Exoskeleton formation in Apis mellifera: Cuticular hydrocarbons profiles and expression of desaturase and elongase genes during pupal and adult development

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Exoskeleton formation in Apis mellifera: Cuticular hydrocarbons profiles and expression of desaturase and elongase genes during pupal and adult development Q3

Tiago Falcón a, Maria Juliana Ferreira-Caliman b, Francis Morais Franco Nunes c, Érica Donato Tanaka a, Fábio Santos do Nascimento b, Márcia Maria Gentile Bitondi b, * a b c

Faculdade de Medicina de Ribeirão Preto, Universidade de São Paulo, Av. Bandeirantes 3900, 14049-900 Ribeirão Preto, SP, Brazil Faculdade de Filosofia, Ciências e Letras de Ribeirão Preto, Universidade de São Paulo, Av. Bandeirantes 3900, 14040-901 Ribeirão Preto, SP, Brazil Centro de Ciências Biológicas e da Saúde, Universidade Federal de São Carlos, Rod. Washington Luís, km 235, 13565-905 São Carlos, SP, Brazil

a r t i c l e i n f o

a b s t r a c t

Article history: Received 11 March 2014 Received in revised form 23 April 2014 Accepted 25 April 2014

Cuticular hydrocarbons (CHCs) are abundant in the superficial cuticular layer (envelope) of insects where they play roles as structural, anti-desiccation and semiochemical compounds. Many studies have investigated the CHC composition in the adult insects. However, studies on the profiles of these compounds during cuticle formation and differentiation are scarce and restrict to specific stages of a few insect species. We characterized the CHCs developmental profiles in the honeybee workers during an entire molting cycle (from pupal-to-adult ecdyses) and in mature adults (forager bees). Gas chromatography/mass spectrometry (GC/MS) analysis revealed remarkable differences in the relative quantities of CHCs, thus discriminating pupae, developing and newly-ecdysed adults, and foragers from each other. In parallel, the honeybee genome database was searched for predicted gene models using known amino acid sequences of insect enzymes catalyzing lipid desaturation (desaturases) or elongation (elongases) as queries in BLASTP analysis. The expression levels of six desaturase genes and ten elongase genes potentially involved in CHC biosynthesis were determined by reverse transcription and real time polymerase chain reaction (RT-qPCR) in the developing integument (cuticle and subjacent epidermis). Aiming to predict roles for these genes in CHC biosynthesis, the developmental profiles of CHCs and desaturase/elongase transcript levels were evaluated using Spearman correlation coefficient. This analysis pointed to differential roles for these gene products in the biosynthesis of certain CHC classes. Based on the assumption that homologous proteins may share a similar function, phylogenetic trees were reconstructed as an additional strategy to predict functions and evolutionary relationships of the honeybee desaturases and elongases. Together, these approaches highlighted the molecular complexity underlying the formation of the lesser known layer of the cuticular exoskeleton, the envelope. Ó 2014 Published by Elsevier Ltd.

Keywords: Cuticular hydrocarbons Apis mellifera Desaturase Elongase Cuticular envelope Insect exoskeleton

1. Introduction The cuticular exoskeleton of insects is mainly formed by proteins, the polysaccharide chitin, and lipids arranged as a complex multilayered structure: the inner procuticle comprising the endocuticle and exocuticle, the epicuticle and an outermost envelope. These functional layers are sequentially produced and are secreted by the epidermis at each molt cycle, and differ from each other in biochemical composition and physiological properties. The envelope

* Corresponding author. Tel.: þ55 1636023805. E-mail address: [email protected] (M.M. Gentile Bitondi).

mainly consists of lipids that form a barrier against water loss and invading pathogens, and also serve as important cues for chemical communication, besides acting as sex pheromones (Wigglesworth, 1970; Blomquist and Dillwith, 1985; Gibbs, 2002; Châline et al., 2005). This lipid layer is largely enriched with hydrocarbons (Hepburn, 1985), which are synthesized in specialized cells called oenocytes (Piek, 1964; Diehl, 1973; Schal et al., 1998; Fan et al., 2003; Billeter et al., 2009). In honeybee workers the oenocytes are localized in close association with the epidermis and the parietal fat body that internally coat the exoskeleton, and are more frequently found closer to the sternites than the tergites (Ruvolo and Landim, 1993). Intermediates and end-products of metabolic pathways, such as fatty acids, in addition to specific enzyme classes, are involved in

http://dx.doi.org/10.1016/j.ibmb.2014.04.006 0965-1748/Ó 2014 Published by Elsevier Ltd.

Please cite this article in press as: Falcón, T., et al., Exoskeleton formation in Apis mellifera: Cuticular hydrocarbons profiles and expression of desaturase and elongase genes during pupal and adult development, Insect Biochemistry and Molecular Biology (2014), http://dx.doi.org/ 10.1016/j.ibmb.2014.04.006

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CHC biosynthesis. Key enzymes in the CHCs biosynthetic pathways are the fatty acid synthases, elongases, desaturases, reductases and a P450 decarbonylase. Two forms of fatty acid synthases are likely involved in the synthesis of unbranched and methyl-branched fatty acids. Elongases catalyze the chain elongation of saturated and unsaturated fatty acids, which are converted to aldehydes by reductase enzymes. Aldehydes are substrates for the last step of hydrocarbons biosynthetic pathway, i.e., the oxidative decarbonylation catalyzed by a P450 enzyme. Desaturation, i.e., the insertion of carbonecarbon double bonds into the saturated fatty acid chain and consequent conversion to an unsaturated fatty acid is catalyzed by desaturases (Kolattukudy, 1965, 1968; Howard and Blomquist, 2005; Blomquist et al., 2012; Qiu et al., 2012). After being synthesized and released from the oenocytes, the CHCs are transported through the hemolymph by lipophorins (Chino and Kitazawa, 1981; Chino et al., 1981). CHCs reach the insect surface via the pore canals of the cuticular exoskeleton (Blomquist and Dillwith, 1985), where they integrate the envelope layer. The multicomponent CHC blend shows a great variation among insect species. CHCs also display both quantitative and qualitative intraspecific variation depending on the developmental stage, environment changes and food availability. The majority of the information on CHC composition comes from studies on adult social insects and is focused on their roles in chemical communication for sex-, kin- and caste-recognition (Dallerac et al., 2000; Roelofs et al., 2003; Liu et al., 2004; Châline et al., 2005; Nunes et al., 2008; Ferreira-Caliman et al., 2010). We have been studying the expression of genes, proteins and enzymes in the honeybee integument as part of a major project aiming to characterize the molecular elements involved in the exoskeleton formation and the regulation of this process (Bitondi et al., 1998; Santos et al., 2001; Zufelato et al., 2004; Soares et al., 2007, 2011, 2013; Elias-Neto et al., 2010). Such information generated using polyacrylamide gel electrophoresis (SDS-PAGE), western blot, RT-qPCR, gene sequencing, cDNA microarrays and fluorescence in situ hybridization (FISH), mainly highlighted the differential gene expression dynamics needed for the adult exoskeleton construction. These approaches, however, have neglected the outmost functional exoskeletal layer, i.e., the envelope, its formation and molecular composition. Trying to fill this gap in part we here used GC/MS to explore the composition as well as developmental profiles of CHCs during exoskeleton formation and maturation. Concomitantly, genes potentially encoding desaturases and elongases were searched in the honeybee genome and their expression patterns were characterized using RT-qPCR. To get an insight on the roles of the honeybee desaturases and elongases genes on cuticular envelope formation, the strength of the correlation between developmental profiles of transcripts and CHCs were estimated. In addition, we built molecular relationship trees for gene function prediction. 2. Material and methods 2.1. Sample collection Africanized Apis mellifera workers were obtained in the experimental apiary of the Faculty of Medicine, University of São Paulo in Ribeirão Preto, SP, Brazil. The samples were collected at successive developmental stages, from pupal ecdysis to a late adult stage, and included newly-ecdysed pupae (Pw phase: white eyes and unpigmented cuticle); pupae-in-apolysis (Pp phase: pink eyes and unpigmented cuticle); early, intermediate and late brown-eyed pharate adults showing unpigmented cuticle (Pb phase), partially pigmented cuticle (Pbm phase) and intensely pigmented cuticle (Pbd phase); newly-ecdysed adults (Ne) and foragers (Fg). These

