Next Article in Journal
A Pesticide Residues Insight on Honeybees, Bumblebees and Olive Oil after Pesticidal Applications against the Olive Fruit Fly Bactrocera oleae (Diptera: Tephritidae)
Previous Article in Journal
Landscape Simplification Modifies Trap-Nesting Bee and Wasp Communities in the Subtropics
Open AccessArticle

New Mitochondrial Gene Rearrangement in Psyttalia concolor, P. humilis and P. lounsburyi (Hymenoptera: Braconidae), Three Parasitoid Species of Economic Interest

1
Department of Genetics, Stellenbosch University, Private Bag X1, Matieland 7602, South Africa
2
Department of Agricultural, Food and Forest Sciences, Università degli Studi di Palermo, Viale delle Scienze, Edificio 5, 90128 Palermo, Italy
3
Unit for Cardiac and Cardiovascular Genetics, Department of Medical Genetics, Oslo University Hospital, Postboks 4956 Nydalen, 0424 Oslo, Norway
4
Norsk Entomologisk Forening, Naturhistorisk Museum, Universitetet i Oslo, Postboks 1172 Blindern, 0318 Oslo, Norway
*
Authors to whom correspondence should be addressed.
Authors contributed equally to this work.
Insects 2020, 11(12), 854; https://doi.org/10.3390/insects11120854
Received: 14 November 2020 / Revised: 23 November 2020 / Accepted: 25 November 2020 / Published: 2 December 2020
(This article belongs to the Section Insect Molecular Biology and Genomics)
Parasitoid wasps in the family Braconidae are generally highly specialized and can be used as agents for biological control of arthropod pests. Psyttalia concolor, Psyttalia humilis and Psyttalia lounsburyi parasitize the larvae of the olive fruit fly (Bactrocera oleae), the most damaging pest of cultivated olives in the world. Psyttalia concolor is native to the Mediterranean, and P. humilis and P. lounsburyi are native to sub-Saharan Africa. Despite their potential for agricultural pest control, these species have been poorly characterized at the genetic level. We sequenced the mitochondrial genome of the three species and compared its organization with other Braconidae. Psyttalia had a unique gene rearrangement involving the positions of transfer RNA genes. We also present a phylogenetic reconstruction of the Braconidae and confirm the phylogenetic placement of Psyttalia in the subfamily Opiinae.

Abstract

The family Braconidae consists mostly of specialized parasitoids, some of which hold potential in biocontrol of agricultural pests. Psyttalia concolor, Psyttalia humilis and Psyttalia lounsburyi are parasitoids associated with Bactrocera oleae, a major pest of cultivated olives. The native range of Psyttalia concolor is the Mediterranean, and P. humilis and P. lounsburyi are native to sub-Saharan Africa. This study reports the mitochondrial genomes of the three species, thus laying the foundation for mitogenomic analyses in the genus Psyttalia. Comparative mitogenomics within Braconidae showed a novel gene arrangement in Psyttalia in involving translocation and inversion of transfer RNA genes. The placement of Psyttalia in the subfamily Opiinae was well-supported, and the divergence between Psyttalia and its closest relative (Diachasmimorpha longicaudata) was at ~55 MYA [95% highest posterior density (HPD): 34–83 MYA]. Psyttalia lounsburyi occupied the most basal position among the three Psyttalia, having diverged from the other two species ~11 MYA (95% HPD: 6–17 MYA). Psyttalia concolor and P. humilis were recovered as sister species diverged at ~2 MYA (95% HPD: 1.1–3.6 MYA). This phylogeny combining new sequences and a set of 31 other cyclostomes and non-cyclostomes highlights the importance of a comprehensive taxonomic coverage of Braconidae mitogenomes to overcome the lack of robustness in the placement of several subfamilies.
Keywords: braconidae; cyclostome wasps; mitogenomics; opiinae; olive; phylogeny; Psyttalia humilis; Psyttalia lounsburyi braconidae; cyclostome wasps; mitogenomics; opiinae; olive; phylogeny; Psyttalia humilis; Psyttalia lounsburyi

