EST and microarray analysis of horn development in Onthophagus beetles
© Kijimoto et al; licensee BioMed Central Ltd. 2009
Received: 23 March 2009
Accepted: 30 October 2009
Published: 30 October 2009
The origin of novel traits and their subsequent diversification represent central themes in evo-devo and evolutionary ecology. Here we explore the genetic and genomic basis of a class of traits that is both novel and highly diverse, in a group of organisms that is ecologically complex and experimentally tractable: horned beetles.
We developed two high quality, normalized cDNA libraries for larval and pupal Onthophagus taurus and sequenced 3,488 ESTs that assembled into 451 contigs and 2,330 singletons. We present the annotation and a comparative analysis of the conservation of the sequences. Microarrays developed from the combined libraries were then used to contrast the transcriptome of developing primordia of head horns, prothoracic horns, and legs. Our experiments identify a first comprehensive list of candidate genes for the evolution and diversification of beetle horns. We find that developing horns and legs show many similarities as well as important differences in their transcription profiles, suggesting that the origin of horns was mediated partly, but not entirely, by the recruitment of genes involved in the formation of more traditional appendages such as legs. Furthermore, we find that horns developing from the head and prothorax differ in their transcription profiles to a degree that suggests that head and prothoracic horns are not serial homologs, but instead may have evolved independently from each other.
We have laid the foundation for a systematic analysis of the genetic basis of horned beetle development and diversification with the potential to contribute significantly to several major frontiers in evolutionary developmental biology.
The origin of novel traits and their subsequent diversification have been central themes in evolutionary biology ever since the discipline's inception over 150 years ago [1, 2]. Specifically, the genetic, developmental, and ecological mechanisms, and the interactions between them, that allow novel phenotypes and functions to arise from pre-existing variation, continue to represent major frontiers in our understanding of phenotypic diversity. With the advent of modern -omics approaches, researchers have increasingly departed from a candidate gene or pathway approach and begun to explore organismal development and evolution from a genome, transcriptome, or proteome perspective, focusing in large part on existing genetic model systems such as Drosophila or Caenorhabditis. However, many key questions in evolutionary biology, including the mechanisms underlying organismal innovation, the role of plasticity in diversification, and the interplay between ecology and developmental evolution, are often difficult to address solely within the confines of classic model systems. Recent efforts have therefore begun to generate genomic and developmental genetic resources for organisms with promise as future model systems in evolutionary developmental biology and ecological genetics (e.g. butterflies: [3, 4]; honey bees: reviewed in ; red flour beetle: ). Here we present and apply the first genomic resources to advance the study of a class of traits that is both novel and highly diverse in a group of organisms that is ecologically complex and experimentally tractable: beetle horns and horned beetles.
Beetle horns possess many characteristics that make them interesting models for integrating genetic, developmental, and environmental perspectives on the development and evolution of complex, novel traits (reviewed in ). First, beetle horns are major structures, often dominating the phenotype of their bearers. Second, beetle horns function as weapons of sexual selection, thus playing a major role in the behavioral ecology of individuals and populations. Third, beetle horns are inordinately variable, both within and between species, including differences in number, size, shape, and location. Moreover, diversity in horn expression is paralleled by amazing species richness. For instance, the genus Onthophagus currently contains over 2,400 extant species, making it the most speciose genus in the animal kingdom [8, 9]. Forth, beetle horns are influenced in their expression by both genetic and environmental factors, ranging from absence of environmental sensitivity to complete determination by nutritional condition. In some cases, both extremes of environmental sensitivity can be found in different horn types expressed by the same individual . Finally, beetle horns lack any obvious homology to structures in other insects or non-insect arthropods. Beetle horns are not modified antennae or mouthparts, but instead horns were "invented" by beetles in addition to their traditional appendages , and now provide their bearers with an important new function: a weapon used in male-male competition. Beetle horns and horned beetles therefore offer rich opportunities to explore the mechanisms of organismal innovation and diversification.
