The mitochondrial genome of Elodia flavipalpis Aldrich (Diptera: Tachinidae) and the evolutionary timescale of Tachinid flies.

Tachinid flies are natural enemies of many lepidopteran and coleopteran pests of forests, crops, and fruit trees. In order to address the lack of genetic data in this economically important group, we sequenced the complete mitochondrial genome of the Palaearctic tachinid fly Elodia flavipalpis Aldrich, 1933. Usually found in Northern China and Japan, this species is one of the primary natural enemies of the leaf-roller moths (Tortricidae), which are major pests of various fruit trees. The 14,932-bp mitochondrial genome was typical of Diptera, with 13 protein-coding genes, 22 tRNA genes, and 2 rRNA genes. However, its control region is only 105 bp in length, which is the shortest found so far in flies. In order to estimate dipteran evolutionary relationships, we conducted a phylogenetic analysis of 58 mitochondrial genomes from 23 families. Maximum-likelihood and Bayesian methods supported the monophyly of both Tachinidae and superfamily Oestroidea. Within the subsection Calyptratae, Muscidae was inferred as the sister group to Oestroidea. Within Oestroidea, Calliphoridae and Sarcophagidae formed a sister clade to Oestridae and Tachinidae. Using a Bayesian relaxed clock calibrated with fossil data, we estimated that Tachinidae originated in the middle Eocene.

Introduction

Since the first insect mitochondrial genome (mitogenome) sequence was reported by Clary and Wolstenholme in 1985 [1], Diptera has remained the primary model system for mitogenomic research. This has included such diverse topics as species identification [2], [3], molecular evolution and phylogenetic inference [4]–[7], population structure and phylogeography [8]–[12], and genome structure and rearrangement [13]–[19]. Because of its small size and relative ease of sequencing, the number of mitogenome sequences has grown rapidly. As of January 2013, there are 64 complete or near-complete dipteran mitogenome sequences in GenBank, accounting for about 17.5% of the 365 insect mitogenomes that have been sequenced. In addition to the model organism Drosophila, most studies of Diptera have focused on taxa of medical and economic importance, such as the anopheline mosquitoes (Culicidae), which are vectors of malaria [20], [21]; the fruit flies Ceratitis capitata and Bactrocera spp. (Tephritidae), which are serious agricultural pests [11], [22]; the blowflies (Calliphoridae) and oestrid flies (Oestridae), which can cause myiasis [23], [24]; and leaf-miners (Agromyzidae), which are vegetable and horticultural pests [16], [25].

Tachinid flies have a worldwide distribution and comprise nearly 10,000 described species [29]. Despite Tachinidae being the second-largest dipteran family, the mitogenomes of only two species have been sequenced completely: Exorista sorbillans (Exoristinae, Exoristini) and Rutilia goerlingiana (Dexiinae; Rutiliini) [26], [27], [28]. They are natural enemies of many lepidopteran and coleopteran pests of forests, agricultural crops, and fruit trees, and thus are of economic importance. The Palaearctic tachinid fly, Elodia flavipalpis Aldrich, 1933 (Exoristinae, Goniini), is usually found in Northern China and Japan and is in the same subfamily, Exoristinae, as Ex. sorbillans [30], [31]. It is one of the primary natural enemies of the leaf-roller moths (Tortricidae), which are major pests of various fruit trees [32], [33]. The monophyly of Tachinidae is broadly supported by phylogenetic studies, but questions remain about its place in the superfamily Oestroidea, particularly the relationship between Tachinidae and several large families of Oestroidea [27], [28], [34]–[36]. There have been various studies of the divergence times of different groups of flies [7], [11], [12], [37], with a recent study placing the rapid radiation of Schizophora 65 mya in the Paleocene [28]. However, owing to the uncertain taxonomic position of Tachinidae in Oestroidea, the evolutionary timescale of tachinid flies has not been well studied.

Here we describe the complete mitochondrial genome of El. flavipalpis. The mitogenome contributes to discussion on the evolutionary relationships and taxonomic positions of the Tachinidae, puts forward a bold hypothesis regarding the timescale in which the family originated, and will aid further molecular research of related taxa through the findings on primer selection for atypical regions and the optimization of PCR experiments.

Materials and Methods

Specimen Collection, DNA Extraction, and DNA Amplification

Adult individuals of Elodia flavipalpis Aldrich, 1933 were collected directly from the pupae of their host species, the leaf-roller moth Spilonota lechriaspis Meyrick (Tortricidae). Moth pupae were collected at an organic apple orchard in Beijing, China, and hatched in the laboratory. The specimens were preserved in 99.5% ethanol and stored at −20°C for preservation of nucleic acids. DNA extraction from a single specimen was performed using the DNeasy Tissue kit (QIAGEN) following the manufacturer’s instructions.

