display specificity to unsaturated substrates with a double bond in either the E or Z configuration [119], a particular chain length and double bond position [120], or a system of conjugated double bonds [118]. Given the multiple FAR paralogs generally present in insect genomes [13], [14], [16], [17], [111], [124] and limited knowledge of their properties, there is a demand to perform functional characterization of these biologically and biotechnologically relevant enzymes.

Table 2:

Overview of functionally characterized insect fatty acyl reductases (FARs).

FARs from insects other than moths are virtually unexplored. A FAR presumably involved in biosynthesis of fatty alcohol precursors of waxes has been identified in Drosophila [29], and honeybee (Apis mellifera) FAR capable of reducing hydroxylated fatty acyl precursors and saturated C16-C22 fatty acyls has been characterized [34]. FARs from pheromone-producing glands in the butterfly Bicyclus anynana reduce C14 and C16 fatty acyls [86]. At least 13 FAR genes have been identified in the labial gland of Bombus terrestris by RNA sequencing [14], and these are currently being functionally characterized in our laboratory. Such a spectrum of FARs from various organisms with distinct substrate specificities presents a potential enzymatic toolbox for tailored biotechnological production of fatty alcohols. Additionally, comparative analysis of these FARs might help uncover the determinants of FAR specificity.

2.3 Other enzymes

In addition to mFADs and FARs, other FA-modifying and FA-biosynthetic enzymes have been studied both in vivo and in vitro. These include (i) acyltransferase involved in synthesis of FA-storing triacylglycerols [125], [126]; (ii) lipases and esterases involved in hydrolytic release of FA pheromone precursors from storage triacylglycerols [127], [128], [129], formation of FA ethyl esters [130], or primary metabolism [131], [132], [133]; and (iii) cytochrome P450, involved in the oxidative decarbonylation of FAs to hydrocarbons [134]. However, for the majority of insect pheromone biosynthetic steps, the genes and enzymes involved have yet to be identified and characterized. Among these are enzymes catalyzing (i) FA elongation [135], [136] and FA chain shortening [137], [138], [139], [140]; (ii) epoxide group formation [141], [142]; (iii) acetate ester formation [143], [144]; (iv) oxidation of fatty alcohols to aldehydes [19], [145], [146], [147]; and (v) FA biosynthesis, e.g. acetyl-CoA carboxylase [148] and fatty acid synthase [149].

3 Enzyme engineering

The specificity of insect pheromone-biosynthetic enzymes can evolve abruptly, presumably as a consequence of their role in reproductive isolation and speciation [11], [12]. The process of functional divergence of pheromone-biosynthetic enzymes, which can be observed in insect subpopulations or closely related species, generates biocatalysts that are highly similar in protein sequence yet distinct in their enzymatic properties. These enzymes are convenient model systems to study the mechanisms of enzyme specificity determinants [82], [119], [150]. Importantly, the amassed knowledge about the function of the FA-modifying enzymes could be used to design novel enzymes with desired enzymatic properties.

3.1 Structural determinants of mFAD function

Substantial research effort has been put into the identification of specificity determinants of mFADs by either random mutagenesis or rational mutagenesis guided by topology predictions and sequence comparisons of mFADs with distinct specificities. The mFADs and their mutants are typically functionally characterized in the yeast S. cerevisiae [151] or in baculovirus-insect cell expression systems [79]. These experiments led to the identification of sequence determinants of both acyl-CoA and acyl-lipid mFAD specificities in diverse organisms, such as transmembrane helices or conserved histidine-rich motifs [55], [59], [152], [153], [154], [155], [156], [157], [158], [159], [160], [161], [162]. Buček et al. [82] identified a critical amino acid residue in the transmembrane domain of an mFAD from Manduca sexta (MsexD3) that determines the specificity and ability of this desaturase to catalyze biosynthesis of FAs containing three conjugated double bonds via E/Z14 desaturation from diunsaturated FAs.

Recently, the crystal structures of two closely related mammalian mFADs with bound fatty acyl-CoA substrate provided the first direct structural insights into the mFADs [163], [164]. In agreement with previous topology predictions [151] and topology-mapping experiments [165], the crystal structures revealed four transmembrane α-helices and a large extramembrane portion of the enzyme including the active center localized on the cytosolic side of the ER membrane. The di-iron active center is coordinated by an ordered water molecule and nine conserved histidine residues. Eight of the coordinating histidines are organized in a tripartite histidine-rich motif, which was previously shown to be essential for mFAD activity [166]. The crystal structures provided direct experimental evidence for a kinked narrow hydrophobic substrate-binding tunnel, which extends approximately 24 Å into the enzyme interior and binds the fatty acyl tail of the fatty acyl-CoA substrate [164]. The kink in the binding tunnel is hypothesized to play a role in correct positioning of the fatty acyl chain toward the active center.

The availability of an mFAD crystal structure enables homology modeling of other related mFADs and can help increase understanding of experimentally obtained biochemical data. Indeed, the homology model of MsexD3 highlighted the prominent position of the critical residue Ile224 in the kink of the fatty acyl substrate binding tunnel, which presumably plays a critical role in positioning the substrate fatty acyl chain with respect to the di-iron active center [82]. Ding et al. found that reciprocal exchange of a single amino acid residue in moth ∆11 mFADs can switch between their E and Z desaturase specificities. The critical residue 258Glu/Asp is predicted to form a secondary coordination sphere of the active center iron ions, based on homology models [150].

3.2 Structural determinants of FAR function

The information about FAR structures is very limited. The N-terminal motif (IVF)X(ILV)TGXTGFL(GA) belonging to the Rossmann fold NAD(P)+ binding domain and the C-terminal FAR_C domain is conserved among the FARs [89], [96], [98], [100]. The common dehydrogenase/reductase active site motif YXXXK was experimentally confirmed to be indispensable for FAR enzymatic activity [167]. Few studies have addressed the sequence determinants of FAR specificity. A study on FAR5 and FAR8 from Arabidopsis, which prefer C18 and C16 acyls, respectively, showed that reciprocal domain swaps and single amino acid mutations (355Ala/Leu and 377Val/Met) in the C-terminal part of the sequence resulted in a transition between C16 and C18 substrate preference [167].

Mutagenesis experiments performed with insect FARs also indicate that a limited number of amino acid substitutions can profoundly change the enzyme specificity [119], [120].

Currently, there is no publicly available protein structure of a FAR, which presents a challenge for performing further mutagenesis studies to infer the mechanisms of FAR specificity determination and to engineer FARs with novel or desired enzymatic properties.

4 Applications of FA-modifying enzymes

The potential biotechnological applications arising from the diverse enzymatic capabilities of insect FA-modifying enzymes are centered mainly on the synthesis of insect pheromones for pest management. However, there are also other, mostly unexplored potential applications for these insect enzymes (Figure 2).

FA-modifying reactions that can be engineered in plant and microbial cells with the help of insect pheromone-biosynthetic enzymes. mFAD, fatty acyl desaturase; LIP, lipase; FAR, fatty acyl reductase; and FA, fatty acid.
Figure 2:

FA-modifying reactions that can be engineered in plant and microbial cells with the help of insect pheromone-biosynthetic enzymes. mFAD, fatty acyl desaturase; LIP, lipase; FAR, fatty acyl reductase; and FA, fatty acid.

Synthetic insect pheromones are used in a variety of pest management strategies in agricultural fields, forests, and urban areas to replace or complement traditional insecticides. The pheromones have several inherent advantages over insecticides: they are active in extremely small amounts (nanogram and sub-nanogram quantities), and they are specific toward the target species and generally non-toxic to other animals or humans (reviewed by Witzgall et al. [168]). Currently, synthetically prepared insect pheromones are used either as attractants for monitoring or mass trapping of insect pests, monitoring of other relevant insect species such as endangered species ([169], reviewed in [170]), or mating disruption of pest species via the release of synthetic sex pheromones that compromise olfactory communication and mate finding in insect pests [168].

The estimated total area of land treated with synthetic insect pheromones is approximately 10,000,000 ha worldwide and, for example, the global production of codlemone, the codling moth (Cydia pomonella) sex pheromone, reached 25,000 tons in 2010 [168]; it is primarily used in apple orchards. The pheromones have also been used widely in vineyards for mating disruption of the grapevine moth Lobesia botrana in Germany, Italy and California [168].

The establishment of economically viable synthesis of pheromones remains a major obstacle to scaling up the use of pheromone chemicals in pest control. In particular, FA-derived pheromone biosynthesis faces several challenging issues, such as the requirement for precisely positioning one or multiple double bonds into the synthesized FA-chain in a particular double bond configuration [171]. In this respect, employment of insect FA-modifying pheromone-biosynthetic enzymes or organisms heterologously expressing these enzymes may be a more economic and environmentally friendly option than traditional chemical synthesis.