1. Introduction

The family Braconidae is a species-rich group that includes 40 subfamilies represented by over 1000 genera and more than 19,000 known species [1,2]. Braconidae are mostly composed of highly specialized parasitoids, and the majority of the subfamilies therein are ectoparasitic idiobionts (i.e., the host is unable to recover after the paralysis induced by the ovipositing wasp), or endoparasitic koinobionts (i.e., the host can recover after oviposition and develop normally, completing all larval instars) [3,4]. In general, Braconidae exhibit host-specific relationships at the subfamily level; however, this is less true for ectoparasitoids [1,4]. For example, the endoparasitic Microgastrinae attack only lepidopteran larvae, with the exception of one species associated with Trichoptera [5], and the endoparasitic Helconinae parasitize coleopteran larvae. In contrast, the ectoparasitic Braconinae attack a variety of holometabolous larvae, and the subfamily has been used as a model for studying the evolutionary transition between ecto- and endoparasitism [1,4]. Braconidae are divided into two major groupings of subfamilies: the cyclostomes and the non-cyclostomes. Cyclostomes are distinguished by a cavity above the mandible (hypoclypeal depression) which is a synapomorphy of the group, and comprise all the ectoparasitoids, some endoparasitoids and all known phytophagous braconids [1,3]. The cyclostome complex has been reported as monophyletic based on morphology [6,7], with the remaining non-cyclostome subfamilies as a sister clade based on integrated molecular and morphological data [8]. Molecular studies using the mitochondrial 16s rRNA and the nuclear 28s rRNA genes also recovered cyclostomes as monophyletic [9,10,11]. However, the phylogenetic relationships within cyclostomes have not been recovered with high statistical support, despite extensive taxon coverage [3]. Although the family Braconidae has received considerable taxonomic attention in recent years, substantial confusion persists over the definitions of several subfamilies, especially among cyclostomes [8].
The olive fruit fly Bactrocera oleae (Rossi, 1790) (Diptera: Tephritidae) has been a major pest of cultivated olives in the Mediterranean Basin since historical times. More recently, the species became an important threat to olive production in California where it quickly spread after the invasion was first detected in 1998 [12]. The olive fruit fly is controlled using primarily insecticides, which have limited success and negatively impact the environment [13]. Moreover, conventional pest control has been associated with increased frequency of insecticide resistance alleles in olive fruit fly populations [14,15,16,17]. Efforts to find agents for the biocontrol of B. oleae started over 100 years ago, and surveys for natural enemies have been conducted in South Africa, Namibia, Kenya, La Réunion, Canary Islands, Morocco, Pakistan, India and China [18]. The highest species diversity of parasitoid wasps (Braconidae and Chalcidoidea) associated with olive fruit flies in a single geographic region was found in the Western Cape province of South Africa, on native African wild olives [Olea europaea L. subsp. cuspidata (Wall ex G. Don Cif.)] [19,20]. The assemblage included four Braconidae koinobiont endoparasitoids endemic to sub-Saharan Africa: Bracon celer (Szépligeti, 1913), Utetes africanus (Szépligeti, 1910), Psyttalia humilis (Silvestri, 1913) and Psyttalia lounsburyi (Silvestri, 1913).