Beetle horns are rigid outgrowths of the exoskeleton that originate as epidermal outbuddings of the head or prothoracic epithelium. Horns lack joints, muscles, and nervous tissue. Several recent studies have begun to shed light on how beetle horns develop and differentiate during ontogeny [11–15] and showed that two developmentally dissociated processes contribute to the final degree of horn expression seen in adults: a prepupal growth phase late in larval development followed by a pupal remodeling phase just prior to the final adult molt (reviewed in ). As such, the development of horns shows many qualitative similarities to the development of traditional appendages, but also exhibits important differences. For instance, prothoracic horn primordia are frequently resorbed during the pupal stage in a sex- and species-specific manner, a phenomenon not usually associated with regular appendages . Furthermore, earlier studies have also begun to question whether horns that develop in different body regions, such as the head vs. prothorax, constitute serial homologs, or instead may have evolved and diversified independently of each other [13, 16].
Here we present the first steps toward a systematic analysis of the genetic and genomic basis of horn development and diversification in the genus Onthophagus. We first present the results of a comprehensive EST analysis of two normalized cDNA libraries obtained from two disparate developmental stages of Onthophagus taurus: larva and pupa. Second, using microarrays developed from our EST libraries we contrast the transcription profiles of the primordia of developing prothoracic horns, head horns, and legs right after the transition from larva to pupa. We then use these contrasts to identify candidate genes involved in the development and diversification of beetle horns. Furthermore, we examine two basic questions regarding the origin and diversification of horns. (a) Are horns highly simplified versions of more traditional appendages such as legs? If so, transcription profiles of developing horn primordia should largely match those of developing legs. If not, transcription profiles of developing horn primordia should only partly match those of developing legs and also include horn specific transcription signatures. (b) Are different horn types produced in different body regions homonomous, i.e. serial homologs of the same ancestral structure? If so, different horn types should exhibit highly similar transcription profiles. However, if different horn types originated and diversified independently of each other, transcription profiles may be predicted to exhibit important horn-type specific differences. We discuss the significance of our findings in the context of the biology of horned beetles in particular, and the origins and diversification of novel traits in general.
Production and analysis of EST sequences
Summary of cDNA libraries and EST sequence analysis
(OtL & OtP)
median cDNA fragment size (nt)
average read length (nt)
It is likely that some of the non-redundant sequences derive from the same transcript but do not overlap, possibly due to 5'-truncated cDNA clones. In order to estimate the magnitude of this redundancy, we aligned the Onthophagus non-redundant sequences to Drosophila proteins, filtered the alignments for highly similar matches (BLASTx, E-value < 10-60), and then manually examined the alignments for separate Onthophagus sequences that align to distinct regions of the same Drosophila protein. Among 534 non-redundant Onthophagus sequences we found 35 pairs of sequences that aligned to the same Drosophila protein. Of these, 19 pairs aligned with highly similar matches to different regions of the same protein, indicating they derive from non-overlapping regions of the same transcript; 12 pairs had co-linear alignments with 95-98% sequence identities, suggesting that they either derive from the same gene with polymorphisms and/or sequencing errors, or derive from highly similar duplicate genes; and 4 pairs appear to be splice variants. Thus, this sample of 534 Onthophagus non-redundant sequences represents approximately 499 distinct genes (93% unique). While this is not a random sample and thus can't be extrapolated to full set of non-redundant sequences, it does however indicate that false-negative assemblies are not a pervasive problem among the non-redundant sequences.
Functional annotation of assembled sequences
Given that insects express a broad diversity of genes during metamorphosis , we expected that the larval and pupal ESTs would be a rich source of gene discovery. In order to provide a first pass annotation for the putative function of the Onthophagus gene sequences, we annotated the non-redundant sequences using the UniProtKB/TrEMBL protein sequence database (E-value < 10-5). This successfully annotated 71.3% of the non-redundant sequences. As expected, these annotations covered a wide diversity of biological and molecular functions including the major expected categories such as cellular processes, metabolic processes, biological regulation, multicellular organismal processes, and developmental processes (see Additional files 1 and 2). This, coupled with the low redundancy within the Onthophagus libraries, indicates the set of ESTs as a rich source for gene discovery.