developmental time points include the sequential events of the pupal-to-adult molt and adult cuticle synthesis, deposition and differentiation/maturation. The pre-ecdysial phases were identified according Michelette and Soares (1993) and all these developmental phases were used to analyze CHCs profiles and the expression of genes encoding enzymes potentially involved in their biosynthetic pathways. 2.2. CHCs extraction, identification and statistical analysis The samples (36 bees per developmental phase, except for Pw phase that comprised 35 bees) were individually added to 1.5 ml of 95% n-hexane (Mallinckrodt Chemicals) and bathed for 1 min to extract the CHCs. The extracts were then dried under N2 flow, resuspended in 160 ml of 95% n-hexane and analyzed in a Gas Chromatograph/Mass Spectrometer (Shimadzu GCMS model QP2010), equipped with a 30 m DB-5MS column and helium as the carrier gas (1 ml/min), using the electronic ionization (EI) method. The injection volume was 1 ml at an initial temperature of 150  C elevated at a rate of 3  C/min to 280  C and keeping this temperature for 15 min. Compounds identification was based on their diagnostic ions and in a standard solution containing different synthetic hydrocarbons (Fluka). To analyze the chromatograms we used the software GCMS solutions for Windows version 2.6 (Shimadzu Corporation). The positions of unsaturations in alkenes and alkadienes were identified according to the dimethyl disulfide derivatization technique (Carlson, 1989) and analyzed at the same GCMS system above mentioned. The initial temperature was 80  C for 2 min, then increased to 180  C at a rate of 30  C/min and then to 300  C at a rate of 3  C/min, keeping 300  C for 80 min. The identification of compounds and positions of unsaturations were done in splitless mode, which is recommended for low concentrated samples (Hü; bschmann, 2009), as those composed by a mixture of CHCs. Quantification of CHCs was based on their peak area in each chromatogram (Singer and Espelie, 1992). We adjusted the compounds percentage to 100% and each peak area was transformed according to the formula Z ¼ ln[Ap/g(Ap)], where Ap is the peak area, g(Ap) is the geometric mean of the peak for each bee sample and Z is the transformed peak area (Aitchison, 1982). To discriminate developmental stages according to their CHCs profile, we performed a PerMANOVA test in R software (version 3.0.0; package vegan: version 2.0e7). This test allowed us to analyze the dispersion of the samples around each group (developmental phase) centroid giving us back a result based on permutation tests (here we utilized 10000 permutations). We plotted the results in a scatterplot with the two first Principal Coordinates as axes. The variation in the proportion of each compound during development was also verified using the One-Way ANOVA associated to the Tukey’s Honestly Significance Difference (Tukey’s HSD) post hoc test (R software). We used this same test to compare variations in the proportions of CHC groups: n-alkanes, methylalkanes, dimethyl-alkanes, alkenes, alkadienes and non-identified compounds. 2.3. Identification of desaturases and elongases potentially involved in hydrocarbons biosynthesis Known desaturase and elongase amino acid sequences of Drosophila melanogaster and lepidopterans, available in the National Center for Biotechnology Information (NCBI) data bank (www.ncbi.nlm.nih.gov/), were used to search for homologous sequences in the honeybee genome (http://hymenopteragenome. org/beebase/) (version 4.5) using a BLASTP search tool (Altschul et al., 1990). The evolutionarily conserved functional motifs were confirmed in all the honeybee desaturase and elongase amino acid sequences, and the structural architecture of the respective

Please cite this article in press as: Falcón, T., et al., Exoskeleton formation in Apis mellifera: Cuticular hydrocarbons profiles and expression of desaturase and elongase genes during pupal and adult development, Insect Biochemistry and Molecular Biology (2014), http://dx.doi.org/ 10.1016/j.ibmb.2014.04.006

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nucleotide sequences, including intron/exon boundaries, was characterized using Artemis 7.0 software (Rutherford et al., 2000). 2.4. RNA extraction and quantification Honeybees at the Pw, Pp, Pbm, Ne and Fg developmental phases (3 bees per phase) were dissected in 0.9% NaCl solution, saving only the integument lining thorax and abdomen. Total RNA was extracted from individual integuments using 1 ml TRIzolÒ (Invitrogen) according to manufacturers’ instruction. After extraction, the RNA was suspended in autoclaved ultra-pure water treated with 0.1% diethylpyrocarbonate v/v (DEPC, Sigma). RNA samples were quantified in NanoDropÒ ND-1000 (NanoDrop Technologies) and A260nm/A280nm wavelength ratios between 1.9 and 2.0 were adopted as indicative of RNA quality and purity (Ausubel et al., 1995). The samples were diluted to a concentration of 0.6 mg/ml (considering that 1 OD unit measured at 260 nm corresponds to 40 mg RNA per ml; Sambrook et al., 1989). To control for genomic DNA contamination the samples were incubated with DNAse (0.1 unit RQ RNAse-free DNAse, Promega) for 30 min at 37  C, followed by 15 min incubation at 70  C to inactivate the enzyme. 2.5. First strand cDNA synthesis First strand cDNA was synthesized from total RNA using the SuperScriptÔ system (InvitrogenÔ, Life Technologies). The reverse transcription reaction was performed by incubating 0.6 mg of total RNA with 1 ml Oligo(dT)12-18 (500 mg/ml) and 1 ml dNTP (10 mM) at 65  C for 5 min, and then adding 1 ml SuperScriptÔ II Reverse Transcriptase 200U (Invitrogen) plus water to make 20 ml (final volume). This mixture was incubated at 42  C for 50 min, and then at 72  C for 15 min to inactivate the enzyme. As negative controls for semi-quantitative RT-PCR and RT-qPCR analyses, the SuperScriptÔ II enzyme was omitted in some reaction mixtures. 2.6. Semi-quantitative RT-PCR analysis The PCR reactions were performed in a Thermo Cycler VeritiÒ (Applied Biosystems). Each cDNA sample was used as template in reaction mixtures containing 10 ml of 2.5x Promega Master Mix [TaqeDNA Polymerase (0.06 unit/ml); 2.5x Taq-reaction buffer (125 mM KCl, 75 mM TriseHCl pH 8.4, 4 mM Mg2þ, 0.25% NonidetÒP40); 500 ml of each dNTP]; 10 pmol of each primer, 1 ml of cDNA and autoclaved ultrapure water to make a final volume of 20 ml. Specific primers were designed for six desaturase sequences and ten elongase sequences in the honeybee genome using the software Primer3 (version 0.4.0) (http://frodo.wi.mit.edu/primer3/). The respective accession numbers, primer sequences and the expected amplicon sizes are listed in Table S1. A primer pair (forward: 50 -TGC CAA CAC TGT CCT TTC TG-30 and reverse: 5 -AGA ATT GAC CCA CCA ATC CA-3 ) was designed for the gene encoding a ribosomal protein, AmRP49, currently renamed as ribosomal protein L32 (RpL32) (GenBank-NCBI accession number NM_001011587.1), which is expressed in similar levels during the honeybee development, and was validated as being a suitable reference gene for PCR normalization (Lourenço et al., 2008). All primers were designed to span an intron, thereby serving as control for genomic DNA contamination. PCR conditions were: 5 min at 94 C, 30 cycles of 30s at 94  C, 30 s at 60  C, 30 s at 72  C and a final extension for 7 min at 72  C. PCR-amplified cDNA aliquots (20 ml) were mixed with DNA loading buffer (0.25% bromophenol blue, 0.25% xylene cyanol FF and 30% glycerol) and analyzed by electrophoresis performed on GelRedÔ (Biotium) stained 1% agarose gel (0.1 ml/ml agarose gel) using 1x TBE (0.45 M Tris base, 0.45 M boric acid, 0.5 M EDTA, pH 8.0) as running buffer. The electrophoresis was