The concept of producing pheromone chemicals in genetically modified plants (“pheromone farming”) or yeasts (“pheromone brewing”) has been tested [23], [25]. In these projects, the researchers semi-synthetically prepared moth pheromones by chemical reduction and consecutive acetylation of Z11-unsaturated FAs produced in genetically modified plants expressing moth Z11-mFAD. Ding et al. reconstructed a complete pheromone biosynthetic pathway by transforming the tobacco plant with a combination of two insect pheromone biosynthetic genes (mFAD and FAR) and two non-insect genes (thioesterase, which modified the length of de novo biosynthesized FAs, and acetyltransferase, which catalyzed the final pheromone biosynthetic step yielding the fatty alkyl acetates) [24]. The approach of combining insect and non-insect genes in the host organism is particularly useful in situations when suitable genes have not yet been isolated from insects. Plant “pheromone factories” possess another attractive feature: if proven non-harmful to the environment, they could be planted directly in fields, thus circumventing the necessity of isolating and dispersing pheromones from specialized dispensers for mating disruption [24].

The yeast S. cerevisiae is a frequently used host for metabolic pathway engineering [172] and represents a promising host organism for heterologous expression of FARs capable of producing biologically active pheromone chemicals [118]. The yeasts co-expressing FAR and Z11-mFAD have been used to produce alcohol pheromone precursors that can be subsequently isolated and chemically oxidized to the biologically active aldehyde pheromones [23].

Muñoz et al. [173] tested another approach to small-scale pheromone synthesis, using immobilized bacterial acetyl transferase to catalyze the terminal step of the biosynthesis of a di-unsaturated conjugated fatty alkyl acetate, a cotton leafworm (Spodoptera littoralis) sex pheromone.

In a semi-synthetic approach, insect enzymes would be used to produce the precursors of compounds that are difficult to access solely by traditional organic synthesis tools, and these would be further chemically modified [23], [24], [25]. Such an approach could yield FAs with multiple double bonds in uncommon positions and geometric configurations or possibly molecules bearing more than one functional group. The insect FARs, with their strict substrate specificity toward uncommon substrates, could serve to efficiently and specifically convert non-conventional FAs into their respective alcohols.

In many cases, handling isolated enzymes is difficult due to their transmembrane character (e.g. mFADs) or requirement for additional protein partners and coenzymes. The only known exception among the FA-modifying enzymes is the lipases, which do not require a coenzyme, are usually soluble, and have been shown to be stable and active in a variety of different environments (including organic and ionic solvents) [174], [175]. The use of lipases from microorganisms and animals has been widely established on an industrial scale [176]; for example, more than 10,000 tons of pure 1-phenylethylamine enantiomer are produced annually using lipase as a catalyst [177]. Lipases are abundant in insect genomes. The majority of them evolved after divergence of insect orders [178], and thus, they potentially possess properties not present among lipases from other organisms. Insect lipase specificities and enzymatic properties, however, remain poorly studied in general. The isolated lipases from non-insects have been used for enantioselective synthesis of pheromones from bark beetles [179], corn rootworms [180], rice moths [181], and many other insect species [182].

To the best of our knowledge, there is currently no commercial application of insect enzymes in the synthesis of oleochemicals, although the implementation of some insect enzymes could substantially expand the toolbox of FA modifications available to both organic chemistry and mass industrial production. One recent example from the research field is the use of honeybee FAR1 for the synthesis of wax esters in engineered S. cerevisiae co-expressing FA elongase and wax synthase [183].

5 Challenges and future perspective

One of the main drawbacks to the use of genetically modified organisms for production of pheromones and other FA-derivatives is the cost of developing such transgenic organisms. Additionally, substantial optimization and scale-up of production capacity are required to convert the heterologous expression systems used in research into feasible production organisms. So far, efforts to produce insect pheromones in host organisms have led to (i) 2 mg/L of fatty alcohols in S. cerevisiae liquid culture [23]; (ii) 44 mg of unsaturated FA-derived methyl ester per kg of plant material [25]; and (iii) hundreds of mg of unsaturated FAs, tens of mg of the respective fatty alcohols, and several mg of the final acetates per kg of fresh leaf tissue from a Nicotiana benthamiana expression system [24]. In host systems transformed with non-insect enzyme sequences, the production of fatty alcohols and wax esters in bacteria (E. coli or Cyanobacteria) or yeasts (S. cerevisiae, Rhodosporidium toruloides, or Yarrowia lipolytica) can reach hundreds to thousands of mg/L cultivation medium after lab-scale optimization [7], [184], [185], [186], [187]. In particular, oleaginous yeasts such as R. toruloides [188] and Y. lipolytica [6], [189], [190], [191], which accumulate large amounts of lipids and have established genetic engineering procedures, might be promising host organism candidates. As a future prospect, the production of desired pheromone chemicals might also be achieved through expression of enzymes with engineered functions.

There are multiple avenues potentially leading to enhanced yields of FA biosynthetic enzymes: (i) genetic modification of the ER retention signal [24], (ii) optimization of the expressed gene codon usage for the host organism [192], (iii) promoter strength and introduction of regulatory sequences such as the eukaryotic Kozak consensus sequence [193], (iv) moving from transient plant transformation and yeast expression plasmids toward stable plant transformants [24] and genome-integrated sequences in yeast [172], and (v) extensive metabolic engineering of the host organisms [7]. In addition to increasing the yield of specific FA derivatives, attention must also be paid to biosynthesis of side products such as undesired FA isomers, which could act as a repellent or inhibitor and thus compromise the pest management strategy.

This review has focused on a relatively minor part of insect biosynthetic capabilities. In a broader context, we envision that insect bioprospecting, the search among insect organisms for commercially valuable resources [194], eventually will exploit their genetic and biosynthetic diversity.

Acknowledgments

This work was supported by the Czech Science Foundation (grant no. 15-06569S) and by the Ministry of Education of the Czech Republic (project no. LD15102 and LO1302).

References

  • 1.

    Rüsch gen. Klaas M, Warwel S. Complete and partial epoxidation of plant oils by lipase-catalyzed perhydrolysis. Ind Crops Prod 1999;9:125–32. CrossrefGoogle Scholar

  • 2.

    Metzger JO, Bornscheuer U. Lipids as renewable resources: current state of chemical and biotechnological conversion and diversification. Appl Microbiol Biotechnol 2006;71:13–22. CrossrefPubMedGoogle Scholar

  • 3.

    Monick JA. Fatty alcohols. J Am Oil Chem Soc 1979;56:853A–60A. CrossrefGoogle Scholar

  • 4.

    Huang Y-S, Pereira SL, Leonard AE. Enzymes for transgenic biosynthesis of long-chain polyunsaturated fatty acids. Biochimie 2004;86:793–8. CrossrefPubMedGoogle Scholar

  • 5.

    Singh SP, Zhou X-R, Liu Q, Stymne S, Green AG. Metabolic engineering of new fatty acids in plants. Curr Opin Plant Biol 2005;8:197–203. PubMedCrossrefGoogle Scholar

  • 6.

    Ledesma-Amaro R, Nicaud J-M. Yarrowia lipolytica as a biotechnological chassis to produce usual and unusual fatty acids. Prog Lipid Res 2016;61:40–50. PubMedCrossrefGoogle Scholar

  • 7.

    Fillet S, Adrio JL. Microbial production of fatty alcohols. World J Microbiol Biotechnol 2016;32:152. CrossrefPubMedGoogle Scholar

  • 8.

    Morgan DE. Biosynthesis in insects. Cambridge: RSC Publishing, 2010. Google Scholar

  • 9.

    El-Sayed AM. The pherobase: database of pheromones and semiochemicals. http://www.pherobase.com, 2016. 

  • 10.

    Liénard MA, Strandh M, Hedenström E, Johansson T, Löfstedt C. Key biosynthetic gene subfamily recruited for pheromone production prior to the extensive radiation of Lepidoptera. BMC Evol Biol 2008;8:270. CrossrefPubMedGoogle Scholar

  • 11.

    Smadja C, Butlin RK. On the scent of speciation: the chemosensory system and its role in premating isolation. Heredity (Edinb) 2009;102:77–97. CrossrefPubMedGoogle Scholar

  • 12.

    Symonds MRE, Elgar MA. The evolution of pheromone diversity. Trends Ecol Evol 2008;23:220–8. CrossrefPubMedGoogle Scholar

  • 13.