Putative Onthophagus taurus orthologs with known functions in insect development and physiology
Ot library ID
Autophagy-specific gene 12
Sptzle 1B (Spz1B)
held out wings
Map kinase-interacting serine/threonine kinase
Broad complex isoform Z1
Programmed cell death
Autophagy-specific gene 1
Juvenile hormone acid methyl transferase
Epidermal growth factor-like protein
Juvenile hormone epoxide hydrolase 1 (EC 3329)
Insulin receptor substrate 1 (IRS-1) (pp185)
Ecdysone 20-hydroxylase (EC 1149922)
Wnt oncogene analog 2
Mago nashi, putative
Enhancer of split protein, putative
Autophagy-specific gene 8a
forkhead box, sub-group O
Ras-related protein 2
Creb protein (Fragment)
Homeodomain transcription factor Prothoraxless
Sex comb on midleg
Ecdysone inducible protein 75 isoform B
absent, small, or homeotic discs 1
Autophagy-specific gene 8b
Insulin receptor tyrosine kinase substrate
ETS-like protein pointed, isoform P2 (D-ETS-2)
Discs large 1 tumor suppressor protein
Additional sex combs
Insulin-like peptide receptor precursor
Bambi (BMP and activin membrane-bound inhibitor)
Ecdysone-induced protein 78C
Comparative analysis of the Onthophagus transcriptome
Datasets used in this study.
Version (date uploaded, YYMMDD)
NCBI 36 release 46 (070803)
Wormpep 180 (070819)
Our analysis also identified 633 of the non-redundant sequences (23.2%) to have "no-hit" (Figure 1 Group 4) to any of the proteomes. This is consistent with the finding that approximately 23% of genes annotated in the Tribolium genome lack sequence matches in a wide range of other species . However, our estimate of Onthophagus specific sequences is likely to be inflated by (i) sequences that are largely, or entirely, within the UTRs of protein coding transcripts, or (ii) sequences that may be non-coding transcripts. Resolving the question of whether these sequences do in fact represent genes that are unique to Onthophagus must await large-scale sequencing of the transcriptomes and/or genomes of Onthophagus and related species. However, the observation that 44% of theses sequences include ORFs of greater than 300 nucleotides (data not shown), suggests that at least some of these represent protein-coding genes that have not yet been identified in the species sequenced to date.
Gene expression profiles in pupal appendage primordia
While our EST analysis identified many genes homologous to interesting Drosophila developmental genes, and such an approach to identify candidate genes has been successful in beetles [11, 15, 19, 20], this approach is limited to identifying obvious candidates. Given that Onthophagus horns appear to be novel structures invented in beetles, it is highly likely that unexpected, or indeed previously uncharacterized genes may be important in their development. We therefore developed a custom microarray spotted with the 3,756 cDNA clones from which the ESTs were derived (Methods), undertook gene expression profiling of developing horns (early pupal stage) as an unbiased means of identifying such candidates. Since there is evidence that head horns and prothoracic horn are quite distinct structures (not simply serial homologs; [13, 16]), we analyzed gene expression in each of these organs separately. Since there is evidence that some, but not all, appendage patterning genes play a role in horn development , we included legs in our analysis in order to distinguish similarities and differences between horns and a canonical appendage. Finally, since beetle horns and legs both develop by out-budding of the epithelium, we use non-appendage bearing epithelium (dorsal abdomen) as a common reference sample.
Identifying candidate genes based on expression in horn primordia
First, we identified genes whose expression in the context of horn development could be expected given existing insights into the developmental biology of horns, and knowledge about the function of these genes in other organisms. For instance, the Hox gene Sex combs reduced (Scr) is enriched in the prothoracic horn (19.2 fold) and legs (7.9 fold) relative to abdominal epithelium (Additional file 4). In Drosophila and Tribolium Scr patterns the identity of the labial and first thoracic segment [21–23]. Preliminary results showed that Onthophagus Scr executes similar functions during labial and thoracic development in addition to playing a major role in the regulation of prothoracic horn development (Wasik, Rose, and Moczek, unpublished data).
Secondly, we identified genes that although functionally well characterized in Drosophila or elsewhere, would not readily be expected to be expressed in the context of horn development. Genes in this category include the putative ortholog of Drosophila doublesex (dsx), enriched more than 2-fold in the head and prothoracic horns relative to abdominal epithelium (Additional file 4). In Drosophila the expression of sex-specific DSX isoforms regulate somatic sex-determination sexually dimorphic differentiation [24, 25]. While Onthophagus horns are sexually dimorphic, our observation that the putative dsx ortholog is expressed preferentially in the male horn tissue when compared to male abdominal tissue was unexpected. Expression and functional studies are now under way to identify the role of dsx in the development and diversification of horns.