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performed under constant voltage (100V). The gel was analyzed in an image scanner KODAK EDAS 290. A 100 bp lDNA/Hind III (InvitrogenÔ, Life Technologies) was used as molecular weight marker. 2.7. RT-qPCR analysis RT-qPCR analysis was performed in a 7500 Real Time PCR System (Applied Biosystems) using primers for desaturase and elongase genes (Table S1) and for the reference gene. Standard curves were performed for each primer to verify amplification efficiency (parameters used: 3.6  Slope  3.3; R2  0.99). Reaction mixtures were prepared with 10 ml of 2x Sybr Green PCR Master Mix (Applied Biosystems), 0.8 ml of each specific primer of a pair, and 2 ml of cDNA in a final volume of 20 ml completed with DEPC treated-autoclaved milli-Q water. The amplification program was: 2 min at 52  C and 10 min at 95  C; 40 cycles of 1 min at 94  C, 30 s at 60  C and 30 s at 72  C. Dissociation curve for each primer pair allowed verifying the quality of amplification. Triplicates were prepared for each sample (technical triplicates), and three independent samples were used for each developmental phase (biological samples). The relative quantification was calculated using 2DDCt (Schmittgen and Livak, 2008). The transcript levels were compared between phases using One-Way ANOVA and Tukey’s HSD post hoc test (p < 0.05) (R software). Desaturase and elongase transcripts profiles were correlated with CHCs profiles using Spearman test and R software (p < 0.05). 2.8. Amplicons validation PCR amplicons were sequenced using the primers cited above. After amplification for 5 min at 94 C, 40 cycles of 30 s at 94  C, 30 s at 60  C, 30 s at 72  C and a final extension for 7 min at 72  C, the cDNA was quantified in NanoDrop and 1 ml was mixed with 1 ml of forward primer (F) or reverse primer (R), 2 ml of Big DyeÒ Terminator solution v3.1 Cycle Sequencing Kit (Perkin Elmer), 2 ml of buffer solution (5x Sequencing Buffer) and 4 ml of ultrapure autoclaved water, totalizing 10 ml of reaction solution. We used 25 cycles of 10 s at 96  C, 5 s at 50  C and 4 min at 60  C. The samples were precipitated with 40 ml of 75% isopropanol, incubated at room temperature for 15 min and centrifuged at 12,100  g during 20 min. The supernatant was discarded, the pellet was dried at 55  C, resuspended in 12 ml of Hi-DiTM formamide and transferred to sequencing tubes to be submitted to a thermal shock (2 min and 5 s, at 95  C, followed by 5 min on ice). Samples were analyzed by the method of Sanger in an automatic sequencer ABI Prism 310 (Applied Biosystems). We also confirmed the sequences through in silico PCR analysis (http://genome.ucsc.edu). 2.9. Molecular relationship analysis of desaturases and elongases Amino acids sequences of desaturases and elongases from arthropods were searched using the same approach described in the section “2.3.”. Only complete amino acid sequences were used for the analysis, except for the desaturase sequences corresponding to the GB42217 and GB48194 accession numbers. The multiple ̊ alignments were performed using the software MAFFT v. 7 (Katoh et al., 2002) (http://mafft.cbrc.jp/alignment/server/) with default parameters. This software has a better performance than other softwares when analyzing a great number of terminals (>50) (Katoh et al., 2005). These alignments were visualized using the software CLustalX 2.1 (Larkin et al., 2007) to identify the enzymes motifs. The arthropod species used in the analysis as well as the desaturases and elongases accession numbers at NCBI are listed in Table S2.

Please cite this article in press as: Falcón, T., et al., Exoskeleton formation in Apis mellifera: Cuticular hydrocarbons profiles and expression of desaturase and elongase genes during pupal and adult development, Insect Biochemistry and Molecular Biology (2014), http://dx.doi.org/ 10.1016/j.ibmb.2014.04.006

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Fig. 1. Cuticular hydrocarbons (CHCs) profiles during A. mellifera development. Compounds were extracted with n-hexane from the cuticle of pupae (Pw and Pp phases), developing adults (Pb, Pbm and Pbd phases), newly-ecdysed adults (Ne) and forager (Fg) bees, and identified by GC/MS.

Please cite this article in press as: Falcón, T., et al., Exoskeleton formation in Apis mellifera: Cuticular hydrocarbons profiles and expression of desaturase and elongase genes during pupal and adult development, Insect Biochemistry and Molecular Biology (2014), http://dx.doi.org/ 10.1016/j.ibmb.2014.04.006

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Fig. 1. (continued).

We performed a model test for both desaturase and elongase sequences using the software ProtTest v. 3.2 (Darriba et al., 2011) with default parameters. The indicated model for both enzymes was LG (Le and Gascuel, 2008), which represents a standard matrix of substitution of amino acids, with an estimative of invariable sites (þI) and a gamma heterogeneity rate between sites (þG) for both groups of enzymes. The eLnL (log likelihood) values were 39,255 for desaturases and 41,945 for elongases. For both groups of enzymes, we used the software PhyML, version 3.0 (Guindon et al., 2010). The molecular phylogeny of desaturases was performed using maximum likelihood analysis (Felsenstein, 1981) with 500 bootstrap replications. Desaturase protein sequences of the crustacean Daphnia pulex and the tick Amblyomma americanum (Liu et al., 1999; Roelofs and Rooney, 2003) were used as external groups. The tick desaturase sequence was used for rooting the tree, once unlike D. pulex it has only one desaturase sequence. We also used BLASTP (Altschul et al., 1990) to compare each sequence with the following desaturase sequences whose functions were already validated: D. melanogaster desat1 (NP_731710), a D9 desaturase that uses preferentially palmitic acid (C16) over stearic acid (C18) (16 > 18) as substrate; D. melanogaster desat2 (NP_650201), a D9 desaturase that prefers myristic acid (C14) and synthesizes D5 hydrocarbons (Dallerac et al., 2000); D. melanogaster desatF (NP_651996), a desaturase that uses the u-7 product from desat1

unsaturation as substrate (Chertemps et al., 2006); Ostrinia furnacalis D14 desaturase (AAL35746); Helicoverpa zea D9 desaturase (AAF81790) that acts on stearic acid (18 > 16) (reviewed in Roelofs and Rooney, 2003); Trichoplusia ni D9 desaturase (AAB92583) with a high specificity for stearic acid (18 > 16) (Liu et al., 1999), and D11 desaturase (AAD03775) (Knipple et al., 1998); Choristoneura rosaceana D9 desaturase (AAN39697) that acts on palmitic acid (16 > 18) (Hao et al., 2002); Danaus plexippus D14 desaturase (EHJ69993); Bombyx mori D11 desaturase (NP_001040141); Lampronia capitella D14-26 desaturase (ABX71629); Choristoneura paralela D14-26 desaturase (AAQ12887); Helicoverpa assulta D1426 desaturase (AF482905) (Liénard et al., 2008). For the molecular phylogeny of elongases we performed a cluster analysis using an approximate likelihood ratio test (aLRT) (Anisimova and Gascuel, 2006). The generated trees were visualized and edited using the software FigTree v.1.3.1 (http://tree.bio.ed. ac.uk/software/figtree/). 3. Results 3.1. CHC profiles during cuticle formation and maturation Five CHC classes including n-alkanes, methyl-alkanes, dimethylalkanes, alkenes and alkadienes, with chain-length of the carbon

Please cite this article in press as: Falcón, T., et al., Exoskeleton formation in Apis mellifera: Cuticular hydrocarbons profiles and expression of desaturase and elongase genes during pupal and adult development, Insect Biochemistry and Molecular Biology (2014), http://dx.doi.org/ 10.1016/j.ibmb.2014.04.006