    Antony B, Soffan A, Jakše J, Alfaifi S, Sutanto KD, Aldosari SA, et al. Genes involved in sex pheromone biosynthesis of Ephestia cautella, an important food storage pest, are determined by transcriptome sequencing. BMC Genomics 2015;16:532. PubMedCrossrefGoogle Scholar

  • 14.

    Buček A, Brabcová J, Vogel H, Prchalová D, Kindl J, Valterová I, et al. Exploring complex pheromone biosynthetic processes in the bumblebee male labial gland by RNA sequencing. Insect Mol Biol 2016;25:295–314. CrossrefPubMedGoogle Scholar

  • 15.

    Jung CR, Kim Y. Comparative transcriptome analysis of sex pheromone glands of two sympatric lepidopteran congener species. Genomics 2014;103:308–15. PubMedCrossrefGoogle Scholar

  • 16.

    Gu S-H, Wu K-M, Guo Y-Y, Pickett JA, Field LM, Zhou J-J, et al. Identification of genes expressed in the sex pheromone gland of the black cutworm Agrotis ipsilon with putative roles in sex pheromone biosynthesis and transport. BMC Genomics 2013;14:636. CrossrefPubMedGoogle Scholar

  • 17.

    Ding B-J, Löfstedt C. Analysis of the Agrotis segetum pheromone gland transcriptome in the light of sex pheromone biosynthesis. BMC Genomics 2015;16:711. PubMedCrossrefGoogle Scholar

  • 18.

    Groot AT, Dekker T, Heckel DG. The genetic basis of pheromone evolution in moths. Annu Rev Entomol 2016;61:99–117. PubMedCrossrefGoogle Scholar

  • 19.

    Vogel H, Heidel AJ, Heckel DG, Groot AT. Transcriptome analysis of the sex pheromone gland of the noctuid moth Heliothis virescens. BMC Genomics 2010;11:29. PubMedCrossrefGoogle Scholar

  • 20.

    Los DA, Murata N. Membrane fluidity and its roles in the perception of environmental signals. Biochim Biophys Acta 2004;1666:142–57. CrossrefPubMedGoogle Scholar

  • 21.

    Zalucki MP, Shabbir A, Silva R, Adamson D, Shu-Sheng L, Furlong MJ. Estimating the economic cost of one of the world’s major insect pests, Plutella xylostella (Lepidoptera: Plutellidae): just how long is a piece of string? J Econ Entomol 2012;105:1115–29. CrossrefGoogle Scholar

  • 22.

    Shorey HH, Gaston LK, Saario CA. Sex pheromones of noctuid moths. XIV. Feasibility of behavioral control by disrupting pheromone communication in cabbage loopers. J Econ Entomol 1967;60:1541–5. CrossrefGoogle Scholar

  • 23.

    Hagström ÅK, Wang H-L, Liénard MA, Lassance J-M, Johansson T, Löfstedt C. A moth pheromone brewery: production of (Z)-11-hexadecenol by heterologous co-expression of two biosynthetic genes from a noctuid moth in a yeast cell factory. Microb Cell Fact 2013;12:125. CrossrefGoogle Scholar

  • 24.

    Ding B-J, Hofvander P, Wang H-L, Durrett TP, Stymne S, Löfstedt C. A plant factory for moth pheromone production. Nat Commun 2014;5:3353. PubMedGoogle Scholar

  • 25.

    Nesnerova P, Sebek P, Macek T, Svatos A. First semi-synthetic preparation of sex pheromones. Green Chem 2004;6:305. CrossrefGoogle Scholar

  • 26.

    Horne I, Gibb N, Damcevski K, Glover K, Haritos VS. Two conserved Z9-octadecanoic acid desaturases in the red flour beetle, Tribolium castaneum. Gene 2010;468:41–7. CrossrefPubMedGoogle Scholar

  • 27.

    Dallerac R, Labeur C, Jallon JM, Knipple DC, Roelofs WL, Wicker-Thomas C. A delta 9 desaturase gene with a different substrate specificity is responsible for the cuticular diene hydrocarbon polymorphism in Drosophila melanogaster. Proc Natl Acad Sci USA 2000;97:9449–54. CrossrefGoogle Scholar

  • 28.

    Eigenheer AL, Young S, Blomquist GJ, Borgeson CE, Tillman JA, Tittiger C. Isolation and molecular characterization of Musca domestica delta-9 desaturase sequences. Insect Mol Biol 2002;11:533–42. CrossrefPubMedGoogle Scholar

  • 29.

    Jaspers MHJ, Pflanz R, Riedel D, Kawelke S, Feussner I, Schuh R. The fatty acyl-CoA reductase waterproof mediates airway clearance in Drosophila. Dev Biol 2014;385:23–31. CrossrefPubMedGoogle Scholar

  • 30.

    Riddervold MH, Tittiger C, Blomquist GJ, Borgeson CE. Biochemical and molecular characterizaton of house cricket (Acheta domesticus, Orthoptera: Gryllidae) Delta9 desaturase. Insect Biochem Mol Biol 2002;32:1731–40. CrossrefPubMedGoogle Scholar

  • 31.

    Zhou X-R, Horne I, Damcevski K, Haritos V, Green A, Singh S. Isolation and functional characterization of two independently-evolved fatty acid Delta12-desaturase genes from insects. Insect Mol Biol 2008;17:667–76. CrossrefPubMedGoogle Scholar

  • 32.

    Matoušková P, Luxová A, Matoušková J, Jiroš P, Svatoš A, Valterová I, et al. A delta9 desaturase from Bombus lucorum males: investigation of the biosynthetic pathway of marking pheromones. Chembiochem 2008;9:2534–41. CrossrefPubMedGoogle Scholar

  • 33.

    Buček A, Vogel H, Matoušková P, Prchalová D, Záček P, Vrkoslav V, et al. The role of desaturases in the biosynthesis of marking pheromones in bumblebee males. Insect Biochem Mol Biol 2013;43:724–31. PubMedCrossrefGoogle Scholar

  • 34.

    Teerawanichpan P, Robertson AJ, Qiu X. A fatty acyl-CoA reductase highly expressed in the head of honey bee (Apis mellifera) involves biosynthesis of a wide range of aliphatic fatty alcohols. Insect Biochem Mol Biol 2010;40:641–9. PubMedCrossrefGoogle Scholar

  • 35.

    Sperling P, Ternes P, Zank TK, Heinz E. The evolution of desaturases. Prostaglandins Leukot Essent Fatty Acids 2003;68:73–95. PubMedCrossrefGoogle Scholar

  • 36.

    Knipple DC, Rosenfield C-L, Nielsen R, You KM, Jeong SE. Evolution of the integral membrane desaturase gene family in moths and flies. Genetics 2002;162:1737–52. PubMedGoogle Scholar

  • 37.

    Eirín-López JM, Rebordinos L, Rooney AP, Rozas J. The birth-and-death evolution of multigene families revisited. Genome Dyn 2012;7:170–96. PubMedCrossrefGoogle Scholar

  • 38.

    Shanklin J, Cahoon EB. Desaturation and related modifications of fatty acids. Annu Rev Plant Physiol Plant Mol Biol 1998;49:611–41. CrossrefPubMedGoogle Scholar

  • 39.

    Shanklin J, Somerville C. Stearoyl-acyl-carrier-protein desaturase from higher plants is structurally unrelated to the animal and fungal homologs. Proc Natl Acad Sci USA 1991;88:2510–4. CrossrefGoogle Scholar

  • 40.

    Aguilar PS, Cronan JE, de Mendoza D. A Bacillus subtilis gene induced by cold shock encodes a membrane phospholipid desaturase. J Bacteriol 1998;180:2194–200. PubMedGoogle Scholar

  • 41.

    Diaz AR, Mansilla MC, Vila AJ, de Mendoza D. Membrane topology of the acyl-lipid desaturase from Bacillus subtilis. J Biol Chem 2002;277:48099–106. PubMedCrossrefGoogle Scholar

  • 42.

    Mustardy L, Los DA, Gombos Z, Murata N. Immunocytochemical localization of acyl-lipid desaturases in cyanobacterial cells: evidence that both thylakoid membranes and cytoplasmic membranes are sites of lipid desaturation. Proc Natl Acad Sci USA 1996;93:10524–7. CrossrefGoogle Scholar

  • 43.

    Sayanova O, Shewry PR, Napier JA. Histidine-41 of the cytochrome b(5) domain of the borage Delta(6) fatty acid desaturase is essential for enzyme activity. Plant Physiol 1999;121:641–6. CrossrefPubMedGoogle Scholar

  • 44.

    Mitchell AG, Martin CE. A novel cytochrome b5-like domain is linked to the carboxyl terminus of the Saccharomyces cerevisiae delta-9 fatty acid desaturase. J Biol Chem 1995;270:29766–72. PubMedCrossrefGoogle Scholar

  • 45.