The fragments were first amplified with the universal PCR primers from Simon et al. [38] and some dipteran-specific primers from Han [39] and Weigl et al. [24] (Table 1). Primer pairs for amplification of the mitochondrial control region were modified according to Lessinger et al. [40] and Oliveira et al. [41]. Species-specific primers were designed using Primer Premier 6.0 software [42], based on the initial fragments aligned with sequences from three closely related species, Ex. sorbillans, Calliphoridae spp., and Oestridae spp. (Table 2). PCR products covering the remaining regions of the mitogenome were amplified using universal and species-specific primers (Table 1). The entire genome of El. flavipalpis was amplified in 21 fragments. All of the primers were synthesized by Shanghai Sangon Bio-technology Co., Ltd (Beijing, China).

Table 1

Details of mitogenome sequencing protocols used in this study.

Region	Primer pairs (F/R)	Sequence (forward and reverse) 5′→3′	Size (bp)	PCR conditions (annealing/extension)
ND2	TM-J-206a/N2-N-732a	GCTAA-ATAAAGCTAACAGGTTCAT/GAAGTTTGGTTTAAACCTCC	∼500	49°C, 45 s/72°C, 1 min
ND2-CO1	N2-J-283d/C1-N-1740d	CATGACTAGGAACTTGAATAGG/AGAACTAAAGCAGGAGGTAA	∼1500	47°C, 45 s/68°C, 4 min
CO1	TY-J-1460a/C1-N-2191a	TACAATTTATCGCCTAAA-CTTCAGCC/CCCGGTAAAATTAAAATATAAACTTC	∼700	49°C, 45 s/72°C, 1 min
CO1	C1-J-1751a/TL2-N-3014a	GGATCACCTGATATAGCATTCCC/TCCAATGCACTAATCTGCCATATTA	∼1200	52°C, 30 s/72°C, 1 min
CO1-CO3	Cl-J-2183a/C3-N-5460a	CAACATTTATTTTGATTTTTTGG/TCAACAAAGTGTCAGTATCA	∼3200	44.5°C, 45 s/68°C, 5 min
ATP8	C2-J-3530d/A6-N-4493d	AAGTTGATGGAACTCCTGGA/GTAAGTCGAACTGCTAATGT	∼950	49°C, 45 s/72°C, 1 min, 50 s
CO3-ND3	C3-J-5005d/E-revb	CTCCATCAGTTGAATTAGGTGCTA/AGTGATAAGCCTCTTTTTGGCTTC	∼1050	49°C, 45 s/72°C, 1 min, 50 s
ND5	F-fwb/N5-N-7707d	CATTTGATTTGCATTCAAAAAGTATTG/AGGATGAGATGGATTAGGAT	∼1800	45°C, 45 s/68°C, 3 min
ND5-ND4	H-fwb/N4-N-8718a	GAAACAGGAGTAGGAGCTGCTATAGC/GCTTATTCATCGGTTGCTCA	∼1200	53°C, 45 s/72°C, 1 min, 50 s
ND4	I-fwb/N4-N-8924a	CAATTCTATTAATTAAAGAAATTTCTCC/AAAGCTCATGTTGAAGCTCC	∼550	48°C, 45 s/72°C, 1 min, 50 s
ND4	N4-J-8614d/N4-N-9061d	TGAGCAACAGAAGAATAAGC/ATCAACCAGAACGATTACAAG	∼400	47°C, 45 s/72°C, 1 min, 50 s
ND4-ND6	N4-J-8944a/I-revb	GGAGCTTCAACATGAGCTTT/CTTATTTTTGATTTACAAGACCAATG	∼850	49°C, 45 s/72°C, 1 min, 20 s
ND4- CYTB	N4-J-9511d/CB-N-11218d	CTAAAATTGATAACCCTAAAGC/TCAGGTTGAATGTGAATTGG	∼1700	45°C, 45 s/68°C, 3 min
CYTB-ND1	CB-J-10933a/N1-N-12051a	TATGTACTACCATGAGGACAAATATC/GATTTTGCTGAAGGTGAATCAGA	∼1100	50.5°C, 45 s/72°C, 1 min, 50 s
ND1- lrRNA	N1-J-11891d/16S-N-12855d	ATCCTCCTCTTCTATATTCAAT/GATTGCGACCTCGATGTT	∼950	48°C, 45 s/72°C, 1 min, 30 s
lrRNA	LR-J-12883c/LR-N-13398a	CTCCGGTTTGAACTCAGATC/CGCCTGTTTATCAAAAACAT	∼550	47°C, 45 s/72°C, 2 min
lrRNA - srRNA	LR-J-12888d/SR -N-14373d	ACGCTGTTATCCCTAAAGTA/AATCCACGATGAACCTTACT	∼1500	45°C, 45 s/68°C, 3 min
srRNA	SR-J-14233a/SR-N-14756a	AAGAGCGACGGGCGATGTGT/GACAAA-ATTCGT-GCCAGCAGT	∼500	50°C, 45 s/72°C, 1 min, 30 s
srRNA	SR-J-14612a/SR-N-14922a	AGGGTATCTAATCCTAGTTT/AAGTTTTATT-TTGGCTTA	∼300	43°C, 45 s/72°C, 1 min
srRNA-ND2	SR-J-14646e/N2-N-757d	GCTGGCACAAATTAAATC/GCTGCAAGTATTCAACTTAAATG	∼1150	43°C, 1 min/68°C, 6 min
Control region	SR-J-14646e/N2-N-309f	GCTGGCACAAATTAAATC/CTAAACCTATTCAAGTTCC	∼650	42°C, 1 min/68°C, 6 min

Note: CO1, CO2, CO3: cytochrome c oxidase subunit 1, 2, and 3 genes; CYTB: cytochrome b gene; ATP6, ATP8: ATP synthase subunit 6 and 8 genes; ND1, ND2, ND3, ND4, ND4L, ND5, ND6: NADH dehydrogenase subunit 1–6 and 4L genes. lrRNA, srRNA: large and small ribosomal RNA.