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backbones ranging from C19 to C35, were identified by GC/MS in the cuticle of newly-ecdysed pupae (Pw phase), pupae-in-apolysis (Pp phase) and in the cuticle of developing adults (Pb, Pbm and Pbd phases), newly-ecdysed adults (Ne) and mature bees (Fg). Individually, the five peaks that better explain the total variation here found were Z-?- C35 (3.754%)/Z-7-C23, Z-9-C23 (3.746%)/ZZ-?-C33a (3.551%)/Z-10-C35, Z-12-C35, Z-14-C35, Z-15-C35 (3.223%)/5-MeC25 (2.995%). Fig. 1 shows the variation in the proportions of the specific compounds in each CHC class during this developmental period (see mean values and statistics in Table S3). The most represented n-alkane in the developing and adult cuticle is C27. C27 is proportionally more abundant in foragers, and C25 is the second more abundant n-alkane in foragers. C24 and C26, which are less represented in the cuticle, also showed increased percentages in foragers. The other n-alkanes showed diverse developmental profiles: C19, C21, and C23 proportions increased significantly at the Pbm or Pbd phases and then decreased in foragers. C28, C29 and C30 were relatively more abundant in the cuticle of the immature developmental phases and newly-ecdysed adults than in foragers. C31 showed similar proportions in the cuticle of some immature phases and foragers (Fig. 1A). The most represented methyl-alkanes in the developing cuticle were 9-;11-;13-MeC27, 9-;11-; 13-;15-MeC29 and 13-;15-MeC31. Except for 7-;9-MeC33, the other sixteen identified methyl-alkanes (94%) were proportionally more concentrated in the immature cuticles (often including the cuticle of the newly-ecdysed adults) than in the cuticle of foragers (Fig. 1B). This developmental pattern is repeated for the dimethyl-alkanes, all of them (100%) presenting relatively higher quantities in the immature cuticle (Fig. 1C). Therefore, the great majority of the methyl-alkanes and all identified dimethyl-alkanes lowered to basal levels in the cuticle of foragers. In contrast to these branched compounds (and similarly to the C24, C25, C26 and C27 n-alkanes), most of the alkenes (Fig. 1D) and alkadienes (Fig. 1E) were overrepresented in the cuticle of foragers. Although two of the alkenes (Z-7-C23, Z-9-C23) has peaked in the Pbd phase and another peak of alkenes (Z-8-C28, Z-9-C28) has decreased in foragers, and one of the alkadienes (ZZ-?-C27) showed the higher concentration in the Pbd phase and newly-ecdysed adults, our data confidently show that in general, the proportion of both unsaturated CHCs, alkenes and alkadienes, increases in the mature cuticle of foragers. Table 1 summarizes the variation in the proportions of n-alkanes, methyl-alkanes, dimethyl-alkanes, alkenes and alkadienes in the cuticle during development. Foragers clearly differentiated from immature bees and newly-ecdysed adults regarding proportions of CHC classes. The overrepresented CHC class in the cuticle of foragers is the class of n-alkanes, which makes up 71.92% of the total CHCs. It is followed by the alkenes, representing 22.28% of the CHCs. The immature cuticles of the developing bees and newly-ecdysed adults also contain a relatively high proportion of nalkanes, comprising 44.88e57.43%, but the methyl-alkanes are also

Table 1 Proportions of CHC chemical classes during adult cuticle formation and maturation.

abundant (35.15e47.47%) in these cuticles. It was evident that methyl-alkanes and dimethyl alkanes exist in significant higher amounts in the immature cuticle in comparison to foragers. In contrast, alkenes and alkadienes are both proportionally more abundant in the cuticle of foragers than in the cuticle of the earlier developmental phases. Although qualitative variations in CHC profiles during development have not been detected, all developmental phases quantitatively differed from each other (PerMANOVA test, PseudoF ¼ 214.64. p < 1e-05) (Fig. 2). This analysis, which included all the identified and non-identified compounds, revealed significant differences across all developmental phases and clearly separated them in three groups according to the relative quantities of CHCs: one group including pupae (Pw and Pp phases) and the successive Pb and Pbm phases, a second group formed by the pre- and postecdysial phases (Pbd and Ne), and a third group formed exclusively by foragers (Fg). Thus, the obtained data indicate that the pupal and adult (immature and mature) cuticles share a similar CHC blend, but with great differences in the relative quantities of the majority of the compounds. All CHC chemical classes were identified by GC/MS in the developing and adult cuticles and were represented in the principal component analysis (Table S3). An example of the chromatograms is shown in Fig. S1. 3.2. Molecular phylogeny of desaturases and elongases All seven desaturases included in the phylogenetic analysis are members of the subfamily First Desaturase (Hashimoto et al., 2008). The deduced amino acid sequences of all desaturases herein characterized contain the eight conserved histidine residues arranged as three box domains, which are typical of integral membrane desaturases and are essential for the catalytic activity and function (Table S2 and Fig. S2). The desaturases grouped according to their functions (position where the unsaturation is created in the fatty acid) (Fig. 3). Six of them (GB42218, GB51236, GB51238, GB48194, GB42217, GB48195) displayed the highest similarity with D9-desaturases (16 > 18) of other organisms, which catalyses the insertion of a double bond at the 9th position from thecarboxyl group of a fatty acid. The desaturase encoded by the gene GB48193 clustered with D4-desaturases creating a carbon/ carbon double bond at the 4th position. The honeybee elongase sequences have the conserved domain formed by three histidine residues and clustered in two groups (Fig. 4, Table S2 and Fig. S2) with different functional motifs. Eleven elongase sequences were included in the phylogenetic analysis. Three of them (GB53873, GB53872 and GB45596) belong to the S/ MUFA cluster and should have roles in elongating Saturated/ Monounsaturated Fatty Acids. Eight elongases (GB51249, GB51247, GB54401, GB54302, GB54396, GB54399, GB40681 and GB46038) clustered into the PUFA class, which has roles in Polyunsaturated Fatty Acids elongation. Interestingly, except for the PUFA elongase genes GB54399 and GB40681, all the other PUFA genes with expression quantified by RT-qPCR showed a positive correlation with the alkadienes.

Proportions of CHC groups (%) Phase

Pw Pp Pb Pbm Pbd Ne Fg

Saturated CHCs

Unsaturated CHCs

n-Alkanes

Methylalkanes

Dimethylalkanes

Alkenes

Alkadienes

47.82 50.87 44.88 45.20 57.44 51.53 71.93

44.10 43.42 47.47 46.57 35.15 38.19 2.59

3.87 2.66 3.72 3.28 1.99 2.80 0.11

3.86 2.60 3.34 4.47 4.97 6.39 22.28

0.01 0.01 0.01 0.01 0.19 0.60 2.60

Nonidentified

3.3. Structure and expression profiles of desaturase and elongase genes in the integument during cuticle formation and maturation

0.34 0.44 0.57 0.47 0.26 0.49 0.49

Six of the seven desaturase sequences and ten of the eleven elongase sequences included in the phylogenetic analyses were characterized in terms of gene structure and expression in the integument. The six desaturase coding sequences (CDSs) have 663 to 1371 nucleotides (stop codon included) and encompass 5 to 8 exons that potentially encode 220 to 456 amino acid residues. The ten elongase CDSs contain 420 to 996 nucleotides distributed into 2

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Fig. 2. Principal coordinate analysis of CHC profiles discriminating developmental phases of A. mellifera workers. Axes are represented by principal coordinates. PerMANOVA test (Pseudo-F ¼ 214.64, p < 1e-05). The centroid is marked in red for each developmental phase. Dashed lines highlight link the more distant samples in each group. Blue lines link samples to centroid. Developmental phases are represented by symbols: pupae (Pw V); pupae-in-apolysis (Pp e >); developing adult phases: Pb (D), Pbm (x) and Pbd (þ); newlyecdysed adult: Ne (⌧); forager bee: Fg (B). (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