    Uemura H. Synthesis and production of unsaturated and polyunsaturated fatty acids in yeast: current state and perspectives. Appl Microbiol Biotechnol 2012;95:1–12. PubMedCrossrefGoogle Scholar

  • 46.

    Certik M, Shimizu S. Biosynthesis and regulation of microbial polyunsaturated fatty acid production. J Biosci Bioeng 1999;87:1–14. CrossrefPubMedGoogle Scholar

  • 47.

    Zhang Z, Dales NA, Winther MD. Opportunities and challenges in developing stearoyl-coenzyme A desaturase-1 inhibitors as novel therapeutics for human disease. J Med Chem 2014;57:5039–56. CrossrefPubMedGoogle Scholar

  • 48.

    Krishnamurthy S, Plaine A, Albert J, Prasad T, Prasad R, Ernst JF. Dosage-dependent functions of fatty acid desaturase Ole1p in growth and morphogenesis of Candida albicans. Microbiology 2004;150:1991–2003. CrossrefPubMedGoogle Scholar

  • 49.

    Nguyen LN, Gacser A, Nosanchuk JD. The stearoyl-coenzyme A desaturase 1 is essential for virulence and membrane stress in Candida parapsilosis through unsaturated fatty acid production. Infect Immun 2011;79:136–45. PubMedCrossrefGoogle Scholar

  • 50.

    Murayama SY, Negishi Y, Umeyama T, Kaneko A, Oura T, Niimi M, et al. Construction and functional analysis of fatty acid desaturase gene disruptants in Candida albicans. Microbiology 2006;152:1551–8. PubMedCrossrefGoogle Scholar

  • 51.

    Alloatti A, Gupta S, Gualdrón-López M, Igoillo-Esteve M, Nguewa PA, Deumer G, et al. Genetic and chemical evaluation of Trypanosoma brucei oleate desaturase as a candidate drug target. PLoS One 2010;5:e14239. CrossrefPubMedGoogle Scholar

  • 52.

    Alloatti A, Testero SA, Uttaro AD. Chemical evaluation of fatty acid desaturases as drug targets in Trypanosoma cruzi. Int J Parasitol 2009;39:985–93. PubMedCrossrefGoogle Scholar

  • 53.

    Shanklin J. Exploring the possibilities presented by protein engineering. Curr Opin Plant Biol 2000;3:243–8. PubMedCrossrefGoogle Scholar

  • 54.

    Tocher DR, Leaver MJ, Hodgson PA. Recent advances in the biochemistry and molecular biology of fatty acyl desaturases. Prog Lipid Res 1998;37:73–117. CrossrefPubMedGoogle Scholar

  • 55.

    Broadwater JA, Whittle E, Shanklin J. Desaturation and hydroxylation. Residues 148 and 324 of Arabidopsis FAD2, in addition to substrate chain length, exert a major influence in partitioning of catalytic specificity. J Biol Chem 2002;277:15613–20. CrossrefGoogle Scholar

  • 56.

    Buist PH. Fatty acid desaturases: selecting the dehydrogenation channel. Nat Prod Rep 2004;21:249–62. CrossrefPubMedGoogle Scholar

  • 57.

    Reed DW, Savile CK, Qiu X, Buist PH, Covello PS. Mechanism of 1,4-dehydrogenation catalyzed by a fatty acid (1,4)-desaturase of Calendula officinalis. Eur J Biochem 2002;269:5024–9. CrossrefPubMedGoogle Scholar

  • 58.

    Rodríguez S, Camps F, Fabriàs G. Synthesis of gem-dideuterated tetradecanoic acids and their use in investigating the enzymatic transformation of (Z)-11-tetradecenoic acid into (E,E)-10,12-tetradecadienoic acid. J Org Chem 2001;66:8052–8. CrossrefGoogle Scholar

  • 59.

    Meesapyodsuk D, Reed DW, Covello PS, Qiu X. Primary structure, regioselectivity, and evolution of the membrane-bound fatty acid desaturases of Claviceps purpurea. J Biol Chem 2007;282:20191–9. PubMedCrossrefGoogle Scholar

  • 60.

    Meesapyodsuk D, Reed DW, Savile CK, Buist PH, Ambrose SJ, Covello PS. Characterization of the regiochemistry and cryptoregiochemistry of a Caenorhabditis elegans fatty acid desaturase (FAT-1) expressed in Saccharomyces cerevisiae. Biochemistry 2000;39:11948–54. PubMedCrossrefGoogle Scholar

  • 61.

    Spychalla JP, Kinney AJ, Browse J. Identification of an animal omega-3 fatty acid desaturase by heterologous expression in Arabidopsis. Proc Natl Acad Sci USA 1997;94:1142–7. CrossrefGoogle Scholar

  • 62.

    Pugh EL, Kates M. Direct desaturation of eicosatrienoyl lecithin to arachidonoyl lecithin by rat liver microsomes. J Biol Chem 1977;252:68–73. PubMedGoogle Scholar

  • 63.

    Knipple DC, Rosenfield CL, Miller SJ, Liu W, Tang J, Ma PW, et al. Cloning and functional expression of a cDNA encoding a pheromone gland-specific acyl-CoA Delta11-desaturase of the cabbage looper moth, Trichoplusia ni. Proc Natl Acad Sci USA 1998;95:15287–92. CrossrefGoogle Scholar

  • 64.

    Wang H-L, Liénard MA, Zhao C-H, Wang C-Z, Löfstedt C. Neofunctionalization in an ancestral insect desaturase lineage led to rare Δ6 pheromone signals in the Chinese tussah silkworm. Insect Biochem Mol Biol 2010;40:742–51. CrossrefGoogle Scholar

  • 65.

    Liénard MA, Lassance J, Wang H, Zhao C, Piskur J, Johansson T, et al. Elucidation of the sex-pheromone biosynthesis producing 5,7-dodecadienes in Dendrolimus punctatus (Lepidoptera: Lasiocampidae) reveals Delta 11- and Delta 9-desaturases with unusual catalytic properties. Insect Biochem Mol Biol 2010;40:440–52. PubMedCrossrefGoogle Scholar

  • 66.

    Liu W, Jiao H, Murray NC, O’Connor M, Roelofs WL. Gene characterized for membrane desaturase that produces (E)-11 isomers of mono- and diunsaturated fatty acids. Proc Natl Acad Sci USA 2002;99:620–4. CrossrefGoogle Scholar

  • 67.

    Liu W, Rooney AP, Xue B, Roelofs WL. Desaturases from the spotted fireworm moth (Choristoneura parallela) shed light on the evolutionary origins of novel moth sex pheromone desaturases. Gene 2004;342:303–11. PubMedCrossrefGoogle Scholar

  • 68.

    Fujii T, Ito K, Tatematsu M, Shimada T, Katsuma S, Ishikawa Y. Sex pheromone desaturase functioning in a primitive Ostrinia moth is cryptically conserved in congeners’ genomes. Proc Natl Acad Sci USA 2011;108:7102–6. CrossrefGoogle Scholar

  • 69.

    Hao G, Liu W, O’Connor M, Roelofs WL. Acyl-CoA Z9-and Z10-desaturase genes from a New Zealand leafroller moth species, Planotortrix octo. Insect Biochem Mol Biol 2002;32:961–6. CrossrefPubMedGoogle Scholar

  • 70.

    Roelofs WL, Liu W, Hao G, Jiao H, Rooney AP, Linn CE. Evolution of moth sex pheromones via ancestral genes. Proc Natl Acad Sci USA 2002;99:13621–6. CrossrefGoogle Scholar

  • 71.

    Liu W, Jiao H, O’Connor M, Roelofs WL. Moth desaturase characterized that produces both Z and E isomers of delta 11-tetradecenoic acids. Insect Biochem Mol Biol 2002;32:1489–95. CrossrefGoogle Scholar

  • 72.

    Hagström ÅK, Albre J, Tooman LK, Thirmawithana AH, Corcoran J, Löfstedt C, et al. A novel fatty acyl desaturase from the pheromone glands of Ctenopseustis obliquana and C. herana with specific Z5-desaturase activity on myristic acid. J Chem Ecol 2014;40:63–70. PubMedCrossrefGoogle Scholar

  • 73.

    Hao G, O’Connor M, Liu W, Roelofs WL. Characterization of Z/E11- and Z9-desaturases from the obliquebanded leafroller moth, Choristoneura rosaceana. J Insect Sci 2002;2:26. PubMedGoogle Scholar

  • 74.