Psyttalia lounsburyi was described by Silvestri (1913) as a parasitoid of olive fruit flies on African wild olives in South Africa. Psyttalia lounsburyi was found to be genetically distinct from P. humilis (see below) and from their Mediterranean counterpart Psyttalia concolor (Szépligeti, 1910) [21]. Psyttalia lounsburyi has been reported in Kenya and South Africa, where it was recovered from B. oleae infesting wild olives [20,22]. Therefore, P. lounsburyi is thought to be a sub-Saharan Africa parasitoid specializing in B. oleae. However, the fact that it also accepts Ceratitis capitata (Wiedemann, 1824) as a host under laboratory conditions raises the possibility that it can parasitize other Bactrocera spp. in the wild, particularly Bactrocera biguttula (Bezzi, 1922), a close relative of B. oleae known to utilize African wild olives in South Africa, Namibia, and Kenya [21,23,24], and B. munroi White, 2004 also found it in Kenya on African wild olives [25]. Psyttalia humilis was described by Silvestri (1913) based on specimens reared from pears infested by C. capitata in Constantia, Cape Town (South Africa). The species has been reared from B. oleae collected from African wild olives in Kenya and South Africa [20,24,25]. Psyttalia humilis is morphologically indistinguishable from the Mediterranean P. concolor and has sometimes been treated as its junior synonym [26]. However, the fact that P. humilis has been recorded only in sub-Saharan Africa and P. concolor only in the in the Mediterranean Basin, and the genetic divergence found in DNA analyses across the genus Psyttalia, supports that P. humilis and P. concolor can be treated as separate species [21]. Psyttalia concolor is an endoparasitoid of B. oleae found on wild and cultivated olives in the Mediterranean region. Psyttalia concolor was first identified as an olive fruit fly parasitoid in Tunisia, and is also considered native to Sicily, southern Sardinia and southern Calabria [27,28]. More recently, the parasitoid was found in various areas of coastal Tuscany [29]. Psyttalia concolor has also reportedly been reared from medfly (C. capitata) infesting argan fruit (Argania spinosa L., Sapotaceae) in Morocco [30]. However, those specimens were not subjected to DNA-based analyses, and the indistinguishability between P. humilis and P. concolor demands caution in the identification of Psyttalia, especially if emerged from hosts other than B. oleae, which is presently the only confirmed host of P. concolor [21].
Psyttalia concolor has been used in trials for biological control of the olive fruit fly in the Mediterranean [31,32]. In 2003, California initiated a program focused on the evaluation and release of P. humilis and P. lounsburyi, but the introductions had limited success, as only P. lounsburyi, the most specialized of the two parasitoids was recovered [18,33,34].
Despite their potential utility and interesting evolutionary specialization as parasitoids of the olive fruit fly, P. concolor, P. humilis and P. lounsburyi have not been fully characterized at the level of the mitochondrial sequence. Insect mitochondrial genomes are powerful sources of information for the reconstruction of phylogenetic relationships due to their maternal inheritance, lack of recombination, conserved gene components and organization and relatively small size [35]. The genus Psyttalia (Walker, 1860) has not been represented in comparative mitogenomics, and its positioning within the family Braconidae has never been assessed in previous phylogenies using complete or near-complete mitogenome sequences [36,37,38]. Mitochondrial gene rearrangements can be particularly interesting in phylogenetic analyses because they occur frequently in certain groups of insects, including Hymenoptera, but are uncommon in closely related taxa [36,39,40]. Therefore, mitochondrial gene rearrangements provide additional information to help resolve deep phylogenetic nodes. A recent study explored the possibility of using mitogenome rearrangements to reconstruct phylogenetic relationships in Braconidae [37], and the results highlighted the importance of obtaining complete mitogenome sequences, as two regions previously known to harbor gene rearrangements in braconids [38] were not sequenced, thus potentially reducing the resolving power of the analysis. The present work lays the foundation for mitogenomics in the genus Psyttalia, and the clarification of the phylogenetic relationships of P. concolor, P. lounsburyi and P. humilis within Braconidae.