to 7 exons that encode proteins formed by 139e331 amino acids. Expected canonical splice sites (GT/AG) were found at all exon/ intron boundaries. The interprimer regions (amplicons) were sequenced and validated (Fig. S3). Some of the honeybee desaturase and elongase genes are organized in tandem in the chromosomes. The desaturase genes GB42217 and GB42218, both encoding amino acid sequences that grouped together with D9-desaturases in the phylogenetic analysis (Fig. 3), are clustered on chromosome 1. Six of the elongase genes are arranged into two clusters on chromosome 16, one cluster including the genes GB54401, GB54302, GB54399 and GB54396, and the other formed by the GB51249 and GB51247, all of them encoding elongases of the PUFA class as suggested by the phylogenetic analysis (Fig. 4). Four D9-desaturase genes (GB51238, GB48195, GB42218 and GB42217) and the D4-desaturase gene (GB48193) showed significant transcript levels increase in the integument of newly-ecdysed (Ne) and/or forager (Fg) bees in comparison to the earlier developmental phases (Pw, Pp and Pbm phases). The desaturase gene identified as GB51236 (D9 desaturase) was the only gene presenting similar levels of transcripts during development (Fig. 5). Ten of the genes potentially encoding elongases showed variable expression profiles. Genes encoding elongases of the PUFA class (GB51249, GB51247 and GB54396) and genes encoding S/ MUFAs (GB53872 and GB45596) showed higher expression in the newly-ecdysed adults and/or forager bees than in the earlier developmental phases. Two other elongase genes in the PUFA class also showed significantly higher expression in foragers but only when compared with the Pp phase (gene GB54401) or Pbm phase (gene GB46038). The gene GB40681potencially encoding an elongase of the PUFA class showed a different expression profile with a higher level of transcripts in the Pbm phase than in the earlier or later phases. The expression profiles of the other PUFA elongase genes, GB54399 and GB53873, did not show significant variation in the developing integument (Fig. 6). 3.4. Desaturase and elongase transcript levels variation versus CHCs profiles during cuticle formation and maturation Sperman’s correlation coefficient was used to measure the interdependence between the changing levels of CHCs and desaturase or elongase transcripts during cuticle development (Fig. 7;

Table S4). In general, we found significant positive correlations between the increase in desaturase transcript levels during development and the increase in the proportions of 13 alkenes peaks (Z-7C23, Z-9-C23/Z-?-C25/Z-6-C27, Z-7-C27, Z-9-C27/Z-?-C27/Z-7-C29, Z-9C29/Z-?-C29/Z-5-C25, Z-7-C25, Z-9-C25/Z-7-C31, Z-8-C31, Z-9-C31, Z-10C31, Z-11-C31, Z-15-C31/Z-8-C33, Z-9-C33, Z-10-C33, Z-12-C33/Z-9-C33, Z-10-C33, Z-13-C33/Z-?-C33/Z-10-C35, Z-12-C35, Z-14-C35, Z-15-C35/Z?-C35) and 6 peaks of alkadienes (ZZ-?-C27/ZZ-?-C31/ZZ-?-C33a/ZZ?-C33b/ZZ-?-C35a/ZZ-?-C35b), representing respectively 93% and 100% of the total of the identified alkenes and alkadienes. Therefore, the increase in the products of desaturation reactions is possibly due to increased transcription of desaturase genes. These correlation data allow us to infer that the six desaturase genes, or rather, their respective protein products, promote an increase in the biosynthesis of alkenes and alkadienes in adult bees by the insertion of double bonds into the fatty acid (hydrocarbon) backbone chains. Interestingly, the expression of five of the desaturase genes (GB42218, GB42217, GB48195, GB51238 and GB48193), with a few exceptions, were positively correlated with the same pool of unsaturated CHCs, alkenes (Z-10-C35, Z-12-C35, Z-14-C35, Z-15-C35/Z?-C29/Z-?-C35/Z-?-C33/Z-5- C25, Z-7-C25, Z-9-C25/Z-9-C33, Z-10-C33, Z-13-C33/Z-7-C23, Z-9-C23/Z-8-C33, Z-9-C33, Z-10-C33, Z-12-C33/Z-7C31, Z-8-C31, Z-9-C31, Z-10-C31, Z-11-C31, Z-15-C31/Z-?-C25) and alkadienes (ZZ-?-C33a/ZZ-?-C33b/ZZ-?-C31/ZZ-?-C35a/ZZ-?-C35b) (see the dark blue area at the left of Fig. 7). The other desaturase gene, GB51236, which in contrast to the five desaturase genes above mentioned did not show a significant increase in expression in foragers, exhibited a more restrictive correlation pattern. Its transcriptional modulation over time was positively correlated only with some of these alkenes (Z-8-C33, Z-9-C33, Z-10-C33, Z-12-C33/Z?-C25) and alkadienes (ZZ-?-C31/ZZ-?-C33b/ZZ-?-C35a/ZZ-?C35b) and with other peaks of alkenes (Z-?-C27/Z-7-C29, Z-9-C29/Z-6-C27, Z-7C27, Z-9-C27) (Fig. 7). Such differences in correlation patterns may reflect enzyme specificities in desaturation reactions. Unexpectedly, significant positive correlation was also found between the levels of four of the desaturase transcripts (GB42217, GB48195, GB51238, GB48193) and some alkanes (C23/C24/C25/C31), and methyl-alkanes (7-MeC33/9-MeC33). Except for the alkane C25, the expression of the desaturase gene GB42218 was also positively correlated with the levels of these same alkanes and methyl-alkanes, and the expression of the desaturase gene GB51236 was positively

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correlated with C27 and C26 levels (Fig. 7). All these alkanes correspond to a fraction (26%) of the total alkanes identified, and the changing levels of the majority of them showed no correlation, or showed negative correlation, with desaturase transcripts levels modulation. As most of the desaturase genes, the expression of five of the elongase genes (GB53872, GB54396, GB51249, GB45596 and GB51247) also increased significantly in newly-ecdysed adults and/ or foragers. Consequently, the correlation pattern of these elongase genes mimicked the correlation pattern of the desaturase genes. The expression profiles of these elongase genes, and also the expression profiles of two other elongases genes, GB53873 and GB54401, showed a positive correlation with the same CHCs as specified above, mainly alkenes and alkadienes, except Z-?-C27 (see the dark blue area at the left of Fig. 7). Such compounds represent respectively 86% and 100% of the total of alkenes and alkadienes herein identified. This suggests that these seven elongase genes act synchronically with the five desaturase genes above mentioned in the biosynthesis of these unsaturated CHCs. The transcript profiles of the other three elongase genes were positively correlated with the profiles of other types of hydrocarbons. Thus, the expression of GB54399 was correlated with the methyl–alkanes 9-;11-;13-;15-MeC29, 5-MeC25 and 7-MeC25, and the expression of GB40681 was correlated with the dimethylalkanes alkanes 3,7-diMeC31, 9,13-diMeC29, 11,15-diMeC27 and the alkenes Z-7-C31, Z-8-C31, Z-9-C31, Z-10-C31, Z-11-C31, Z-15-C31. The elongase gene GB46038 showed a more extensive correlation pattern including alkanes (C27/C26/C25), alkenes (Z-7-C29, Z-9-C29/Z6-C27, Z-7-C27, Z-9-C27/Z-8-C33, Z-9-C33, Z-10-C33, Z-12-C33/Z-7-C31, Z-8-C31, Z-9-C31, Z-10-C31, Z-11-C31, Z-15-C31) and alkadienes (ZZ?-C31/ZZ-?-C33b/ZZ-?-C35a/ZZ-?-C35b). Interestingly, the developmental profiles of some compounds were specifically correlated with the changing expression of a single elongase gene. For example, the methyl-alkanes 9-;11-;13-MeC23, 9-;11-;13-MeC25, 7MeC25, 5-MeC25 and 9-;11-;13-; 15-MeC29 showed a high positive correlation exclusively with the expression of the GB54399 elongase gene. Similarly, the dimethyl-alkanes 11,15-diMeC27, 9,13diMeC29 and 3,7-diMeC31 were positively correlated exclusively with the GB40681 elongase gene (see Fig. 7). This may indicate that the enzymes encoded by these genes act preferentially on the biosynthesis of methyl-branched CHCs. The developmental profiles of a fraction of alkanes (C30/C28/C29), methyl- or dimethyl alkanes (13- MeC31,15- MeC31,17-MeC31/3MeC31,5-MeC31,7-MeC31,10-MeC31,12-MeC31,13-MeC31,15-MeC31/9MeC31,12-MeC31,14-MeC31/5-MeC33,11-MeC33,13-MeC33,14-MeC33, 15-MeC33,16-MeC33/13-MeC35,15-MeC35,16-MeC35,17-MeC35/10,15diMeC29/11-MeC26,12-MeC26/7,11diMeC27, 7,15diMeC27/6,10diMeC27, 11,15-diMeC27/4,8-diMeC27, 13,15-diMeC27/7-MeC27/5MeC27/9- MeC27,11- MeC27,13MeC27/12- MeC28,13- MeC28,14MeC28) (see green area in Fig. 7) in general showed negative correlation with the expression of all the elongase genes herein characterized. This suggests that the products of these genes do not have roles in the biosynthesis of these specific saturated CHCs. 4. Discussion 4.1. Overrepresented CHC classes in the immature and mature cuticle Fig. 3. Evolutionary relationships of insect desaturases. Different clusters are marked in different colors with their putative function. Branches: Hymenoptera (blue); Diptera (red); Lepidoptera (green); Coleoptera (yellow); Crustacea (purple); Chelicerata (orange). The A. mellifera enzymes are written in blue. The horizontal bar indicates the scale of amino acids substitutions in branches. The values in the nodes are the branch support after 500 bootstrap replications. The first letter in each enzyme name indicates the gender and the following three letters indicate the species, followed by the