    Jeong SE, Eun S, Rosenfield CL, Marsella-herrick P, You KM, Knipple DC. Multiple acyl-CoA desaturase-encoding transcripts in pheromone glands of Helicoverpa assulta, the oriental tobacco budworm. Insect Biochem Mol Biol 2003;33:609–22. CrossrefPubMedGoogle Scholar

  • 75.

    Park HY, Kim MS, Paek A, Jeong SE, Knipple DC. An abundant acyl-CoA (Delta9) desaturase transcript in pheromone glands of the cabbage moth, Mamestra brassicae, encodes a catalytically inactive protein. Insect Biochem Mol Biol 2008;38:581–95. CrossrefPubMedGoogle Scholar

  • 76.

    Rodríguez S, Hao G, Liu W, Piña B, Rooney AP, Camps F, et al. Expression and evolution of delta9 and delta11 desaturase genes in the moth Spodoptera littoralis. Insect Biochem Mol Biol 2004;34:1315–28. PubMedCrossrefGoogle Scholar

  • 77.

    Liénard MA, Löfstedt C. Functional flexibility as a prelude to signal diversity?: role of a fatty acyl reductase in moth pheromone evolution. Commun Integr Biol 2010;3:586–8. PubMedCrossrefGoogle Scholar

  • 78.

    Albre J, Liénard MA, Sirey TM, Schmidt S, Tooman LK, Carraher C, et al. Sex pheromone evolution is associated with differential regulation of the same desaturase gene in two genera of leafroller moths. PLoS Genet 2012;8:e1002489. CrossrefPubMedGoogle Scholar

  • 79.

    Moto K, Suzuki MG, Hull JJ, Kurata R, Takahashi S, Yamamoto M, et al. Involvement of a bifunctional fatty-acyl desaturase in the biosynthesis of the silkmoth, Bombyx mori, sex pheromone. Proc Natl Acad Sci USA 2004;101:8631–6. CrossrefGoogle Scholar

  • 80.

    Matoušková P, Pichová I, Svatoš A. Functional characterization of a desaturase from the tobacco hornworm moth (Manduca sexta) with bifunctional Z11- and 10,12-desaturase activity. Insect Biochem Mol Biol 2007;37:601–10. CrossrefPubMedGoogle Scholar

  • 81.

    Serra M, Piña B, Abad JL, Camps F, Fabriàs G. A multifunctional desaturase involved in the biosynthesis of the processionary moth sex pheromone. Proc Natl Acad Sci USA 2007;104: 16444–9. CrossrefGoogle Scholar

  • 82.

    Buček A, Matoušková P, Vogel H, Šebesta P, Jahn U, Weißflog J, et al. Evolution of moth sex pheromone composition by a single amino acid substitution in a fatty acid desaturase. Proc Natl Acad Sci USA 2015;112:12586–91. CrossrefGoogle Scholar

  • 83.

    Ding B, Liénard MA, Wang H, Zhao C, Löfstedt C. Terminal fatty-acyl-CoA desaturase involved in sex pheromone biosynthesis in the winter moth (Operophtera brumata). Insect Biochem Mol Biol 2011;41:715–22. PubMedCrossrefGoogle Scholar

  • 84.

    Lee M, Lenman M, Banaś A, Bafor M, Singh S, Schweizer M, et al. Identification of non-heme diiron proteins that catalyze triple bond and epoxy group formation. Science 1998;280:915–8. CrossrefPubMedGoogle Scholar

  • 85.

    Serra M, Gauthier LT, Fabrias G, Buist PH. Delta11 desaturases of Trichoplusia ni and Spodoptera littoralis exhibit dual catalytic behaviour. Insect Biochem Mol Biol 2006;36:822–5. CrossrefPubMedGoogle Scholar

  • 86.

    Liénard MA, Wang H-L, Lassance J-M, Löfstedt C. Sex pheromone biosynthetic pathways are conserved between moths and the butterfly Bicyclus anynana. Nat Commun 2014;5:3957. doi: 10.1038/ncomms4957. PubMedGoogle Scholar

  • 87.

    Rosenfield CL, You KM, Marsella-Herrick P, Roelofs WL, Knipple DC. Structural and functional conservation and divergence among acyl-CoA desaturases of two noctuid species, the corn earworm, Helicoverpa zea, and the cabbage looper, Trichoplusia ni. Insect Biochem Mol Biol 2001;31:949–64. PubMedGoogle Scholar

  • 88.

    Liu W, Ma PWK, Marsella-Herrick P, Rosenfield CL, Knipple DC, Roelofs T. Cloning and functional expression of a cDNA encoding a metabolic acyl-CoA Delta 9-desaturase of the cabbage looper moth, Trichoplusia ni. Insect Biochem Mol Biol 1999;29:435–43. CrossrefPubMedGoogle Scholar

  • 89.

    Liénard MA, Hagström AK, Lassance J-M, Löfstedt C. Evolution of multicomponent pheromone signals in small ermine moths involves a single fatty-acyl reductase gene. Proc Natl Acad Sci USA 2010;107:10955–60. CrossrefGoogle Scholar

  • 90.

    Haritos VS, Horne I, Damcevski K, Glover K, Gibb N. Unexpected functional diversity in the fatty acid desaturases of the flour beetle Tribolium castaneum and identification of key residues determining activity. Insect Biochem Mol Biol 2014;51:62–70. PubMedCrossrefGoogle Scholar

  • 91.

    Riendeau D, Meighen E. Enzymatic reduction of fatty acids and acyl-CoAs to long chain aldehydes and alcohols. Experientia 1985;41:707–13. CrossrefGoogle Scholar

  • 92.

    Kolattukudy PE. Reduction of fatty acids to alcohols by cell-free preparations of Euglena gracilis. Biochemistry 1970;9:1095–102. CrossrefPubMedGoogle Scholar

  • 93.

    Vioque J, Kolattukudy PE. Resolution and purification of an aldehyde-generating and an alcohol-generating fatty acyl-CoA reductase from pea leaves (Pisum sativum L.). Arch Biochem Biophys 1997;340:64–72. CrossrefPubMedGoogle Scholar

  • 94.

    Hagström AK, Walther A, Wendland J, Löfstedt C. Subcellular localization of the fatty acyl reductase involved in pheromone biosynthesis in the tobacco budworm, Heliothis virescens (Noctuidae: Lepidoptera). Insect Biochem Mol Biol 2013; 43:510–21. PubMedCrossrefGoogle Scholar

  • 95.

    Moore C, Snyder F. Properties of microsomal acyl coenzyme A reductase in mouse preputial glands. Arch Biochem Biophys 1982;214:489–99. PubMedCrossrefGoogle Scholar

  • 96.

    Aarts MGM, Hodge R, Kalantidis K, Florack D, Wilson ZA, Mulligan BJ, et al. The Arabidopsis MALE STERILITY 2 protein shares similarity with reductases in elongation/condensation complexes. Plant J 1997;12:615–23. CrossrefPubMedGoogle Scholar

  • 97.

    Cheng JB, Russell DW. Mammalian wax biosynthesis. I. Identification of two fatty acyl-Coenzyme A reductases with different substrate specificities and tissue distributions. J Biol Chem 2004;279:37789–97. PubMedCrossrefGoogle Scholar

  • 98.

    Hellenbrand J, Biester E-M, Gruber J, Hamberg M, Frentzen M. Fatty acyl-CoA reductases of birds. BMC Biochem 2011;12:64. CrossrefPubMedGoogle Scholar

  • 99.

    Nagan N, Zoeller RA. Plasmalogens: biosynthesis and functions. Prog Lipid Res 2001;40:199–229. CrossrefPubMedGoogle Scholar

  • 100.

    Metz JG, Pollard MR, Anderson L, Hayes TR, Lassner MW. Purification of a jojoba embryo fatty acyl-coenzyme A reductase and expression of its cDNA in high erucic acid rapeseed. Plant Physiol 2000;122:635–44. CrossrefPubMedGoogle Scholar

  • 101.

    Teerawanichpan P, Qiu X. Molecular and functional analysis of three fatty acyl-CoA reductases with distinct substrate specificities in copepod Calanus finmarchicus. Mar Biotechnol (NY) 2012;14:227–36. CrossrefPubMedGoogle Scholar

  • 102.

    Teerawanichpan P, Qiu X. Fatty acyl-CoA reductase and wax synthase from Euglena gracilis in the biosynthesis of medium-chain wax esters. Lipids 2010;45:263–73. PubMedCrossrefGoogle Scholar

  • 103.

    Khannoon ER, Flachsbarth B, El-Gendy A, Mazik K, Hardege JD, Schulz S. New compounds, sexual differences, and age-related variations in the femoral gland secretions of the lacertid lizard Acanthodactylus boskianus. Biochem Syst Ecol 2011;39:95–101. CrossrefGoogle Scholar

  • 104.