Similarly, we found that the putative Onthophagus orthologs of yellow-c, -e, and -f were enriched more than 2-fold in head and/or prothoracic horns relative to abdominal epithelium (Additional file 4). The functions of yellow family genes are remarkably diverse and include the regulation of pigmentation [26, 27], the production of a major component of royal jelly in the honeybee  as well as expression of normal male courtship behavior in Drosophila . Combined, these observations suggest that yellow genes may be involved in the regulation of a wide array of sex- or caste-specific functions, at least among insects, though it remains to be determined, what, if any, function the gene family may be executing in Onthophagus beetles.
Lastly, we identified 74 genes that were significantly differentially expressed in either head horns or prothoracic horns, or both, that lack obvious homology to proteins in any of the datasets used in this study. Of those 74, at least 29 (39%) contained predicted ORFs with longer than 300 nucleotides (100 codons).
Horned beetles, most notably in the genus Onthophagus, are increasingly being recognized as an emerging model system in evo-devo and eco-devo studies [13, 30–34]. Below we discuss the major findings of our study and their applicability to ongoing and future research efforts in horned beetles and beyond.
Onthophagus taurus expressed sequences as a resource
The expressed sequences and the corresponding cDNAs presented here provide a valuable entry point for studies of gene function in Onthophagus taurus. The sequences derived from normalized larval and pupal cDNA libraries had a low level of redundancy. The 3,488 high quality EST sequences from both libraries assembled into 2,781 non-redundant sequences (contigs and singletons). The low level of redundancy resulted in a sample of sequences derived from a wide range of biological functions.
The Onthophagus transcriptome
This study provides a first pass survey of genes found in Onthophagus. Prior to this study, Tribolium castaneum was the only species of beetle for which comprehensive sequence information was available [6, 35]. Comparative analyses indicate that the gene repertoire of Tribolium is consistent with the general trends seen across sequenced insects and vertebrates [6, 36]. Our estimates of the proportions of Onthophagus sequences that are common to other species are consistent with those in Tribolium . For instance, we found that 39% of Onthophagus sequences had sequence matches to proteins in all the datasets searched, which is consistent with the Tribolium genome in which ~35% of genes have orthologs in all species examined . Of particular interest are the 23% of Onthophagus sequences that lack orthology (Group 4 in Figure 1) to proteins from six proteomes including the non-redundant dataset which is very close to the corresponding estimate of 23% of annotated Tribolium genes . About 40% of these Group 4 Onthophagus sequences exhibited appreciable putative open reading frames and thus need to be considered potentially protein-coding. This group of genes likely contains genes unique to, or fast evolving in, Onthophagus beetles, and studies are under way to further characterize and analyze the significance of these genes for the evolution, diversification, and radiation of horned beetles.
From ESTs to candidate genes for the evolutionary biology of beetle horns and horned beetles
Beetle horns and horned beetles are attractive system to address several current frontiers in evolutionary biology. The EST resources and array results presented here provide the first genomic resources to identify candidate genes, pathways, and networks underlying morphological, behavioral, and developmental aspects of the biology of horned beetles, as well as providing insights into their respective evolutionary histories. Below we briefly highlight two broad categories of current research efforts and how they are being advanced by the results presented here.
The origins of horns
Beetle horns have attracted attention because they lack obvious homology to other appendages or outgrowths in the insects. Horns therefore constitute an evolutionary novelty. Understanding how novel traits arise from pre-existing variation remains one of the most challenging and poorly understood questions in evolutionary biology.
One hypothesis that has been proposed toward explaining the origin of horns is based on the observation that horns share many morphological and developmental features with traditional appendages (e.g. epithelial origin, prepupal growth, dorso-ventral axis formation, or pupal remodeling presumably via programmed cell death; ). Furthermore, in several other respects horns are much simpler than legs or mouthparts (e.g. they lack nerves, muscles, or joints). Horns may therefore have evolved via the large-scale co-option of genes ancestrally used to instruct appendage development. Our microarray results suggest that horns and legs are indeed highly similar in gene expression profiles and support the hypothesis that many genes involved in leg formation may also play a role in horn development. Earlier research has begun to implicate a small subset of appendage patterning genes in horn development (Distal-less, dachshund, extradenticle, homothorax, [11, 13, 15]). The results presented here add a substantial list of gene candidates (Additional file 4) that may have mediated the origin of horns via co-option from traditional appendage development.