We characterized the CHC profiles of honeybee workers during an entire molting cycle, from the pupal ecdysis to the adult ecdysis,

accession number of their respective amino acid sequences (see Table S2). (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

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and also in mature adults. The genuine honeybee worker pupa, represented by the Pw phase, exists during a relatively short period, lasting approximately 40 h from the pupal ecdysis. Apolysis, i.e., the detachment of the pupal cuticle occurs at the Pp phase and signals the end of the pupal phase and the onset of adult cuticle deposition throughout the Pb to Pbd phases. The developing adult cuticle becomes increasingly thicker, and this is followed by the ecdysis and emergence of the adult bee from the comb cell. Such sequential events characterize the molting cycle that transforms the pupa into an adult bee. CHC profiles determination through GC/MS allowed us to discriminate the cuticles of all these molting cycle-phases and also the mature cuticle of forager bees. The Principal Coordinate Analysis clustered these developmental time points into three groups: one including the pupal cuticle (Pw and Pp phases) and the cuticle of the subsequent Pb and Pbm phases, one other formed by the preand post-ecdysial phases (Pbd phase and the newly-ecdysed adults), and the other exclusively formed by mature foragers. This means that these groups significantly differ from each other in the relative quantity of CHCs. Variations in the relative quantities of a number of CHCs between the pupal cuticle (Pw and Pp phases) and the cuticle of the subsequent Pb and Pbm phases may be interpreted as being due to the incorporation of hydrocarbons into the growing (nascent) adult cuticle. Our analysis showed that the mature cuticle of the forager bees is by far the most dissimilar in CHC proportions. These results go along with the hypothesis that, in social bees, the cuticle is fully mature only later in the adult life when they start foraging activities. This hypothesis is based on comparative morphological study that revealed striking differences in the cuticular maturation, and thus in cuticle properties, of workers of social and solitary bee species. In contrast to the solitary bee species that emerge from the nest with a heavily pigmented/sclerotized cuticle and immediately starts foraging, the social species showed the higher degree of pigmentation/sclerotization only several days after the emergence when they leave the intranidal activities for foraging for nectar and pollen (Elias-Neto et al., 2013). In this context, the next step would consist in comparing the CHCs profiles of social and solitary bees at definite time points of the adult life, which certainly is an interesting way of testing this hypothesis. Approximately half of the identified n-alkanes, and almost all of the identified unsaturated CHCs (alkenes and alkadienes) showed higher proportions in the mature cuticle of forager bees than in the earlier developmental phases. The opposite was verified for the saturated, branched CHCs (methyl-alkanes and dimethyl-alkanes), which were less represented in the cuticle of foragers, but overrepresented in the immature cuticles of pupae (Pw and Pp phases), developing adults (Pb, Pbm and Pbd phases) and newly-ecdysed adults. Such differences seem due to the intensification of the biosynthesis and/or deposition of n-alkanes, alkenes and alkadienes on the cuticle of foragers, since 40% of the total of the identified compounds (n ¼ 55) were proportionally more concentrated in foragers than in the earlier developmental phases. The higher relative quantities of methyl-alkanes and dimethyl-alkanes in the immature cuticles suggest that branched compounds are important for the structure of the envelope at these stages, or for the communication between brood and worker bees inside the

Fig. 4. Evolutionary relationships of insect elongases. Enzymes were clustered based on their functional domains: S/MUFAs (blue area) and PUFAs (green area). Branches: Hymenoptera (blue); Diptera (red); Lepidoptera (green); Coleoptera (yellow);

Crustacea (purple); Chelicerata (orange). A. mellifera elongases are written in blue. The horizontal bar indicates the scale of amino acids substitutions in branches. The values in nodes are the aLRT support. The first letter in each enzyme name indicates the gender and the following three letters indicate the species, followed by the accession number of their respective amino acid sequences (see Table S2). (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

Please cite this article in press as: Falcón, T., et al., Exoskeleton formation in Apis mellifera: Cuticular hydrocarbons profiles and expression of desaturase and elongase genes during pupal and adult development, Insect Biochemistry and Molecular Biology (2014), http://dx.doi.org/ 10.1016/j.ibmb.2014.04.006

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Fig. 5. Expression of desaturase genes in the pupal integument (Pw and Pp phases) and in the integument of developing adults (Pbm phase), newly-ecdysed adult (Ne) and forager bees (Fg). Transcript levels (columns) were determined by RT-qPCR. Different letters above each error bar indicate significant statistical difference (ANOVA associated to post hoc Tukey’s HSD test, p < 0.05) between developmental phases. The genes are identified by their accession numbers in the honeybee genome (version 4.5).

hive. Interestingly, new types of compounds were not identified as the cuticle became mature indicating that the blends of CHCs (but not the proportions of individual CHCs) are similar in the pupal and adult cuticles. Therefore, the CHC classes and types present in the pupal cuticle are also present in the growing and mature adult

cuticles, but show clear differences in relative quantities. The results of our CHC analysis in the newly-ecdysed adults and foragers are comparable with the results of Blomquist et al. (1980a). These authors similarly found a higher proportion of saturated than unsaturated compounds and an increase in the proportions of

Fig. 6. Expression of elongase genes in the pupal integument (Pw and Pp phases) and in the integument of developing adults (Pbm phase), newly-ecdysed adults (Ne) and foragers (Fg). Transcript levels (columns) were determined by RT-qPCR. Different letters above each error bar indicate significant statistical difference (ANOVA associated to post hoc Tukey’s HSD test, p < 0.05) between developmental phases. The genes are identified by their accession numbers in the honeybee genome (version 4.5).

Please cite this article in press as: Falcón, T., et al., Exoskeleton formation in Apis mellifera: Cuticular hydrocarbons profiles and expression of desaturase and elongase genes during pupal and adult development, Insect Biochemistry and Molecular Biology (2014), http://dx.doi.org/ 10.1016/j.ibmb.2014.04.006

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Fig. 7. Correlation between the developmental profiles of desaturases and elongases transcript levels and the relative proportions of saturated and unsaturated CHCs during adult cuticle formation and maturation (Spearman test, p < 0.05).

pentacosane (C25), heptacosane (C27), pentacosenes (C25:1) and heptacosenes (C27:1) in older bees (26 day-old), possibly foragers, in comparison to the younger bees (7-day old) used in their study. Like Blomquist et al. (1980a, 1980b) we detected 23 to 35 carbons chainlength alkenes, and 31, 33 and 35 carbons chain-length alkadienes, but another alkadiene (27 carbons chain-length) was also detected in our analysis of the adult cuticle. Concerning some discrepancies between both data, they could be due to the use of adult bees of different ages and subspecies: our data were obtained with Africanized A. mellifera, the hybrid produced by cross-breeding of the African A. m. scutellata and European honeybees, whereas Blomquist et al. (1980a, b) worked with the Italian race. Our results on the adult stage also match those of Kather et al. (2011) in demonstrating that the pool of CHCs in forager honeybees is mainly composed of n-alkanes and that these compounds and alkenes have higher proportions in foragers than in newlyecdysed adults. In addition to their structural and physiological

functions in the cuticular envelope, the alkenes may be important for nestmate recognition (Châline et al., 2005; Dani et al., 2005; Kather et al., 2011) whereas n-alkanes are less informative in this aspect of insect sociality (Krasnec and Breed, 2013). To our knowledge only two previous studies have investigated CHCs in immature stages of the honeybee, and even so, using punctual stages. Nation et al. (1992) and Martin et al. (2001) used GC/MS to investigate the CHC mimicry of honeybee host by the ectoparasitic mite Varroa jacobsoni. At a stage identified as “purpleeye pupa”, apparently equivalent to our pupa-in-apolysis (Pp phase), Nation et al. (1992) found relatively low proportions of alkanes in comparison with the adults, these having significantly less branched alkanes than the pupa. This was confirmed by our analysis. Alkenes were found in proportionally higher quantities in the adult cuticle. This was also evident in our analysis, which additionally showed a significant increase in the proportions of alkadienes on the adult cuticle. Martin et al. (2001) used newly-ecdysed