    Wood WF. Volatile components in metatarsal glands of sika deer, Cervus nippon. J Chem Ecol 2003;29:2729–33. CrossrefPubMedGoogle Scholar

  • 105.

    Butenandt von A, Beckmann R, Stamm D, Hecker E. Über den sexuallockstoff des seidenspinners Bombyx mori. Reindarstellung und konstitution. Z Naturforsch B 1959;14:283–4. Google Scholar

  • 106.

    Jirošová A, Sillam-Dussès D, Kyjaková P, Kalinová B, Dolejšová K, Jančařík A, et al. Smells Like Home: Chemically mediated co-habitation of two termite species in a single nest. J Chem Ecol 2016;42:1070–81. CrossrefGoogle Scholar

  • 107.

    Kullenberg B, Bergström G, Ställberg-Stenhagen S. Volatile components of the cephalic marking secretion of male bumble bees. Acta Chem Scand 1970;24:1481–5. PubMedCrossrefGoogle Scholar

  • 108.

    Urbanová K, Valterová I, Hovorka O, Kindl J. Chemotaxonomical characterisation of males Bombus lucorum (Hymenptera: Apidae) collected in Czech Republic. Eur J Entomol 2001;127:111–5. Google Scholar

  • 109.

    Pickett JA, Williams IH, Martin AP. (Z)-11-eicosen-1-ol, an important new pheromonal component from the sting of the honey bee, Apis mellifera L. (Hymenoptera, Apidae.). J Chem Ecol 1982;8:163–75. CrossrefGoogle Scholar

  • 110.

    Tillman JA, Seybold SJ, Jurenka RA, Blomquist GJ. Insect pheromones – an overview of biosynthesis and endocrine regulation. Insect Biochem Mol Biol 1999;29:481–514. CrossrefPubMedGoogle Scholar

  • 111.

    Antony B, Ding B-J, Moto K, Aldosari SA, Aldawood AS. Two fatty acyl reductases involved in moth pheromone biosynthesis. Sci Rep 2016;6:29927. CrossrefPubMedGoogle Scholar

  • 112.

    Rowland O, Zheng H, Hepworth SR, Lam P, Jetter R, Kunst L. CER4 encodes an alcohol-forming fatty acyl-coenzyme A reductase involved in cuticular wax production in Arabidopsis. Plant Physiol 2006;142:866–77. PubMedCrossrefGoogle Scholar

  • 113.

    Doan TTP, Carlsson AS, Hamberg M, Bülow L, Stymne S, Olsson P. Functional expression of five Arabidopsis fatty acyl-CoA reductase genes in Escherichia coli. J Plant Physiol 2009;166:787–96. PubMedCrossrefGoogle Scholar

  • 114.

    Wang A, Xia Q, Xie W, Dumonceaux T, Zou J, Datla R, et al. Male gametophyte development in bread wheat (Triticum aestivum L.): molecular, cellular, and biochemical analyses of a sporophytic contribution to pollen wall ontogeny. Plant J 2002;30:613–23. CrossrefPubMedGoogle Scholar

  • 115.

    Miklaszewska M, Banaś A. Biochemical characterization and substrate specificity of jojoba fatty acyl-CoA reductase and jojoba wax synthase. Plant Sci 2016;249:84–92. CrossrefPubMedGoogle Scholar

  • 116.

    Willis RM, Wahlen BD, Seefeldt LC, Barney BM. Characterization of a fatty acyl-CoA reductase from Marinobacter aquaeolei VT8: a bacterial enzyme catalyzing the reduction of fatty acyl-CoA to fatty alcohol. Biochemistry 2011;50:10550–8. PubMedCrossrefGoogle Scholar

  • 117.

    Hofvander P, Doan TTP, Hamberg M. A prokaryotic acyl-CoA reductase performing reduction of fatty acyl-CoA to fatty alcohol. FEBS Lett 2011;585:3538–43. CrossrefPubMedGoogle Scholar

  • 118.

    Moto K, Yoshiga T, Yamamoto M, Takahashi S, Okano K, Ando T, et al. Pheromone gland-specific fatty-acyl reductase of the silkmoth, Bombyx mori. Proc Natl Acad Sci USA 2003;100:9156–61. CrossrefGoogle Scholar

  • 119.

    Lassance J-M, Groot AT, Liénard MA, Antony B, Borgwardt C, Andersson F, et al. Allelic variation in a fatty-acyl reductase gene causes divergence in moth sex pheromones. Nature 2010;466:486–9. CrossrefGoogle Scholar

  • 120.

    Lassance J-M, Liénard MA, Antony B, Qian S, Fujii T, Tabata J, et al. Functional consequences of sequence variation in the pheromone biosynthetic gene pgFAR for Ostrinia moths. Proc Natl Acad Sci USA 2013;110:3967–72. CrossrefGoogle Scholar

  • 121.

    Antony B, Fujii T, Moto K, Matsumoto S, Fukuzawa M, Nakano R, et al. Pheromone-gland-specific fatty-acyl reductase in the adzuki bean borer, Ostrinia scapulalis (Lepidoptera: Crambidae). Insect Biochem Mol Biol 2009;39:90–5. PubMedCrossrefGoogle Scholar

  • 122.

    Carot-Sans G, Muñoz L, Piulachs MD, Guerrero A, Rosell G. Identification and characterization of a fatty acyl reductase from a Spodoptera littoralis female gland involved in pheromone biosynthesis. Insect Mol Biol 2015;24:82–92. CrossrefPubMedGoogle Scholar

  • 123.

    Hagström AK, Liénard MA, Groot AT, Hedenström E, Löfstedt C. Semi-selective fatty acyl reductases from four heliothine moths influence the specific pheromone composition. PLoS One 2012;7:e37230. CrossrefPubMedGoogle Scholar

  • 124.

    Zhang Y-N, Zhu X-Y, Fang L-P, He P, Wang Z-Q, Chen G, et al. Identification and expression profiles of sex pheromone biosynthesis and transport related genes in Spodoptera litura. PLoS One 2015;10:e0140019. PubMedCrossrefGoogle Scholar

  • 125.

    Du M, Zhang S, Zhu B, Yin X, An S. Identification of a diacylglycerol acyltransferase 2 gene involved in pheromone biosynthesis activating neuropeptide stimulated pheromone production in Bombyx mori. J Insect Physiol 2012;58:699–703. CrossrefPubMedGoogle Scholar

  • 126.

    Du M, Liu X, Liu X, Yin X, Han S, Song Q, et al. Glycerol-3-phosphate O-acyltransferase is required for PBAN-induced sex pheromone biosynthesis in Bombyx mori. Sci Rep 2015;5:8110. CrossrefPubMedGoogle Scholar

  • 127.

    Du M, Yin X, Zhang S, Zhu B, Song Q, An S. Identification of lipases involved in PBAN stimulated pheromone production in Bombyx mori using the DGE and RNAi approaches. PLoS One 2012;7:e31045. CrossrefPubMedGoogle Scholar

  • 128.

    Zhang SD, Li X, Bin Z, Du MF, Yin XM, An SH. Molecular identification of a pancreatic lipase-like gene involved in sex pheromone biosynthesis of Bombyx mori. Insect Sci 2014;21:459–68. PubMedCrossrefGoogle Scholar

  • 129.

    Brabcová J, Prchalová D, Demianová Z, Bučánková A, Vogel H, Valterová I, et al. Characterization of neutral lipase BT-1 isolated from the labial gland of Bombus terrestris males. PLoS One 2013;8:e80066. CrossrefPubMedGoogle Scholar

  • 130.

    Castillo C, Chen H, Graves C, Maisonnasse A, Le Conte Y, Plettner E. Biosynthesis of ethyl oleate, a primer pheromone, in the honey bee (Apis mellifera L.). Insect Biochem Mol Biol 2012;42:404–16. CrossrefPubMedGoogle Scholar

  • 131.

    Smith GM, Rothwell K, Wood SL, Yeaman SJ, Bownes M. Specificity and localization of lipolytic activity in adult Drosophila melanogaster. Biochem J 1994;304:775–9. CrossrefPubMedGoogle Scholar

  • 132.

    Arreguín-Espinosa R, Arreguín B, González C. Purification and properties of a lipase from Cephaloleia presignis (Coleoptera, chrysomelidae). Biotechnol Appl Biochem 2000;31:239–44. PubMedCrossrefGoogle Scholar

  • 133.

    Arrese EL, Wells MA. Purification and properties of a phosphorylatable triacylglycerol lipase from the fat body of an insect, Manduca sexta. J Lipid Res 1994;35:1652–60. PubMedGoogle Scholar

  • 134.