Primers from Simon et al. [27].

Primers from Weigl et al. [21].

Primers from Han [28].

Primers designed specially for this genome, using the nomenclature of Simon et al. [27].

Primers modified from Lessinger et al. [29]. f Primers modified from Oliveira et al. [30].

Table 2

Summary of mitogenome sequences from Diptera and an outgroup species from Lepidoptera.

Species	Family	Length (bp)	Accession Number	Reference
Elodia flavipalpis	Tachinidae	14932	NC_018118	Present study
Exorista sorbillans	Tachinidae	14960	NC_014704	Shao et al., 2012
Dermatobia hominis	Oestridae	16360	NC_006378	Azeredo-Espin et al.,2004
Hypoderma lineatum	Oestridae	16354	NC_013932	Weigl et al., 2010
Chrysomya putoria	Calliphoridae	15837	NC_002697	Junqueira et al., 2004
Cochliomyia hominivorax	Calliphoridae	16022	NC_002660	Lessinger et al., 2000
Lucilia sericata	Calliphoridae	15939	NC_009733	Stevens et al., 2008
Sarcophaga impatiens	Sarcophagidae	15169	NC_017605	Nelson et al., 2012
Haematobia irritans	Muscidae	16078	NC_007102	Oliveira et al., 2008
Drosophila littoralis	Drosophilidae	16017	NC_011596	Sorokina et al., 2010
Drosophila sechellia	Drosophilidae	14950	NC_005780	Ballard, 2000 a
Drosophila simulans	Drosophilidae	14927	NC_005781	Ballard, 2000 a
Drosophila mauritiana	Drosophilidae	14964	NC_005779	Ballard, 2000 b
Drosophila melanogaster	Drosophilidae	19517	NC_001709	Lewis et al., 1995
Drosophila yakuba	Drosophilidae	16019	NC_001322	Clary & Woolstenholme, 1985
Drosophila pseudoobscura	Drosophilidae	14914	NC_018348	Torres et al., 2009
Liriomyza sativae	Agromyzidae	15551	NC_015926	Yang et al., 2011
Liriomyza trifolii	Agromyzidae	16141	NC_014283	Wang, 2010
Liriomyza bryoniae	Agromyzidae	16183	NC_016713	Yang et al. unpublished
Liriomyza huidobrensis	Agromyzidae	16236	NC_016716	Yang et al. unpublished
Fergusonina taylori	Fergusoninidae	16000	NC_016865	Nelson et al., 2011
Ceratitis capitata	Tephritidae	15980	NC_000857	Spanos et al. 2000
Bactrocera carambolae	Tephritidae	15915	NC_009772	Ye et al., 2010
Bactrocera dorsalis	Tephritidae	15915	NC_008748	Yu et al., 2007
Bactrocera minax	Tephritidae	16043	NC_014402	Zhang et al.,2010
Bactrocera oleae	Tephritidae	15815	NC_005333	Nardi et al.,2003
Bactrocera papayae	Tephritidae	15915	NC_009770	Ye et al., 2010
Bactrocera philippinensis	Tephritidae	15915	NC_009771	Ye et al.,2010
Bactrocera tryoni	Tephritidae	15925	NC_014611	Nardi et al.,2010
Bactrocera cucurbitae	Tephritidae	15825	NC_016056	Wu,P.-F. unpublished
Simosyrphus grandicornis	Syrphidae	16141	NC_008754	Cameron et al., 2007
Trichophthalma punctata	Nemestrinidae	16396	NC_008755	Cameron et al., 2007
Cydistomyia duplonotata	Tabanidae	16247	NC_008756	Cameron et al., 2007
Rhopalomyia pomum	Cecidomyiidae	14503	NC_013063	Beckenbach and Joy, 2009
Mayetiola destructor	Cecidomyiidae	14759	NC_013066	Beckenbach and Joy, 2009
Culicoides arakawae	Ceratopogonidae	18132	NC_009809	Matsumoto et al.,2009
Aedes aegypti	Culicidae	16655	NC_010241	Behura et al., 2011
Aedes albopictus a	Culicidae	16665	NC_006817	Ho, C.-M. et al. unpublished
Anopheles darlingi	Culicidae	15386	NC_014275	Moreno et al., 2010
Anopheles funestusb	Culicidae	–	NC_008070	Krzywinski et al., 2006
Anopheles gambiae	Culicidae	15363	NC_002084	Beard et al., 1993
Anopheles quadrimaculatus	Culicidae	15455	NC_000875	Mitchell et al., 1993
Anopheles albitarsis	Culicidae	15413	HQ_335344	Krzywinski et al., 2011
Anopheles deaneorum	Culicidae	15424	HQ_335347	Krzywinski et al., 2011
Anopheles janconnae	Culicidae	15425	HQ_335348	Krzywinski et al., 2011
Anopheles oryzalimnetes	Culicidae	15422	HQ_335345	Krzywinski et al., 2011
Culex pipiens	Culicidae	14856	NC_015079	Atyame et al., 2011
Culex quinquefasciatus	Culicidae	15587	NC_014574	Behura et al., 2011
Chironomus tepperi	Chironomidae	15652	NC_016167	Beckenbach, 2012
Trichocera bimacula	Trichoceridae	16140	NC_016169	Beckenbach, 2012
Paracladura trichoptera	Trichoceridae	16143	NC_016173	Beckenbach, 2012
Sylvicola fenestralis	Anisopodidae	16234	NC_016176	Beckenbach, 2012
Bittacomorphella fenderiana	Ptychopteridae	15609	JN_861745	Beckenbach, 2012
Ptychoptera sp.	Ptychopteridae	15214	NC_016201	Beckenbach, 2012
Protoplasa fitchii	Tanyderidae	16154	NC_016202	Beckenbach, 2012
Cramptonomyia spenceri	Pachyneuridae	16274	NC_016203	Beckenbach, 2012
Arachnocampa flava	Keroplatidae	16923	NC_016204	Beckenbach, 2012
Tipula abdominalis	Tipulidae	14566	JN_861743	Beckenbach, 2012
Spilonota lechriaspis	Tortricidae (Lepidoptera)	15368	NC_014294	Zhao et al., 2010