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adult bees, five-day-old larvae and a stage identified as “eight-dayold pupae”, which seems equivalent to our Pbm phase, in a comparative analysis with CHCs from the mite. Like our findings, these analyses did not reveal qualitative differences, but quantitative differences in the CHC pools of developing honeybees. CHC developmental profiles were also characterized in other insect species. Similarly to our findings, the n-alkanes detected in pupae of T. ni were in general also detected in the 3-day-old adult females, and the proportions of these compounds were higher in the adult moths. However, differently from the honeybees, the branched CHCs of T. ni showed qualitative variation during development (de Renobales and Blomquist, 1983). There was also no qualitative variation in the CHC profiles in larvae, pupae and adults of Sarcophaga bullata, but quantitative variations (Armold and Reignier, 1975). Taken together, these few studies on characterization of CHC profiles revealed remarkable differences in the relative quantities of these compounds during development. There is evidence in T. ni that biosynthesis and transport of hydrocarbons to the cuticular surface are highest during the feeding periods of the successive larval instars. As the larva prepares for each molt and stops feeding, hydrocarbon biosynthesis and transport decrease at different rates, the decrease in transport being even greater, such as the internal hydrocarbons accumulate and are then used as a source of CHC for the next larval instar (Dwyer et al., 1986). During the pupal-to-adult molt of T. ni, hydrocarbons biosynthesis was limited to two specific time periods: soon after the pupal molt and preceding the adult ecdysis. The produced hydrocarbons remained stored internally up to the adult ecdysis and then were deposited on the adult cuticle (de Renobales et al., 1988). Similar results were obtained in another lepidopteran, Spodoptera eridania (Guo and Blomquist, 1991) and in the dipteran S. bullata (Armold and Regnier, 1975). Even in hemimetabolous such as Blattella germanica, the patterns of hydrocarbons biosynthesis were correlated to molting and food intake during nymphal stages (Young and Schal, 1997). It follows that the deposition of CHCs does not reflect the rate of biosynthesis, since most of the CHC pool in a given stage were produced in the previous stage. These data on the dynamics of biosynthesis and deposition of CHCs allow us to infer that the CHCs integrating the pupal cuticle (Pw and Pp phases) and the cuticle of the subsequent developmental phases (Pb, Pbl and Pbm phases) of the honeybee are remnant of larval synthesis. The CHCs detected at the time of adult ecdysis, i.e., at the Pbd and Ne phases, may result from post-pupal molt biosynthesis. The increase in the proportions of several CHCs in forager bees is then possibly due to feeding and de novo biosynthesis at the expenses of precursors derived from the metabolism of dietary compounds. This idea receives support from the expression profiles of several desaturase and elongase genes, which showed increased transcript levels in forager bees in comparison to the immature phases. 4.2. Evolution of desaturases and elongases in Hymenoptera Genes encoding proteins/enzymes using fatty acid as substrates generally show duplications and have accumulated mutations, and this may imply in new functions (Karanth et al., 2009). The desat1 and desat2 desaturase genes of D. melanogaster are tandemly arrayed in a region of the chromosome 3 and could be evolved from a single gene through an initial duplication (Wicker-Thomas et al., 1997). The in tandem chromosomal organization of some of the honeybee desaturase and elongase genes supports the occurrence of duplication events during evolution. Four genes putatively encoding desaturases belonging to the First Desaturase subfamily, with higher affinity for stearic acid (C18:0) to form oleic acid (C18:1), were previously characterized in

A. mellifera (Hashimoto et al., 2008). In the current version of the honeybee genome (version 4.5) we could detect at least seven members of this subfamily. Computational and phylogenetic data including desaturase sequences displaying known functions were then performed to predict the functions of the honeybee desaturases. The premise was that, in a phylogenetic analysis of a family of enzymes encoded by multiple related genes from closely related species, the sequences will be clustered according to the function of enzyme/protein, or gene, or by the duplication history, but not by species (Roelofs and Rooney, 2003). In addition, the use of bootstrap confidence levels for assessing the accuracy of the phylogenetic tree allows inferring that members of a cluster of desaturases with a bootstrap support 90% would probably display the same function (Liu et al., 2004). Such analysis allowed us to classify the honeybee enzymes as being D9 (16 > 18) or D4 desaturases. Evidently, this needs experimental confirmation, especially because subtle differences between the sequences may imply different functions, as detected for the group of the D10,11 desaturases of lepidopterans (Liu et al., 2004). Our phylogenetic analysis of desaturases highlighted the following points: (1) the desaturases of some hymenopterans (Nasonia vitripennis and Harpegnathos saltator), a coleopteran (Tribolium castaneum) and a lepidopteran (D. plexippus) clustered together as D14 desaturases (desaturation at the position D14) (see Fig. 3, blue clade); (2) the desaturase encoded by the GB48193 gene is possibly a D4 desaturase since it groups with the D4 desaturases of Bombus terrestris (XP_003395145) and B. impatiens (XP_003492440), with a branch support of 0.963. The B. terrestris desaturase is an important enzyme for pheromones production (Bu cek et al., 2013); (3) one desaturase encoding gene, GB48195, shares an apparent orthology relationship with the D. melanogaster DesatF (see NP_651966 in Fig. 3), which creates a second unsaturation in the incipient fatty acid that will originate polyunsaturated CHCs. DesatF is expressed only in females and is involved in the synthesis of alkadienes, which are used as pheromones by these flies (Dallerac et al., 2000). Interestingly, the GB48195 gene is highly expressed in forager bees and showed a high positive correlation with the unsaturated alkenes and alkadienes. Further studies may inform whether this honeybee desaturase is femalespecific and involved in the biosynthesis of pheromones. Two elongase genes of the S/MUFA subfamily and four pertaining to the PUFA subfamily were previously identified in A. mellifera (Hashimoto et al., 2008). In the A. mellifera genome version 4.5 we found three S/MUFA members and eight PUFA members (see Fig. 4). Seven elongases that clustered into the S/ MUFA subfamily (GB53872, GB53873, GB45596) and into the PUFA subfamily (GB54396, GB51249, GB51247, GB54401), were mainly correlated to the production of alkenes and alkadienes, thus reinforcing the suggestion that they have roles in the elongation of monounsaturated or polyunsaturated CHCs. This premise was strengthened by the results of RT-qPCR transcript profiles that revealed that except for the genes GB53873 and GB54401, these elongase genes were highly expressed in the adult integument where they may contribute to the biosynthesis of unsaturated CHCs for the forager cuticle. In contrast, the elongases encoded by the genes GB54399 and GB40681 were mainly correlated to the production of methyl- and dimethyl-alkanes, but were clustered into the PUFA class. This apparently discrepant result deserves further investigation. 4.3. Expression of desaturase and elongase genes in the context of cuticle formation and maturation In the current work, our attention focused on genes putatively encoding two enzyme classes in the hydrocarbons biosynthetic