    Qiu Y, Tittiger C, Wicker-Thomas C, Le Goff G, Young S, Wajnberg E, et al. An insect-specific P450 oxidative decarbonylase for cuticular hydrocarbon biosynthesis. Proc Natl Acad Sci USA 2012;109:14858–63. CrossrefGoogle Scholar

  • 135.

    Tillman-Wall JA, Vanderwel D, Kuenzli ME, Reitz RC, Blomquist GJ. Regulation of sex pheromone biosynthesis in the housefly, Musca domestica: relative contribution of the elongation and reductive steps. Arch Biochem Biophys 1992;299:92–9. CrossrefPubMedGoogle Scholar

  • 136.

    Wicker-Thomas C, Chertemps T. Molecular biology and genetics of hydrocarbon production. In: Insect Hydrocarbons: biology, Biochemistry, and Chemical Ecology. Cambridge, UK: Cambridge University Press, 2010:53–74. Google Scholar

  • 137.

    Plettner E, Slessor KN, Winston ML, Oliver JE. Caste-selective pheromone biosynthesis in honeybees. Science (80-) 1996;271:1851–3. CrossrefGoogle Scholar

  • 138.

    Rosell G, Hospital S, Camps F, Guerrero A. Inhibition of a chain shortening step in the biosynthesis of the sex pheromone of the Egyptian armyworm Spodoptera littoralis. Insect Biochem Mol Biol 1992;22:679–85. CrossrefGoogle Scholar

  • 139.

    Jurenka RA, Haynes KF, Adlof RO, Bengtsson M, Roelofs WL. Sex pheromone component ratio in the cabbage looper moth altered by a mutation affecting the fatty acid chain-shortening reactions in the pheromone biosynthetic pathway. Insect Biochem Mol Biol 1994;24:373–81. CrossrefGoogle Scholar

  • 140.

    Bjostad LB, Roelofs WL. Sex pheromone biosynthesis in Trichoplusia ni: key steps involve Delta-11 desaturation and chain-shortening. Science 1983;220:1387–9. CrossrefPubMedGoogle Scholar

  • 141.

    Jurenka RA, Subchev M, Abad J-L, Choi M-Y, Fabrias G. Sex pheromone biosynthetic pathway for disparlure in the gypsy moth, Lymantria dispar. Proc Natl Acad Sci USA 2003;100:809–14. CrossrefGoogle Scholar

  • 142.

    Millar JG. Polyene hydrocarbons, epoxides, and related compounds as components of lepidopteran pheromone blends. In: Insect Hydrocarbons: Biology, Biochemistry, and Chemical Ecology. Cambridge, UK: Cambridge University Press, 2010:390–447. Google Scholar

  • 143.

    Jurenka RA, Roelofs WL. Characterization of the acetyltransferase used in pheromone biosynthesis in moths: Specificity for the Z isomer in tortricidae. Insect Biochem 1989;19:639–44. CrossrefGoogle Scholar

  • 144.

    Zhao C-H, Lu F, Bengtsson M, Löfstedt C. Substrate specificity of acetyltransferase and reductase enzyme systems used in pheromone biosynthesis by Asian corn borer, Ostrinia furnacalis. J Chem Ecol 1995;21:1495–510. CrossrefPubMedGoogle Scholar

  • 145.

    Luxová A, Svatoš A. Substrate specificity of membrane-bound alcohol oxidase from the tobacco hornworm moth (Manduca sexta) female pheromone glands. J Mol Catal B Enzym 2006;38:37–42. CrossrefGoogle Scholar

  • 146.

    Fang N, Teal PEA, Tumlinson JH. Characterization of oxidase (s) associated with the sex pheromone gland in Manduca sexta (L.) females. Insect Biochem Physiol 1995;257:243–57. Google Scholar

  • 147.

    Teal PE, Tumlinson JH. Terminal steps in pheromone biosynthesis by Heliothis virescens and H. zea. J Chem Ecol 1986;12:353–66. CrossrefGoogle Scholar

  • 148.

    Tsfadia O, Azrielli A, Falach L, Zada A, Roelofs W, Rafaeli A. Pheromone biosynthetic pathways: PBAN-regulated rate-limiting steps and differential expression of desaturase genes in moth species. Insect Biochem Mol Biol 2008;38:552–67. CrossrefPubMedGoogle Scholar

  • 149.

    Alabaster A, Isoe J, Zhou G, Lee A, Murphy A, Day WA, et al. Deficiencies in acetyl-CoA carboxylase and fatty acid synthase 1 differentially affect eggshell formation and blood meal digestion in Aedes aegypti. Insect Biochem Mol Biol 2011;41:946–55. PubMedCrossrefGoogle Scholar

  • 150.

    Ding B-J, Carraher C, Löfstedt C. Sequence variation determining stereochemistry of a Δ11 desaturase active in moth sex pheromone biosynthesis. Insect Biochem Mol Biol 2016;74:68–75. CrossrefPubMedGoogle Scholar

  • 151.

    Stukey JE, McDonough VM, Martin CE. The OLE1 gene of Saccharomyces cerevisiae encodes the delta 9 fatty acid desaturase and can be functionally replaced by the rat stearoyl-CoA desaturase gene. J Biol Chem 1990;265:20144–9. PubMedGoogle Scholar

  • 152.

    Meesapyodsuk D, Qiu X. Structure determinants for the substrate specificity of acyl-CoA Δ9 desaturases from a marine copepod. ACS Chem Biol 2014;9:922–34. PubMedCrossrefGoogle Scholar

  • 153.

    Libisch B, Michaelson L V, Lewis MJ, Shewry PR, Napier JA. Chimeras of Δ6-fatty acid and Δ8-sphingolipid desaturases. Biochem Biophys Res Commun 2000;279:779–85. PubMedCrossrefGoogle Scholar

  • 154.

    Watanabe K, Ohno M, Taguchi M, Kawamoto S, Ono K, Aki T. Identification of amino acid residues that determine the substrate specificity of mammalian membrane-bound front-end fatty acid desaturases. J Lipid Res 2016;57:89–99. CrossrefPubMedGoogle Scholar

  • 155.

    Hoffmann M, Hornung E, Busch S, Kassner N, Ternes P, Braus GH, et al. A small membrane-peripheral region close to the active center determines regioselectivity of membrane-bound fatty acid desaturases from Aspergillus nidulans. J Biol Chem 2007;282:26666–74. CrossrefPubMedGoogle Scholar

  • 156.

    Hongsthong A, Subudhi S, Sirijuntarat M, Cheevadhanarak S. Mutation study of conserved amino acid residues of Spirulina delta 6-acyl-lipid desaturase showing involvement of histidine 313 in the regioselectivity of the enzyme. Appl Microbiol Biotechnol 2004;66:74–84. CrossrefPubMedGoogle Scholar

  • 157.

    Lim ZL, Senger T, Vrinten P. Four amino acid residues influence the substrate chain-length and regioselectivity of Siganus canaliculatus Δ4 and Δ5/6 desaturases. Lipids 2014;49: 357–67. PubMedCrossrefGoogle Scholar

  • 158.

    Shi H, Chen H, Gu Z, Song Y, Zhang H, Chen W, et al. Molecular mechanism of substrate specificity for delta 6 desaturase from Mortierella alpina and Micromonas pusilla. J Lipid Res 2015;56:2309–21. PubMedCrossrefGoogle Scholar

  • 159.

    Gagné SJ, Reed DW, Gray GR, Covello PS. Structural control of chemoselectivity, stereoselectivity, and substrate specificity in membrane-bound fatty acid acetylenases and desaturases. Biochemistry 2009;48:12298–304. CrossrefPubMedGoogle Scholar

  • 160.

    Rawat R, Yu XH, Sweet M, Shanklin J. Conjugated fatty acid synthesis: Residues 111 and 115 influence product partitioning of Momordica charantia conjugase. J Biol Chem 2012;287:16230–7. CrossrefPubMedGoogle Scholar

  • 161.

    Broun P, Shanklin J, Whittle E, Somerville C. Catalytic plasticity of fatty acid modification enzymes underlying chemical diversity of plant lipids. Science 1998;282:1315–7. PubMedCrossrefGoogle Scholar

  • 162.

    Vanhercke T, Shrestha P, Green AG, Singh SP. Mechanistic and structural insights into the regioselectivity of an acyl-CoA fatty acid desaturase via directed molecular evolution. J Biol Chem 2011;286:12860–9. PubMedCrossrefGoogle Scholar

  • 163.