Note ‘−’ not available (unknown or incomplete data).

In order to reduce time required for sequencing and walking, we used both standard and nested PCR techniques. The PCR conditions for all of the fragments are shown in Table 1. The control region was amplified using a nested PCR approach. The “external” primers SR-J-14646 and N2-N-757 were used for the first step of the PCR (long target), followed by nested amplification (specific target) with the “internal” primers SR-J-14646 and N2-N-309. All of the fragments were amplified using TaKaRa LA Taq (Takara Co., Dalian, China), and performed on an Eppendorf Mastercycler gradient in 50 µl reaction volumes. The reaction volume consisted of 25.5 µl of sterilized distilled water, 5 µl of 10× LA PCR Buffer II (Takara), 5 µl of 25 mM MgCl2, 8 µl of dNTPs Mixture, 1.5 µl of each primer (10 µM), 3 µl of DNA template, and 0.5 µl (1.25 U) of TaKaRa LA Taq polymerase (Takara).

Sequence Assembly and Annotation

The PCR products were detected via electrophoresis in 1% agarose gel, purified using the 3S Spin PCR Product Purification Kit, and sequenced directly with the ABI-377 automatic DNA sequencer. All amplified products were sequenced directly using upstream and downstream primers from both directions. Sequencing was performed using ABI BigDye ver 3.1 dye-terminator sequencing technology and run on ABI PRISM 3730 × 1 capillary sequencers. Raw sequences were manually checked and assembled using the software BioEdit version 7.0.9.0 [43] and SeqMan (DNAStar, Steve ShearDown, 1998–2001 version reserved by DNASTAR Inc., Madison, Wisconsin, USA).

Protein-coding and ribosomal RNA genes and gene boundaries were identified by BLAST search (http://www.ncbi.nlm.nih.gov/BLAST) and by comparison with sequences from other brachyceran species. The genomic positions and secondary structures of 20 of the 22 transfer RNAs (tRNAs) were identified by tRNAscan-SE software v.1.21 [44], with the remaining two (tRNAArg and tRNASer(AGN)) identified by visual inspection of genome sequence followed by inference using RNAstructure Ver.5.3 [45] (Table 3). Nucleotide composition was calculated using MEGA 4.1 [46]. Sequence data have been deposited in the NCBI database [GenBank: NC_018118].

Table 3

Organization of the mitogenome of Elodia flavipalpis Aldrich.