2. Materials and Methods

2.1. Sample Collection and Species Identification

Adult specimens of P. humilis and P. lounsburyi were reared from African wild olives (Olea europaea L. subsp. cuspidata) collected in April and May 2016 in Grahamstown (33.3195° S, 26.5171° E) and Stellenbosch (33.9951° S, 18.8676° E), respectively situated in the Eastern and the Western Cape province of South Africa. Adult specimens of P. concolor were reared from cultivated olives (O. europaea L. subsp. europaea var. europaea) collected in November 2014 in Constância (39.4781° N, 8.3372° W), in the Ribatejo province of Portugal. Morphological species identification was performed on ethanol-preserved adult specimens, using the taxonomic keys and photographic images available in the Parasitoids of Fruit-Infesting Tephritidae (PAROFFIT) database (http://paroffit.org), and previous descriptions [41] (Figure 1). Morphological species identification was confirmed by comparing DNA barcodes (650 bp; COI-5’) of P. concolor, P. humilis and P. lounsburyi and homologous sequences available on GenBank as of 18 November 2020. Intra- and interspecific genetic divergences were estimated as pairwise distances (p-distances) under the Kimura 2-parameter (K2P) model [42] in MEGA X [43]. Standard errors were calculated from 1000 bootstrap replicates.

2.2. DNA Extraction, Polymerase Chain Reaction (PCR) Amplification and Sequencing

Total DNA was extracted from a single adult specimen representative of each species using a standard SDS/Proteinase K method for P. concolor, and a standard phenol–chloroform method for P. humilis and P. lounsburyi.

2.2.1. Sanger Sequencing

The complete mitogenome of P. concolor (15,308 bp) was obtained by Sanger sequencing, after PCR amplification in 18 overlapping segments, with shorter versions of four of these also used for sequencing (Table S1). All PCR and sequencing primers were designed by the authors specifically for this study except for segment S03*, which was amplified and sequenced using arthropod universal DNA barcoding primers [44]. PCR was carried out in 25 μL total volume containing 75 mM Tris-HCl (pH 8.8), 20 mM (NH4)2SO4, 0.01% (v/v) Tween 20 (Fermentas), 3.0 mM MgCl2 (Fermentas), 0.5 mM of each dNTP (Fermentas), 25 pmol of each primer (Macrogen) and 2.5 U of Taq DNA polymerase (Fermentas). Three hotstart and touchdown cycling protocols were used: (a) PAS1, consisting of 95 °C for 5 min; three cycles of 95 °C for 30 s; 60 °C for 1 min and 72 °C for 2 min; three cycles of 95 °C for 30 s; 57 °C for 1 min and 72 °C for 2 min; three cycles of 95 °C for 30 s; 54 °C for 1 min and 72 °C for 2 min; 38 cycles of 95 °C for 30 s, 58 °C for 1 min and 72 °C for 2 min; and 72 °C for 5 min; (b) PAS2, consisting of 95 °C for 5 min; two cycles of 95 °C for 30 s, 60 °C for 1 min and 72 °C for 2 min; 2 cycles of 95 °C for 30 s; 58 °C for 1 min and 72 °C for 2 min; two cycles of 95 °C for 30 s; 56 °C for 1 min and 72 °C for 2 min; 38 cycles of 95 °C for 30 s; 58 °C for 1 min and 72 °C for 2 min; and 72 °C for 5 min; and (c) PAS3, differing from PAS2 only in the extension step (68 °C for 3 min). PCR products were purified by treatment with ExonucleaseI (Fermentas) and Shrimp Alkaline Phosphatase (Fermentas), and Sanger-sequenced by Macrogen Europe (Amsterdam, The Netherlands). Mitochondrial DNA is usually overrepresented in Next Generation Sequencing (NGS) reads, allowing for easy retrieval of mitogenomes [45], as well as completion and curation of published mitochondrial sequences [23]. However, the correct assembly of insect mitogenomes from NGS data can be hampered by lack of adequate reference sequences, particularly in groups where gene rearrangements are common (e.g., Hymenoptera), and complete mitogenomes for some subfamilies are not available (e.g., Opiinae). Therefore, the mitogenome of P. concolor was sequenced using Sanger technology to obtain a reference sequence for subsequent mapping of the NGS reads from P. humilis and P. lounsburyi. The complete mitogenome sequence of P. concolor was recovered using a multi-step strategy starting with the PCR amplification of seed regions with primers designed on conserved regions of the mitochondrial genomes of Diachasmimorpha longicaudata (GenBank accession GU097655.1), Spatius agrili (GenBank accession NC_014278.1) and Cotesia vestalis (GenBank accession NC_014272.1). The seed regions were then iteratively extended using PCR primers specific for the newly obtained P. concolor sequences and primers based on the other Braconidae sequences, and the gaps between the seed regions were bridged using PCR primers specifically for P. concolor.