At the same time, horn-specific transcription profiles also included genes not represented in developing legs, suggesting that horns should not be viewed solely as being simplified appendages. While this fraction of genes was small in comparison, it nevertheless highlights a possible class of genes involved in developmental processes of horn formation that are not represented, or at least not to the same degree, during the development of traditional appendages. If correct, this would suggest that the origin of horns may have been mediated by the co-option of appendage patterning genes alongside integration of genes and pathways unrelated to appendage formation. Clearly, additional contrasts including the sampling of other developmental time points, as well as gene function studies, are needed to establish the general validity of these conclusions.
The diversification of beetle horns and horned beetles
Beetle horns and horned beetles are attractive study organisms because they permit investigation of the mechanisms underlying phenotypic diversification on many interesting levels. First, species differ in the body region involved in horn expression: horns may extend from the head, prothorax, or both, and while their function as weapons in male combat appears to be conserved across species, recent studies suggest that different horn types may have originated and diversified at least in part independently of one another . Our results support this scenario by identifying a list of genes whose expression differs significantly across horn types such as yellow-e (head horns), tailup (encodes a LIM-homeodomain protein; prothoracic horns), or Scr (prothoracic horns and legs). While the function, if any, of these candidate genes in the context of horn development remains to be explored our results presented here provide an important starting point toward untangling shared, independent, and convergent aspects in the evolution of different horn types across horned beetles.
Substantial diversity in horn expression also exists within species in the form of sexual and male dimorphisms. Sexual dimorphisms are brought about via sex-specific regulation of horn expression whereas male dimorphisms are predominantly the product of nutritional differences experienced during larval life (reviewed in ). Endocrine factors such as juvenile hormone (JH) are likely to play important roles in the regulation of both types of diversity [38–40]. Furthermore, the same nutritional or hormonal manipulations affect sexual and male dimorphisms differently in different species and populations, suggesting that evolutionary changes in the interplay between endocrine factors, nutrition, and sexual differentiation have contributed to the diversification of horned beetles [40, 41]. Our EST resources and microarray results provide an important starting point to begin exploring putative candidate genes that may be associated with sex-specific (such as doublesex, transformer-2 or members of the yellow gene family) or nutrition-dependent (e.g. foxo) expression of horns. Moreover, the resources presented here should support the development of experiments towards characterizing sex- and morph-specific transcriptomes in O. taurus and closely related species in the genus (Snell-Rood, Cash, Kijimoto, Andrews, Moczek; unpublished data).
In conclusion, the EST resources and microarray results present here provide a first step toward a systematic analysis of the molecular basis of horn development and diversification in beetles with the potential to inform several major frontiers in evolutionary developmental biology.
cDNA library construction
Adult Onthophagus taurus were collected from pastures near Bloomington, IN and reared as described previously . We constructed two separate libraries from larval and pupal stages. For the larval library we dissected heads and thoraces from mid third instar larvae, late third instar larvae, and early and late prepupal stages. For the pupal library tissues included whole individuals one, two, three and four days after pupation. For both libraries we harvested at least two individuals for each stage and sex, and all samples were frozen in liquid nitrogen, immediately transferred to -80°C for storage until RNA extraction. Total RNA was extracted using TRIreagent (Sigma, MO), precipitated with ethanol and stored at -80°C. The normalized cDNA libraries were each constructed from 1 μg of total RNA, using the TRIMMER-DIRECT cDNA normalization kit (Evrogen, Moscow, Russia) for the library normalization, followed by the Creator SMART cDNA library construction kit (Clontech, CA) for cDNA library construction, as described in Zhulidov et al. 2004 . We followed the manufacturers protocols with the following modifications and specific conditions: (i) the cycle conditions for the PCR-based double-strand cDNA synthesis were 16 cycles of [95°C for 7 sec, 66°C for 30 sec, and 72°C for 6 min]; (ii) we used 2 μl of cDNA mixture for PCR during cDNA library construction and normalization; and (iii) the conditions for the two step amplification of the normalized cDNA were 18 cycles [95°C for 7 sec, 66°C for 30 sec, and 72°C for 6 min] for the first step, and the second amplification was cycled for 12 cycles using the same conditions. Normalized and amplified cDNA fragments were size-fractionated, digested by Sfi I, and ligated with the plasmid vector pDNR-LIB according to manufacturer's instruction. Electro-transformed E. coli cells were spread on LB plate containing chloramphenicol (final concentration of 30 μg/ml). The estimated titer of both of the libraries were ~1 × 10-8CFU. A total of 3,756 colonies were picked at random. Unless stated otherwise standard molecular procedures were used to execute basic molecular analyses .