Gene	Strand	Locationa		Length(bp)	IGNb	CodonStart/Anti Stop		AT%
tRNAIle	+		1–65	65	−3	GAT		77.0
tRNAGln	−		63–131	69	6	TTG		86.2
tRNAMet	+		138–206	69	0	CAT		71.0
ND2	+		207–1217	1011	−2	ATT	TAA	83.4
tRNATrp	+		1216–1283	68	−8	TCA		80.9
tRNACys	−		1276–1342	67	2	GCA		74.6
tRNATyr	−		1344–1408	65	6	GTA		80.0
CO1	+		1415–2953	1539	−5	TCG	TAA	72.9
tRNALeu(UUR)	+		2949–3014	66	4	TAA		77.3
CO2	+		3019–3706	688	0	ATG	T	77.0
tRNALys	+		3707–3777	71	−1	CTT		69.0
tRNAAsp	+		3777–3847	71	0	GTC		87.4
ATP8	+		3848–4012	165	−7	ATT	TAA	86.8
ATP6	+		4006–4683	678	−1	ATG	TAA	78.4
CO3	+		4683–5471	789	6	ATG	TAA	73.0
tRNAGly	+		5478–5542	65	0	TCC		83.1
ND3	+		5543–5896	354	0	ATT	TAA	82.2
tRNAAla	+		5897–5963	67	0	TGC		79.1
tRNAArg	+		5966–6027	62	11	TCG		72.6
tRNAAsn	+		6039–6103	65	0	GTT		78.5
tRNASer(AGN)	+		6104–6171	68	0	GCT		75.0
tRNAGlu	+		6172–6233	62	18	TTC		92.0
tRNAPhe	−		6252–6316	65	0	GAA		80.0
ND5	−		6317–8036	1720	15	ATT	T	80.9
tRNAHis	−		8052–8115	64	1	GTG		84.4
ND4	−		8117–9455	1339	−7	ATG	T	81.4
ND4L	−		9449–9745	297	2	ATG	TAA	84.5
tRNAThr	+		9748–9812	65	0	TGT		86.1
tRNAPro	−		9813–9877	65	2	TGG		81.5
ND6	+		9880–10404	525	−1	ATT	TAA	86.7
CYTB	+		10404–11540	1137	−2	ATG	TAG	75.9
tRNASer(UCN)	+		11539–11605	67	16	TGA		82.1
ND1	−		11622–12569	948	1	TTG	TAA	81.1
tRNALeu(CUN)	−		12571–12635	65	0	TAG		81.6
lrRNA	−		12636–13972	1337	0			84.6
tRNAVal	−		13973–14044	72	0	TAC		80.6
srRNA	−		14045–14827	783	0			81.7
Control region	−		14828–14932	105	0			92.4

Note:

Gene positions with parentheses indicate the genes encoded by major strand; plus (+) and minus (−) symbols represent major and minor strands, respectively.

IGN: Intergenic nucleotide, minus indicates overlapping between genes. tRNA: where X is the abbreviation of the corresponding amino acid.

Phylogenetic Analysis

We conducted a phylogenetic analysis using all of the mitogenomes of Diptera that were available as of August 2012 (58 species from 33 genera and 23 families), along with a lepidopteran outgroup species (Spilonota lechriaspis) (Table 2). To assess the impact of saturation at third codon positions, we constructed two mitochondrial sequence alignments. The first dataset consisted of all 13 protein-coding genes and the 2 ribosomal RNA genes. The second dataset was the same, except for the exclusion of the third codon sites from the protein-coding genes. Sequences of protein-coding genes were translated into amino acid sequences, aligned using ClustalX v2.0.9 [47], then back-translated. Ribosomal RNA genes were also aligned using ClustalX, and refined by eye to account for their secondary structures. The GTR+I+G model was selected by the Akaike information criterion for both datasets in MODELTEST v.3.7 [48]. Maximum likelihood (ML) and Bayesian methods were used for phylogenetic analysis. For the ML analyses, we used RAxML v7.0.4 [49] with 1000 bootstrap replicates. The Bayesian phylogenetic analysis was performed using MrBayes v3.1.2. [50], with posterior distributions estimated using Markov chain Monte Carlo (MCMC) sampling. We conducted two independent runs, each with one cold and three heated chains. Samples were drawn every 100 MCMC steps over a total of 2 million steps. The first 25% of steps were discarded as burn-in.

Divergence times were estimated using the Bayesian phylogenetic method implemented in BEAST v.1.6.2 [51]. Rate variation among lineages was modelled using an uncorrelated lognormal relaxed clock [52], with the tree prior generated using a Yule speciation model. Based on previous research on divergence times in Diptera [7], [11], [12], [28], [37], we implemented fossil-based minimum ages for three clades and added one maximum age constraint. The origin of Diptera was assumed to be between 270 and 230 million years ago (mya) [28], [53]. We also specified minimum age constraints of 195 mya for Brachycera and 64 mya for Schizophora [53]–[55]. Posterior estimates of parameters were obtained by MCMC sampling. Samples were drawn every 1000 steps over a total of 10 million steps. Tracer v1.5 [56] was used to confirm that the effective sample size of each parameter exceeded 100. The maximum-clade-credibility tree was calculated using TreeAnnotator v1.6.2, with the node times scaled to match mean posterior estimates.

Results and Discussion

Genome Organization

The 14932 bp mitogenome of El. flavipalpis contained all 37 genes usually present in bilaterians: 13 protein-coding genes, 22 tRNA genes, and 2 rRNA genes (Figure 1, Table 3). The gene order corresponds to the typical pattern of brachyceran flies, such as Drosophila spp. [1], [22], [57]. Gene order is typically conserved in brachyceran flies, except for a few species in which additional tRNAs have been detected in the control region (Liriomyza trifolii, Chrysomya spp., and Cydistomyia duplonotata), and Dermatobia hominis, in which an additional tRNAVal is found on the minor strand. In contrast, there is an elevated number of tRNA rearrangements in nematoceran flies [13], [14], [20], [21].

Figure 1

Mitochondrial genome map of Elodia flavipalpis.

Numbers indicate non-coding nucleotides between genes (positive values) or gene overlap (negative values). Arrows indicate orientation on (+) strand (clockwise) or (−) strand (counterclockwise).