2.2.2. Next Generation Sequencing

Psyttalia humilis and P. lounsburyi were sequenced using the Ion™ Torrent Proton™ platform (ThermoFisher Scientific, Waltham, MA, USA) available at the Central Analytical Facilities of Stellenbosch University, South Africa. Sequence libraries were prepared using the NEXTflex™ DNA Sequencing Kit for Ion Platforms (PerkinElmer, Waltham, MA, USA) according to the BI00 Scientific v15.12 protocol. Libraries were diluted to a target concentration of 60 pM. The diluted, barcoded libraries were combined in equimolar amounts for template preparation using the Ion PI™ Hi-Q™ Chef Kit (Thermo Fisher Scientific, Waltham, MA, USA). Twenty five microliters of diluted, pooled library was loaded onto the Ion Chef liquid handler (Thermo Fisher Scientific) for template preparation and enrichment using Ion PI™ Hi-Q™ Chef reagents, solutions and supplies according to the protocol, MAN0010967 REVB.0. Enriched ion sphere particles were loaded onto an Ion PI™ v3 chip. Massively parallel sequencing was performed on the Ion Torrent Proton system using sequencing solutions, reagents and supplies according to the protocol MAN0010967 REV B.0. Flow space calibration and basecaller analysis were performed using standard analysis parameters in the Torrent Suite version 5.10.0 software.

2.3. Mitogenome Assembly, Annotation and Analyses

The complete mitogenome of P. concolor was assembled using the CLCBio Main Workbench v6.9 (QIAGEN Bioinformatics), with manual curation. The NGS reads for P. humilis and P. lounsburyi were mapped and assembled to the complete mitogenome sequence of P. concolor using the mapper functionality available on Geneious Prime v2019.1 (https://www.geneious.com) with medium/low sensitivity option, and fine tuning up to five iterations. The consensus sequences were calculated using Geneious Prime. Open reading frames of protein-coding genes (PCGs) were identified using Geneious Prime, with the invertebrate mitochondrial genetic code. The position and secondary structure of transfer RNA genes (tRNAs) were predicted with ARWEN software [46] using the composite metazoan mitochondrial genetic code, and with MITOS WebServer (http://mitos.bioinf.uni-leipzig.de/index.py) using the invertebrate genetic code. Ribosomal RNA genes (rRNAs) were estimated by BLASTn search on NCBI (https://blast.ncbi.nlm.nih.gov). Overlapping regions and intergenic spacers were counted manually. Nucleotide composition and AT- and GC-skews were calculated using Geneious Prime, as AT-skew = (A − T)/(A + T) and GC-skew = (G − C)/(G + C). The sequences were deposited in GenBank under the accession numbers MW279212, MW279213 and MW279214.