DNA samples were prepared for sequencing using a Beckman Coulter Biomek FX Laboratory Automation Workstation as described in Burr et al. 2006 . Each picked clone was incubated overnight at 37°C in 96-well tissue culture plates with 100 μl of SOC medium with chloramphenicol (final concentration of 30 μg/ml), without rotation. 20 μl of the cultured cells were mixed with 80 μl of water and heat-punctured at 95°C for 10 min. Insert DNA was PCR-amplified using cell lysate (10 μl) as template, 0.1 μM M13fw primer (5'-GTG TAA AAC GAC GGC CAG TAG-3'), 0.1 μM M13rev primer (5'-AAA CAG CTA TGA CCA TGT TCA C-3'), 0.2 mM each dNTP, 0.5 U/20 μl reaction Taq polymerase (Bioline, MA), and 1× reaction buffer (Bioline, MA). The reaction was incubated at 95°C for 5 min then 35 cycles of [95°C for 1 min, 54°C for 1 min, and 2 min at 72°C]. The amplified DNA was purified using the Multiscreen-PCR 96-well purification system (Millipore, MA). The purified DNA was subjected to agarose gel electrophoresis against molecular weight standard and visualized using a Kodak 440cf imaging station. Sequencing reactions were performed with the primer pDNRlib30-50 (5'-TAT ACG AAG TTA TCA GTC GAC G-3') and ABI BigDye chemistry and ABI Prism 3730 sequencer (Applied BioSystems, CA).
EST processing, assembly, and annotation
ESTPiper  was used to analyze EST sequences including base calling, data cleaning, de novo assembly, and annotation. A total of 3,756 EST sequences were generated in FASTA format with quality scores after base calling. For data cleaning, ESTPiper first removed low quality and vector sequences using LUCY  program with the default parameter settings. PolyA/T tails were then trimmed, where within 50 bp searching range from both ends of the sequences, the minimum length of continuous polyA/T region was set to 9 bp and the maximum number of mismatches within the polyA/T region was set to 3. Potentially chimeric clones, which were defined as sequences with at least 30 bp continuous A/T or adaptors occurring in the middle of sequences, were removed. Finally, shorter sequences (< 100 bp) were also removed. A total of 3,488 high quality sequences passed data cleaning procedure. We then performed de novo assembly to assemble EST sequences into contigs and singletons. Parameters were set as follows: (i) overlap percent identity cutoff was 95%, (ii) overlap length cutoff was 49, and (iii) maximum number of word matches was 10,000 (this parameter defines the maximum number of matches that the program will consider for a given sequence, and was set high to improve accuracy . For annotation, ESTPiper matched contigs/singletons to UniProt database  using BLASTX with an E-value cutoff of 1 × 10-5 and only the top match was taken.
We developed the cDNA microarray using all the clones used for the EST analysis (3,756 clones) as well as GAPDH and actin-5c (internal positive controls). Insert DNA was PCR amplified and purified as described above in the EST sequencing section. We followed the protocol of Indiana University Drosophila Genomics Resource Center  to print microarrays with a minor revision to post-print washes. Purified insert DNA was dried completely, re-dissolved in DGRC spotting solution (1.5 M Betaine in 3 × SSC), and spotted to GAPSII Microarray Slides (Corning) using an OmniGrid 300 printing. The microarray design included 4,320 spots arranged in 48 blocks of 90. A total of 3,756 of these spots were cDNA fragments (each spotted once) and 564 of these consisted of control spots (GAPDH, actin-5c, and spotting buffer only). The gene list and platform description is available at Gene Expression Omnibus http://www.ncbi.nlm.nih.gov/geo/ accession number GPL7555. After printing, the microarrays were heated at 85°C for 3 hrs and rinsed with 5 × SSC/0.1%SDS (55°C), water (twice at RT, once at 95°C, and once again at RT) and then centrifuged to dry. All microarrays were kept dry at room temperature until they were used.