In total, there were 37 overlapping nucleotides between neighboring genes at 10 locations, with the length of overlapping sequence ranging from 1 to 7 bp. Excluding the control region, there were 90 intergenic nucleotides at 12 locations, in stretches ranging from 1 to 18 bp (Table 3).

As with other insects, the nucleotide composition of the El. flavipalpis mitogenome was biased towards A and T. The overall A+T content was 79.97%, slightly higher than that of Ex. sorbillans (78.4%) and R. goerlingiana (77.7%), and is among the highest of the sequenced dipteran species (67.2%–85.2%; Table S1). A detailed comparison of nucleotide composition, AT-skew, and GC-skew between the two closely related species, El. flavipalpis, Ex. sorbillans and R. goerlingiana, is given in Table 4.

Table 4

Comparison of mitochondrial nucleotide composition in three tachinid flies.

Region	A+T %			G+C %			AT-skew			GC-skew
	El. fla	Ex. sor	R. goe	El. fla	Ex. sor	R. goe	El. fla	Ex. sor	R. goe	El. fla	Ex. sor	R. goe
Whole mitogenome	79.9	78.4	77.7	20.1	21.5	22.3	0.00	0.02	0.04	−0.15	−0.17	−0.23
Protein-coding genes	79.1	77.7	76.2	20.9	22.3	23.8	−0.14	−0.15	−0.15	0.04	0.00	0.00
1st codon position	71.2	70.4	70.1	28.8	29.6	29.9	−0.19	−0.18	−0.07	0.12	0.09	0.22
2nd codon position	79.1	78.0	67.0	20.9	22.0	33.0	−0.19	−0.20	−0.08	−0.17	−0.16	−0.16
3rd codon position	87.1	84.7	91.2	12.9	15.3	8.8	−0.07	−0.07	−0.04	0.19	0.07	−0.16
tRNA genes	79.8	76.8	77.2	20.2	23.2	22.8	0.03	0.02	0.02	−0.11	−0.11	−0.12
lrRNA	84.6	83.2	82.6	15.5	16.8	17.4	0.00	0.05	0.05	−0.30	−0.30	−0.34
srRNA	81.7	79.4	80.5	18.3	20.5	19.5	−0.04	0.01	0.00	−0.30	−0.30	−0.28
Control region	92.4	98.1	92.6	7.7	1.9	7.4	0.11	−0.05	0.11	0.74	1.00	−0.71

Note: El. fla indicates Elodia flavipalpis, Ex. sor indicates Exorista sorbillans and R. goe indicates Rutilia goerlingiana. The A+T and G+C biases of protein-coding genes were calculated by AT-skew = [A−T]/[A+T] and GC-skew = [G−C]/[G+C], respectively.

Protein-coding Genes and Nucleotide Composition

Thirteen protein-coding genes were identified in the mitogenome of El. flavipalpis, with characteristics similar to those of other dipteran species (Table S1). The average A+T content across all protein-coding genes was 79.1%, similar to that of Ex. sorbillans (77.7%) and R. goerlingiana (76.2%). Table 4 shows the AT-skews and CG-skews of the three codon positions for El. flavipalpis, Ex. sorbillans and R. goerlingiana. In all three of these species, the A+T content of the third codon positions (87.1%, 84.7%, and 91.2%, for El. flavipalpis, Ex. sorbillans, and R. goerlingiana, respectively) was higher than those of the first (71.2%, 70.4%, and 70.1) and second codon positions (79.1%, 78.0%, and 67%). This result is in agreement with studies of other dipteran taxa (Table S1). AT-skew and GC-skew were used to analyse the biases in nucleotide composition. The A content was slightly lower than the T content at all three codon positions, but almost equal over the whole genome.

Except for CO1 and ND1, all of the protein-coding genes have one of the common start codons for mitochondrial DNA, ATG, ATA, or ATT (Table 3). The start codon TCG (Serine) in CO1 is also found in Ex. sorbillans, R. goerlingiana, and other Oestroidea species (Table 2). CO1 commonly uses nonstandard start codons in many other flies [5], [16], [20], [58], [59]. Within Diptera, the start codon TTG (Leucine) in ND1 of El. flavipalpis is same as that in Dermatobia hominis (Oestroidea). Of the 13 protein-coding genes, CO2, ND4, and ND5 have incomplete stop codons and terminate with only a single thymine. Similar structural features have also been described for the mitogenomes of other dipteran taxa [4], [5]. In addition, CYTB terminates with a complete stop codon TAG, whereas TAA or only a single thymine is utilized in some other fly species.