2.4. Phylogenetic Analyses

The phylogenetic positioning of P. humilis, P. concolor and P. lounsburyi within Braconidae was reconstructed using 31 complete and partial mitogenomes of cyclostomes and non-cyclostomes, with two species of Ichneumonidae (Diadegma semiclausum and Enicospilus sp.) as outgroups (Table 1). In line with a previous study [37], analyses were restricted to all PCGs except ND2 due to incompleteness of several mitogenomes, and the first and second codon positions due to the temporal depth of the phylogeny. Sequences for each of the PCGs were aligned using the Translator-X server (translatorx.co.uk) [47], with alignment cleaning under less stringent selection and additional minor manual corrections. The 24 partitions corresponding to the first and second codon positions were separated using MEGA7 [48]. Subsequent analyses were performed using either the 24 partitions, or the 15-partition subset used by Li et al. (2016) [37], which was selected by excluding individual gene partitions with lower quality phylogenetic information. Three different partition-clustering schemes were tested for the datasets: (a) the partition scheme selected by PartitionFinder2 [49], run on the CIPRES Science Gateway V3.3 portal (www.phylo.org) [50] using a greedy algorithm (in line with Li et al. 2016); (b) a partition by codon position alone; (c) a partition by codon position and strand. In addition to facilitating comparisons with previous work, using PartitionFinder provides a “best-fit” (from a maximum likelihood perspective) partitioning scheme. However, it is generally advisable to test other partition-clustering schemes, particularly when phylogenetic analyses are conducted within a Bayesian framework. Partitioning by codon position alone or codon position + strand are commonly used in such comparisons as, on the one hand they have a biological basis, and on the other represent an intermediate between no partitioning and no partition clustering. Dated phylogenetic trees were obtained with a Bayesian method implemented in BEAST1.8.4 [51], with separate GTR + I + G (4 gamma categories) substitution models and lognormal relaxed clock models for each partition, but a single global tree model. The tree was left unconstrained except for monophyly requirements for both Braconidae and Ichneumonidae. A Yule process tree prior was used, and priors for divergence dates of Braconidae and Braconidae–Ichneumonidae were based on recently published data [52]. Priors for mutation rates were chosen based on previous results for insects [53], and values obtained with jModelTest2 [54]. Runs were performed for 30 million generations (main run: 15 partition data set, partition-clustering scheme selected by PartitionFinder) or 10 million generations (alternative runs, using the other partition-clustering schemes or/and the 24 partition data set), with a 10% generation burn-in and sampling every 1000 generations. Trees were summarized and annotated using TreeAnnotator v1.8.1 [51], and drawn using FigTree 1.4 (http://tree.bio.ed.ac.uk/software/figtree/).

3. Results and Discussion

3.1. Morphological and Molecular Species Identification

The adult specimens used for the recovery of the mitochondrial genomes of P. concolor, P. humilis and P. lounsburyi were identified morphologically by co-author V. Caleca. To confirm the morphological identification, intra- and interspecific genetic divergences using 650 bp of COI extracted for the new mitogenomes and all homologous sequences available on GenBank (P. concolor, n = 7; P. humilis, n = 10; P. lounsburyi, n = 32) were calculated. Intraspecific maximum p-distance was low in the three species (P. concolor = 0.32%; P. humilis, n = 0.71%; P. lounsburyi, n = 0.44%). Interspecific average p-distances were lowest for the pair P. concolor/P. humilis (4.76%; SE = 0.80), followed by P. humilis/P. lounsburyi (8.92%; SE = 1.16), and P. concolor/P. lounsburyi (9.66%; SE = 1.18). These results confirmed correct morphological identification of the specimens used for the recovery of the mitogenomes presented in the sections below.