Target RNA preparation, hybridization and obtaining data sets
Tissues were dissected from 20 male O. taurus (day 1 pupae) that were collected from our laboratory colony. Dissections and RNA extractions (RNeasy Mini kit, Qiagen, CA) of head horn, prothoracic horn, leg, and abdominal epithelium were performed separately for each animal. Independent biological replicates of RNA samples were created by pooling an equal mass of RNA isolated from the same type of tissue from 4 individuals. For each RNA sample 1 μg of RNA was reverse transcribed using Oligo(dT)-T7 primer (Ambion, TX) and SuperScriptIII reverse transcriptase (Invitrogen, CA), and DNA polymerase and RNase H (Invitrogen, CA) were used for second strand synthesis. Amplified RNA (aRNA) was generated by in vitro transcribing the cDNA using the MEGAscript kit (Ambion, TX). The aRNA was directly labeled with Cy3 or Cy5 using the ULS aRNA Fluorescent Labeling Kit (KREATECH, Amsterdam, The Netherlands). Three sets of amplified RNA samples from head horns, prothoracic horns, and legs were labeled with Cy5, while abdominal epithelial tissue samples were labeled with Cy3. The remaining two sets of samples were labeled in the opposite way. After measuring the quantity and labeling efficiency, amplified and labeled RNA samples from test (head horns, prothoracic horns, and legs) samples and abdomen (reference sample) were mixed and hybridized onto arrays. aRNA with 50 pmol dye from the test sample and reference sample were mixed with KREAblock (ULS aRNA Fluorescent Labeling kit) and 2 × enhanced cDNA hybridization buffer (Genisphere, PA), then heated at 80°C for 10 min. Arrays were pre-treated for more than one hour at 55°C in pre-hybridization buffer (5 × SSC, 0.1%SDS, 1% I-block (Applied Biosystems, CA)). Both mixed sample and microarray were kept at 55°C until the hybridization step. Hybridization was performed in a dark humidified chamber at 55°C overnight. The microarray was rinsed in buffer A (2 × SSC/0.2%SDS) at 55°C then incubated in buffer A at 65°C for 10 min. The microarray was transferred to 2 × SSC (room temperature) for 10 min, followed by incubation in 0.2% SSC for 10 min at room temperature. The rinsed microarray was dried by centrifuging at 500rcf for 4 min.
The hybridized microarrays were scanned by GenePix scanner 4200 (Molecular Devices, CA) to obtain raw data sets. After initial quality check of results using OLIN in Bioconducter (Basic Hybridization Analysis, Costello et al. 2005, https://dgrc.cgb.indiana.edu/microarrays/support/bha.html), differential expression was assessed using Limma . The values for each spot were shown as log2 ratios between the two signal intensities (M-values). The microarray data are available at Gene Expression Omnibus http://www.ncbi.nlm.nih.gov/geo/, accession number GPL7555.
We performed clustering analysis and support tree construction using TIGR MultiExperiment Viewer of the TM4 system . We performed hierarchical clustering by using Cosine Correlation with average linkage to obtain the cluster and tree.
This manuscript benefited greatly from comments by Amy Cash, Emilie Snell-Rood, and two anonymous reviewers. We thank the Center for Genomics and Bioinformatics at Indiana University and its staff, especially John Colborne, Zhao Lai, Zach Smith, Jade Buchanan-Carter, and Heejung Yang for their advice and expertise in executing this study. Erin Yoder and Sarah Jones provided expert beetle care. Funding for this study, as well as for work in the Center for Genomics and Bioinformatics, was provided in part by the METACyt Initiative of Indiana University, funded in part through a major grant from the Lilly Endowment. Additional support was provided by National Science Foundation grants IOS 0820411 to JA and APM.
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