    Wang H, Klein MG, Zou H, Lane W, Snell G, Levin I, et al. Crystal structure of human stearoyl-coenzyme A desaturase in complex with substrate. Nat Struct Mol Biol 2015;22:581–5. CrossrefPubMedGoogle Scholar

  • 164.

    Bai Y, McCoy JG, Levin EJ, Sobrado P, Rajashankar KR, Fox BG, et al. X-ray structure of a mammalian stearoyl-CoA desaturase. Nature 2015;524:252–6. PubMedCrossrefGoogle Scholar

  • 165.

    Man WC, Miyazaki M, Chu K, Ntambi JM. Membrane topology of mouse stearoyl-CoA desaturase 1. J Biol Chem 2006;281:1251–60. CrossrefPubMedGoogle Scholar

  • 166.

    Shanklin J, Whittle E, Fox BG. Eight histidine residues are catalytically essential in a membrane-associated iron enzyme, stearoyl-CoA desaturase, and are conserved in alkane hydroxylase and xylene monooxygenase. Biochemistry 1994;33:12787–94. CrossrefGoogle Scholar

  • 167.

    Chacón MG, Fournier AE, Tran F, Dittrich-Domergue F, Pulsifer IP, Domergue F, et al. Identification of amino acids conferring chain length substrate specificities on fatty alcohol-forming reductases FAR5 and FAR8 from Arabidopsis thaliana. J Biol Chem 2013;288:30345–55. PubMedCrossrefGoogle Scholar

  • 168.

    Witzgall P, Kirsch P, Cork A. Sex pheromones and their impact on pest management. J Chem Ecol 2010;36:80–100. CrossrefPubMedGoogle Scholar

  • 169.

    Millar JG, McElfresh JS, Romero C, Vila M, Marí-Mena N, Lopez-Vaamonde C. Identification of the sex pheromone of a protected species, the Spanish moon moth Graellsia isabellae. J Chem Ecol 2010;36:923–32. PubMedCrossrefGoogle Scholar

  • 170.

    Larsson MC. Pheromones and other semiochemicals for monitoring rare and endangered species. J Chem Ecol 2016;42:853–68. CrossrefPubMedGoogle Scholar

  • 171.

    Howse PE, Stevens IDR, Jones OT. Synthesis of pheromones. In: Insect pheromones and their use in pest management. Netherlands: Springer Science+Business Media B.V. 1998:226–45. Google Scholar

  • 172.

    Da Silva NA, Srikrishnan S. Introduction and expression of genes for metabolic engineering applications in Saccharomyces cerevisiae. FEMS Yeast Res 2012;12:197–214. PubMedCrossrefGoogle Scholar

  • 173.

    Muñoz L, Dimov N, Carot-Sans G, Bula WP, Guerrero A, Gardeniers HJGE. Mimicking insect communication: release and detection of pheromone, biosynthesized by an alcohol acetyl transferase immobilized in a microreactor. PLoS One 2012;7:e47751. CrossrefGoogle Scholar

  • 174.

    de Miranda AS, Miranda LSM, de Souza ROMA. Lipases: Valuable catalysts for dynamic kinetic resolutions. Biotechnol Adv 2015;33:372–93. CrossrefPubMedGoogle Scholar

  • 175.

    Gao W-W, Zhang F-X, Zhang G-X, Zhou C-H. Key factors affecting the activity and stability of enzymes in ionic liquids and novel applications in biocatalysis. Biochem Eng J 2015;99:67–84. CrossrefGoogle Scholar

  • 176.

    Sharma R, Chisti Y, Banerjee UC. Production, purification, characterization, and applications of lipases. Biotechnol Adv 2001;19:627–62. PubMedCrossrefGoogle Scholar

  • 177.

    Wandrey C, Liese A, Kihumbu D. Industrial biocatalysis: past, present, and future. Org Process Res Dev 2000;4:286–90. CrossrefGoogle Scholar

  • 178.

    Horne I, Haritos VS, Oakeshott JG. Comparative and functional genomics of lipases in holometabolous insects. Insect Biochem Mol Biol 2009;39:547–67. PubMedCrossrefGoogle Scholar

  • 179.

    Mastihuba V, Čepec P, Vlčková S, Farkašová E, Mastihubová M, Bobal P. Enzymatic synthesis of a chiral chalcogran intermediate. Chem Pap 2014;68:745–50. Google Scholar

  • 180.

    Sonnet PE, Baillargeon MW. Kinetic resolution of secondary alcohols with commercial lipases. J Chem Ecol 1987;13:1279–92. CrossrefPubMedGoogle Scholar

  • 181.

    Naoshima Y, Kamezawa M, Tachibana H, Munakata Y, Fujita T, Kihara K, et al. Enzymatic preparation of enantiomerically pure alkan-2- and -3-ols by lipase-catalysed hydrolysis with Pseudomonas cepacia in the presence of organic media. J Chem Soc Perkin Trans 1993;1:557–61. Google Scholar

  • 182.

    Mori K. Chemoenzymatic preparation of enantiopure building blocks of synthetic utility. In: Biocatalysis in the pharmaceutical and biotechnology industries. USA: CRC Press, 2007:564–90. Google Scholar

  • 183.

    Wenning L, Yu T, David F, Nielsen J, Siewers V. Establishing very long-chain fatty alcohol and wax ester biosynthesis in Saccharomyces cerevisiae. Biotechnol Bioeng 2017;114:1025–35. CrossrefPubMedGoogle Scholar

  • 184.

    Runguphan W, Keasling JD. Metabolic engineering of Saccharomyces cerevisiae for production of fatty acid-derived biofuels and chemicals. Metab Eng 2014;21:103–13. PubMedCrossrefGoogle Scholar

  • 185.

    Liu Y, Chen S, Chen J, Zhou J, Wang Y, Yang M, et al. High production of fatty alcohols in Escherichia coli with fatty acid starvation. Microb Cell Fact 2016;15:129. PubMedCrossrefGoogle Scholar

  • 186.

    Rutter CD, Rao CV. Production of 1-decanol by metabolically engineered Yarrowia lipolytica. Metab Eng 2016;38:139–47. CrossrefPubMedGoogle Scholar

  • 187.

    Wang W, Wei H, Knoshaug E, Van Wychen S, Xu Q, Himmel ME, et al. Fatty alcohol production in Lipomyces starkeyi and Yarrowia lipolytica. Biotechnol Biofuels 2016;9:227. CrossrefPubMedGoogle Scholar

  • 188.

    Zhang S, Skerker JM, Rutter CD, Maurer MJ, Arkin AP, Rao C V. Engineering Rhodosporidium toruloides for increased lipid production. Biotechnol Bioeng 2016;113:1056–66. PubMedCrossrefGoogle Scholar

  • 189.

    Chuang L-T, Chen D-C, Nicaud J-M, Madzak C, Chen Y-H, Huang Y-S. Co-expression of heterologous desaturase genes in Yarrowia lipolytica. N Biotechnol 2010;27:277–82. CrossrefPubMedGoogle Scholar

  • 190.

    Wang G, Xiong X, Ghogare R, Wang P, Meng Y, Chen S. Exploring fatty alcohol-producing capability of Yarrowia lipolytica. Biotechnol Biofuels 2016;9:107. PubMedCrossrefGoogle Scholar

  • 191.

    Xu P, Qiao K, Ahn WS, Stephanopoulos G. Engineering Yarrowia lipolytica as a platform for synthesis of drop-in transportation fuels and oleochemicals. Proc Natl Acad Sci USA 2016;113:10848–53. CrossrefGoogle Scholar

  • 192.

    Gustafsson C, Govindarajan S, Minshull J. Codon bias and heterologous protein expression. Trends Biotechnol 2004;22:346–53. PubMedCrossrefGoogle Scholar

  • 193.

    Kozak M. Point mutations define a sequence flanking the AUG initiator codon that modulates translation by eukaryotic ribosomes. Cell 1986;44:283–92. CrossrefPubMedGoogle Scholar

  • 194.

    Srivastava SK. Insect bioprospecting especially in India. In: Topics of Biodiversity and Conservation. Switzerland: Springer International Publishing, 2017:245–67. Google Scholar

About the article

Michal Tupec and Aleš Buček: These authors contributed equally to this work.


Received: 2017-02-28

Revised: 2017-04-19

Accepted: 2017-07-25

Published Online: 2017-07-25

Published in Print: 2017-09-26


Citation Information: Zeitschrift für Naturforschung C, Volume 72, Issue 9-10, Pages 387–403, ISSN (Online) 1865-7125, ISSN (Print) 0939-5075, DOI: https://doi.org/10.1515/znc-2017-0031.

Export Citation

©2017 Walter de Gruyter GmbH, Berlin/Boston.Get Permission

Comments (0)

Please log in or register to comment.
Log in