3.2. Sequencing of the Psyttalia Mitogenomes

The complete mitogenome of P. concolor was recovered by Sanger sequencing of overlapping PCR-amplified fragments (see Material and Methods section for the amplification strategy), while the mitogenomes of P. humilis and P. lounsburyi were recovered using Ion Proton technology, resulting in 34.0 million reads (average read length of 177 bp) for P. humilis, and 11.6 million reads (average read length of 142 bp) for P. lounsburyi. Average sequence coverage was 304× for P. humilis and 285× for P. lounsburyi.
Hymenopteran mitogenomes are notoriously difficult to sequence due to very high A+T content, as well as frequent gene rearrangements. At the time of this study, sequences for all the mitochondrial genes had only been previously reported for two braconids (S. agrili and C. vestalis). In the present work, we obtained the sequences for all mitochondrial genes of P. concolor, P. humilis and P. lounsburyi. However, the AT-rich region was not successfully assembled for the last two species due to a combination of short NGS read length and extreme A+T content (90.9% in P. concolor). The AT-rich region had been previously annotated in only four braconids (S. agrili, C. vestalis, D. longicaudata and Macrocentrus camphoraphilus), and it is unclear whether it has been fully recovered in the last two species, as the reported mitogenomes lack the regions adjacent to ND2. Sequencing of the AT-rich region in Braconidae may be hampered not only by the typically high A+T content, but also by gene rearrangements and stable stem-and-loop structures commonly found in hymenopteran mitogenomes [55,56,57,58].

3.3. Organization and Composition of the Psyttalia Mitogenomes

3.3.1. General Mitogenome Organization and Gene Content

The mitogenomes of the three Psyttalia had the typical metazoan gene content with 13 PCGs, two rRNAs, and 22 tRNAs, and gene arrangement was conserved among the three species (Table 2; Figure 2). Nine PCGs (ND2, COI, COII, ATP8, ATP6, COIII, ND3, ND6, CTYB) and 12 tRNAs (tRNATrp, tRNALeu1, tRNAHis, tRNALys, tRNAGly, tRNAAla, tRNAArg, tRNAAsn, tRNASer1, tRNAGlu, tRNAThr, tRNASer2) were encoded on the majority strand (J-strand), and the remaining four PCGs (ND5, ND4, ND4L, ND1), 10 tRNAs (tRNAGln, tRNATyr, tRNACys, tRNAAsp, tRNAPhe, tRNAPro, tRNALeu2, tRNAVal, tRNAIle, tRNAMet) and the two rRNAs were encoded on the minority strand (N-strand). The tRNAGln region was poorly recovered in P. humilis and P. lounsburyi due to its location between the AT-rich region and ND2; therefore, this gene was only annotated in P. concolor.
The longest intergenic space averaged 164 bp across the three species and was located between COX2 and tRNAAsp. Psyttalia had few and short gene overlapping regions, with an average of 16 locations, and the longest overlap between ATP6 and ATP8 (22 bp).

3.3.2. Nucleotide Composition and Strand Asymmetry

The three mitogenomes were highly biased towards A and T (average A+T content = 83.8), as typically is the case in insects (Table 3). The A+T content for PCGs on the N-strand (average = 83.8%) was higher than on the J-strand (average = 83.1%), with the highest in ATP8 (90.4%) in P. humilis and P. lounsburyi, and in ND6 in P. concolor (89.9%). The three Psyttalia had strand asymmetry with negative AT-skew (average = −0.06) and positive GC-skew (average = 0.19), similarly to the trend in other the 28 Braconidae (average AT-skew = −0.04, average CG-skew = 0.15) (Table 4). All genes in Psyttalia had negative AT-skews (PCG average = −0.07, tRNA average = −0.01 and rRNA average = −0.08), and positive GC-skews (PCG average = 0.20, tRNA average = 0.13 and rRNA average = 0.09). PCGs on the J-strand had negative AT-skew (−0.17) and positive GC-skew (average = 0.23), and PCGs on the N-strand had positive AT-skew (average = 0.83) and positive GC-skew (average = 0.14). Strand compositional bias (strand asymmetry) is frequent in insect mitogenomes, and is presumed to be the result of a prolonged single-stranded state of either the J-strand or the N-strand during transcription and replication, exposing one of the strands to a higher chance of DNA damage and repair. Exposed single-stranded DNA has a greater probability of deamination of C and A nucleotides, resulting in greater frequencies of C and A content on the complementary strand [36]. Consequently, positive AT-skew and negative GC-skew are usually observed on the J-strand. However, in some arthropods strand asymmetry is reversed, with negative AT-skews and positive GC-skews on the J-strand [58,59,60,61,62]. Psyttalia had strand asymmetry reversal on the J-strand, as had all other species in our dataset except Proterops sp. This feature may be explained by the inversion of the replication of origin in the AT-rich region in some insect lineages, as demonstrated previously [38].