Other Regions of the Mitogenome

The mitogenome of El. flavipalpis bears all of the 22 standard tRNAs found in metazoan mitogenomes (Figure 1, Table 3). The total length of the tRNA genes is 1463 bp, with individual genes ranging from 62 to 71 bp and with A+T contents from 71% (tRNAMet) to 92% (tRNAGlu). With the exception of tRNASer (AGN), all tRNAs possess the typical clover-leaf secondary structure (Figure 2). The DHU-arm of tRNASer (AGN) is entirely absent, as observed in other insects [20], [24], [33], [60]. In the secondary structures, the lengths of the amino acid acceptor arms (7 bp), anticodon arms (5 bp), and loops (7 bp) are relatively conserved, while the TψC loop (3–10 bp) is more variable. There are 21 mismatched base pairs in the tRNA genes, 14 of which are weak G–U matches (nine sites in DHU arms, three sites in amino acid acceptor arms, and two sites in anticodon arms). The other seven include U–U (5 bp), C–U (1 bp), and A–A (1 bp).

Figure 2

Putative secondary structures of tRNAs found in the mitochondrial genome of Elodia flavipalpis.

All tRNAs can be folded into the usual clover-leaf secondary structure.

The lengths of the lrRNA and srRNA genes are 1337 bp and 783 bp, respectively. Both are encoded on the minor strand and the ends of those genes were assumed to be at the boundaries of the flanking genes [61]. As in other dipteran species, the lrRNA gene is flanked by tRNALeu (CUN) and tRNAVal, while the srRNA gene is between tRNAVal and the control region. Their A+T contents were 84.6% for lrRNA and 81.7% for srRNA, which are within the range of other dipteran species (Table S1).

The length of the control region of El. flavipalpis is identical to that of Ex. sorbillans (105 bp), the shortest among the sequenced dipteran mitogenomes. It has an A+T content of 92.38%, which is lower than that of Ex. sorbillans (98.1%) and R. goerlingiana (92.6%) but higher than those of most other dipteran species. Owing to its short length, there is no distinct duplicate fragment found in this region. It should be noted that all three of the sequenced tachinid mitogenomes bear a control region that is shorter than those of most known in flies [62], [63].

The phylogenies estimated using likelihood and Bayesian approaches were similar for both of the datasets that were analysed (Figures 3, 4, 5 and 6). Various higher-level relationships were consistent across the analyses. The monophyly of Brachycera, Cyclorrhapha, and Calyptratae were consistently supported (posterior probability = 1.00, ML bootstrap = 100), as was the monophyly of Schizophora (posterior probability = 1.00, ML bootstrap = 55, 66). The monophyly of superfamily Oestroidea has been widely accepted and has traditionally received good support from morphological characters [64]–[67]. Here we add support from the Bayesian analysis of mitochondrial DNA sequences that Oestroidea is a sister group of Muscidae (posterior probability = 1.00). However, our ML analyses of both datasets placed the family Muscidae within Oestroidea, although support was not high (bootstrap = 70, 60).

Figure 3

Bayesian tree of Diptera, inferred from a mitochondrial data set comprising 13 protein-coding genes and 2 ribosomal RNA genes.

The tree was rooted using the outgroup taxon Spilonota lechriaspis (Lepidoptera). Numbers denote posterior probabilities of nodes. The lengths of very long branches have been reduced to aid viewing. The symbol ‘//’ indicates a contracted branch, with the value above giving the length of contraction. Red lines indicate the differences among the four phylogenetic trees.

Figure 4

Maximum-likelihood tree of Diptera, inferred from a mitochondrial data set comprising 13 protein-coding genes and 2 ribosomal RNA genes.

The tree was rooted using the outgroup taxon Spilonota lechriaspis (Lepidoptera). Numbers denote bootstrap values in percentages. Red lines indicate the differences among the four phylogenetic trees.

Figure 5

Bayesian tree of Diptera, inferred from a mitochondrial data set comprising 13 protein-coding genes (without third codon sites) and 2 ribosomal RNA genes.

The tree was rooted using the outgroup taxon Spilonota lechriaspis (Lepidoptera). Numbers denote posterior probabilities of nodes. The lengths of very long branches have been reduced to aid viewing. The symbol ‘//’ indicates a contracted branch, with the value above giving the length of contraction. Red lines indicate the differences among the four phylogenetic trees.

Figure 6

Maximum-likelihood tree of Diptera, inferred from a mitochondrial data set comprising 13 protein-coding genes (without third codon sites) and 2 ribosomal RNA genes.

The tree was rooted using the outgroup taxon Spilonota lechriaspis (Lepidoptera). Numbers denote bootstrap values in percentages. Red lines indicate the differences among the four phylogenetic trees.

In Oestroidea, all four families are monophyletic, and Calliphoridae+Sarcophagidae was inferred as a sister group to Oestridae+Tachinidae. This result is similar to that obtained in analyses of morphology [34] and of 18S and 16S ribosomal DNAs [35]. The tree topology is broadly similar to that inferred from whole mitogenomes by Nelson et al. [27], except that Oestridae and a subfamily of Calliphoridae (Polleninae) are nested within Tachinidae. However, Nelson et al. [27] focused on the relationships within Calliphoridae, while species from other families of Oestroidea were only used as an outgroup in the analysis.