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pathways, desaturases and elongases, both determining fatty acid structure and function. The increased expression of five of the six desaturase genes and five of the ten elongase genes in newlyecdysed adults and/or forager bees coincided with the period of enlargement of the smooth endoplasmic reticulum in the oenocytes described by Hepburn et al. (1991). This structural change in the CHC-synthesizing cells suggests increasing synthesis of these compounds and strengthens our premise that the genes here analyzed have a role in CHCs biosynthesis. The increase in desaturase transcript levels in the integument of adult honeybees correlated positively with the increase in the proportion of most of the unsaturated compounds (alkenes and alkadienes). Although such correlation requires further experimental validation, it is important because it has the potential to inform us whether, and how strongly, these variables (levels of CHCs and transcripts) are interconnected. Together, the phylogenetic and the correlation analysis were relevant as the first steps toward the elucidation of the function of desaturase and elongase genes in the biosynthesis of CHCs of interest. A suitable way to advance in this direction consists in using RNA interference for silencing these desaturase or elongase genes, followed by CHC analysis in the silenced bees. In a minor fraction of our correlation data, we could detect a positive relationship between levels of desaturase transcripts and levels of saturated CHCs (alkanes and methyl-alkanes). At first glance, the positive correlation between these variables seemed merely casual, i.e., not representing causation effects, since the products of desaturase reactions are unsaturated, and not saturated compounds. However, saturated compounds are substrates for desaturases, and cell regulatory mechanisms may ensure substrate availability when the reaction product is needed for the organism. There are examples in biochemistry demonstrating that substrates may induce the transcription of enzymes-encoding genes, thus controlling the production of metabolites through biosynthetic pathways and the rate at which both substrates and enzymes are synthesized. This may tentatively explain why the expression of desaturase genes was positively correlated with the increase in the proportion of certain saturated compounds. Elongases catalyze the elongation of saturated and unsaturated fatty acids, however, a fraction of the CHC compounds, all of them being alkanes, methyl- or dimethyl-alkanes, showed a negative correlation with the expression of all the elongase genes herein characterized. Such genes may not have roles in the biosynthesis of these saturated CHCs. Interestingly, the elongase gene GB40681, which is significantly more expressed in the developing adult cuticle of the Pbm phase, was exclusively correlated with dimethylalkanes, coincidently present in higher proportions in the immature than in the mature cuticle. Similarly, the expression of the elongase gene GB54399 was exclusively correlated with methylalkanes, suggesting functional specificity. Thus, our correlative data points to strong causeeeffect relationships between the expression of desaturase/elongase genes and levels of CHCs thus suggesting that these genes have roles in the biosynthesis of these compounds. In conclusion, our data identified the hydrocarbon composition of the developing envelope layer, the less studied structural component of the exoskeleton, thus increasing our understanding of its formation and maturation. To our knowledge, this is the first approach on the changes in CHCs profiles in the immature cuticle during an entire molting cycle in comparison to the mature cuticle. Such analysis clearly demonstrated that different CHC classes predominate in immature and mature cuticles. In addition, correlation analysis allowed us to evaluate the strength of interconnection between the changing developmental patterns of CHCs and the fluctuations in the expression of genes potentially involved in CHCs

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biosynthesis. Additional insights into specific functions of these genes were provided by molecular phylogenetic analyses that also highlighted the evolutionary relationships between the desaturases and elongases of the honeybee and other insects. All this information is important for further studies on the molecular mechanisms leading to the diversity of CHC profiles and their developmental regulation. Acknowledgments This research was supported by FAPESP (São Paulo Research Foundation, Processes 2010/16380-9 and 2011/03171-5). Fellowships from CAPES and FAPESP (São Paulo Research Foundation, Process 2012/24284-4) were provided to T. Falcon. We are very grateful to Dr. D. G. Pinheiro for bioinformatics support and to Dr. N. Châline for helpful comments on the manuscript. We also thank L.R. Aguiar for his valuable technical assistance in the apiary. Appendix A. Supplementary data Supplementary data related to this article can be found at http:// dx.doi.org/10.1016/j.ibmb.2014.04.006. References Aitchison, J., 1982. The statistical analysis of compositional data. J. R. Stat. Soc. B 44, 139e177. Altschul, S.F., Gish, W., Miller, W., Myers, E.W., Lipman, D.J., 1990. Basic local alignment search tool. J. Mol. Biol. 215, 403e410. Le, S.Q., Gascuel, O., 2008. An improved general amino acid replacement matrix. Mol. Biol. Evol. 25, 1307e1320. Anisimova, M., Gascuel, O., 2006. Approximate likelihood-ratio test for branches: a fast, accurate, and powerful alternative. Syst. Biol. 55, 539e552. Armold, M.T., Regnier, F.E., 1975. Stimulation of hydrocarbon biosynthesis br ecdysterone in the flesh fly Sarcophaga bullata. J. Insect Physiol. 21, 1827e1833. Ausubel, F., Brent, R., Kingston, R.E., Moore, D.D., Seidman, J.G., Smith, J.A., Struhl, K., 1995. Short Protocols in Molecular Biology. John Wiley & Sons, New York. Billeter, J.C., Atallah, J., Krupp, J.J., Millar, J.G., Levine, J.D., 2009. Specialized cells tag sexual and species identity in Drosophila melanogaster. Nature 461, 987e991. Bitondi, M.M., Mora, I.M., Simões, Z.L.P., Figueiredo, V.L., 1998. The Apis mellifera pupal melanization program is affected by treatment with a juvenile hormone analogue. J. Insect Physiol. 44, 499e507. Blomquist, G.J., Dillwith, J.W., 1985. Cuticular lipids. In: Kerkut, G.A., Gilbert, L.I. (Eds.), Comprehensive Insect Physiology, Biochemistry and Pharmacology, Integument, Respiration and Circulation, vol. 3. Pergamon, Oxford, pp. 117e154. Blomquist, G.J., Chu, A.J., Remaley, S., 1980a. Biosynthesis of wax in the honeybee, Apis mellofera L. Insect Biochem 10, 313e321. Blomquist, G.J., Howard, R.W., McDaniel, C.A., Remaley, S., Dwyer, L.A., Nelson, D.R., 1980b. Application of methoxymercuration-demercuration followed by mass spectrometry as a convenient microanalytical technique for double-bond location in insect-derived alkenes. J. Chem. Ecol. 6, 257e269. Blomquist, G.J., Jurenka, R., Schal, C., Tittiger, C., 2012. Pheromone Production: biochemistry and molecular biology. In: Gilbert, L.I. (Ed.), Insect Endocrinology. Elsevier, New York, pp. 523e567. Bu cek, A., Vogel, H., Matousková, P., Prchalová, D., Zácek, P., Vrkoslav, V., Sebesta, P., Svatos, A., Jahn, U., Valterová, I., Pichová, I., 2013. The role of desaturases in the biosynthesis of marking pheromones in bumblebee males. Insect Biochem. Mol. Biol. 43, 724e731. Carlson, D.A., 1989. Dimethyl disulfide derivatives of long chain alkenes, alkadienes, and alkatrienes for gas chromatography/mass spectrometry. Anal. Chem. 61, 1564e1571. Châline, N., Sandoz, J.C., Martin, S.J., Ratnieks, F.L.W., Jones, G.R., 2005. Learning and discriminating of individual cuticular hydrocarbons by honeybees (Apis mellifera). Chem. Senses 30, 327e335. Chertemps, T., Duportets, L., Labeur, C., Ueyama, M., Wicker-Thomas, C., 2006. A female-specific desaturase gene responsible for diene hydrocarbon biosynthesis and courtship behaviour in Drosophila melanogaster. Insect Mol. Biol. 15, 465e473. Chino, H., Kitazawa, K., 1981. Diacylglycerol-carrying lipoprotein of hemolymph of the locustand some insects. J. Lipid Res. 22, 1042e1052. Chino, H., Katase, H., Downer, R.G., Takahashi, K., 1981. Diacylglycerol-carrying lipoprotein of hemolymph of the American cockroach: purification, characterization, and function. J. Lipid. Res. 22, 7e15. Dallerac, R., Labeur, C., Jallon, J.M., Knipple, D.C., Roelofs, W.L., Wicker-Thomas, C., 2000. 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. U. S. A 97, 9449e9454.

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Please cite this article in press as: Falcón, T., et al., Exoskeleton formation in Apis mellifera: Cuticular hydrocarbons profiles and expression of desaturase and elongase genes during pupal and adult development, Insect Biochemistry and Molecular Biology (2014), http://dx.doi.org/ 10.1016/j.ibmb.2014.04.006

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