Our phylogenetic estimate differs from that obtained by Kutty et al. [36] in their analysis of four nuclear and four mitochondrial genes. They inferred the relationships (((Tachinidae, Oestridae), Calliphoridae), Sarcophagidae), with both Calliphoridae and Tachinidae paraphyletic (Oestridae is nested within Tachinidae, as are some calliphorid subfamilies). In contrast, Wiegmann et al. [28], considered Tachinidae to be more closely related to Calliphoridae, and as a sister taxon to Oestridae and Sarcophagidae. Owing to the morphological similarities shared by these four families, it is difficult to distinguish among these phylogenetic hypotheses using morphological data. The disparities among the molecular estimates are probably due to differences in the taxa sampled and the data being analysed. Kutty et al. [36] and Wiegmann et al. [28] used both nuclear and mitochondrial DNA sequences, but most were only partial sequences. Moreover, most of the Oestroidea species analysed by Wiegmann et al. [28] were represented by only two or three genes. Given that our analysis involved a larger and more complete data set, we believe that our results are more strongly supported.

The primary difference among the four phylogenetic trees here is in the placement of the superfamily Opomyzoidae, which consists of the families Furgusoninidae and Agromyzidae. It is closer to Drosophoridae in the Bayesian analysis (posterior probability = 1.00), but its position is unstable in the ML analysis (bootstrap = 55, 27). A similar result was obtained by Wiegmann et al. [28]. Another difference is seen in the placement of Bibionomorpha (Nematocera), which is the closest group to Brachycera in both of the trees inferred without third codon positions, but is unstable in the tree inferred from all three gene positions. It is even non-monophyletic in the Bayesian analysis, hinting at the possible negative phylogenetic effects of including third codon sites. The placement of Cecidomyiidae is problematic, which is also indicated by the long branch leading to this group. The two Cecidomyiidae species have undergone substantial reduction in mitogenome size, which results in their apparent distinctiveness and causes problems for sequence alignment. With additional sampling in Cecidomyiidae, future studies will be better equipped to reconstruct the molecular evolution of these mitochondrial genomes.

Estimates of Divergence Times

We used a Bayesian relaxed clock to estimate the evolutionary timescale of Brachycera (Figure 7). Our analysis suggests that the last common ancestor of extant Brachycera existed in the early Jurassic (∼199 mya) (95% credibility interval: 195.9–206.7 mya). The schizophoran radiation took place during the late Cretaceous (∼84 mya), and the clade Calyptratae containing Oestroidea and Muscidae (nested in Acalyptrata) split in the early Eocene (∼60 mya) (95% credibility interval: 45.6–91.1 mya). These results are consistent with evidence from a number of amber specimens from Acalyptrata and Calyptratae [68]. Subsequently, Oestroidea divided into two groups in the middle Palaeocene (∼56 mya) (95% credibility interval (CI): 42.7–85.0 mya). The split of Tachinidae with Oestridae is estimated to have occurred in the early Eocene (∼48 mya) (95% CI: 37.8–77.9 mya), with this clade becoming the most speciose in Brachycera. Finally, the two tachinid flies El. flavipalpis and Ex. sorbillans are estimated to have separated in the early Oligocene (∼33 mya) (95% CI: 15.5–60.1 mya).

Figure 7

Evolutionary timescale for Diptera inferred from a mitochondrial data set comprising 13 protein-coding genes and 2 ribosomal RNA genes.

Numbers at nodes indicate mean estimated divergence times (in mya) and node bars indicate 95% credibility intervals. Red circles indicate the three nodes used for calibration. The yellow circle indicates the hypothesised origin of tachinid flies. In the geological time scale: Pala indicates Palaeocene; Eoce indicates Eocene; Oligo indicates Oligocene; Mioc indicates Miocene; P indicates Pliocene; Q indicates Quaternary.

Our estimates suggest that the most recent common ancestor of tachinid flies existed between 33 and 48 mya. The two tachinid samples used in this study are from the same subfamily Exoristinae, but are in separate tribes Goniini (El. flavipalpis) and Exoristini (Ex. sorbillans). Among the four subfamilies of Tachinidae, Exoristinae was probably the latest to emerge [69]–[71]. Therefore, the time of origin of tachinid flies should be in the earlier half of the time interval described above. We speculate that the most recent common ancestor of tachinid flies existed in the middle Eocene (∼42–48 mya). Tachinid flies have a worldwide distribution, but are mainly found in Palaearctic and Nearctic regions. For a globally distributed family with significant differences in distribution between northern and southern continents, the relevant split is between the supercontinents Laurasia and Gondwana in the mid-Mesozoic. Our estimates for the origin of Tachinidae is much more recent than this. However, given that most tachinid species are distributed throughout the Holarctic Region, we suggest that the evolutionary history of tachinid flies is probably tied to the split of Laurasia in the Eocene rather than that between Laurasia and Gondwana.

We have shown that the availability of additional mitogenomes can make a valuable contribution to our understanding of the phylogeny and divergence times of Diptera. Our study serves as a useful primer for the evolution of tachinid flies, but the accuracy of divergence-time estimates can be improved by denser sampling of tachinid mitogenomes, a greater number of fossil calibrations, and combination with nuclear genes or morphological data. In addition, we suggest that the stable gene order among brachyceran species might be due to their comparatively short evolutionary timeframe.

Nucleotide composition of all available mitogenome sequences from Diptera.

(DOC)

Click here for additional data file.