3.4. tRNA Genes and Mitochondrial Gene Rearrangements in Braconidae

3.4.1. tRNA Structure and Anticodons

The positions and structure of the tRNAs predicted by ARWEN and MITOS were identical. All tRNAs were predicted to fold into a cloverleaf structure except tRNASer1, for which the dihydrouridine (DHU) arm was reduced to a simple loop, a frequent occurrence in metazoans [63]. tRNALys and tRNASer2 used the TTT and TCT anticodons instead of the regular CTT and GCT, respectively. In our dataset of mitogenomes, tRNALys was annotated in all species and all used the TTT anticodon, except Diadegma semiclausum and Enicospilus sp. which used the regular CTT. tRNASer2 used the TCT anticodon in all species for which the gene was annotated (27/33). The usage of irregular anticodons in these two tRNAs could be associated with gene rearrangements [64].

3.4.2. tRNA Rearrangements in Braconidae

Comparative mitogenomics have shown that gene rearrangements are infrequent in closely related taxa, and have a low likelihood of convergence owing to the large number of different possible combinations [65,66]. As such, mitochondrial gene rearrangements can be useful for resolving ancient evolutionary relationships [65,67]. The three Psyttalia shared the same gene arrangement but several differences relative to the other cyclostomes were present, all involving tRNAs (Figure 3). Twelve of the tRNA genes were organized into three clusters: D-H-K, A-R-N-S1-E-F and W-Y-C (genes in the N strand are underlined). In contrast, the ancestral insect mitogenome [68] is thought to have the A-R-N-S1-E-F, I-Q-M and W-C-Y organization, and the COX2 and ATP8 junction is COX2-K-D-ATP8. All Psyttalia had the derived state COX2-D-H-K-ATP8, with D inverted and having switched positions with K, and H inverted and translocated from its original position between ND5 and ND4 to its new location between D and K. Interestingly, H is found between COX2 and ATP8 in all cyclostomes except Aphidius gifuensis and Histeromerus sp., which occupy the most basal positions in the phylogeny of the clade (see below), suggesting that the translocation took place ~90 MYA. In congruence with the phylogeny, the basal A. gifuensis has preserved the ancestral K-D gene order, whereas Histeromerus sp. has the D-K arrangement. The different derived state COX2-D-K-H-ATP8, found in Pambolus sp., is therefore likely due to a subsequent exchange of positions between K and H.
In insects, the ancestral I-Q-M cluster is situated between the AT-rich region and ND2. All Psyttalia had the derived state I-M-AT-rich region-Q, with I and M inverted and translocated to their new position between 12S rRNA and the AT-rich region, a state which is shared with S. agrili and C. vestalis. Furthermore, the same topology might be present in D. longicaudata and A. gifuensis as they both have an inverted I (as well as M, in the case of D. longicaudata) adjacent to 12S rRNA, while the positions of the remaining elements are unknown due to incompleteness of the sequences. Interestingly, Ichneumonidae, the sister group to Braconidae, have the intermediate topology AT-rich region-I-M-Q. This suggests that the ancestral state in the common ancestor of Braconid–Ichneumonid is the state found in Ichneumonidae and raises the possibility that the srRNA-I-M gene arrangement is a synapomorphy of Braconidae.
In the insect ancestral mitogenome, the W-C-Y cluster is situated between ND2 and COX1. All Psyttalia had the derived state W-Y-C, in that C and Y swapped positions. This state is also found in species of Ichneumonidae, Pteromalidae and Eulophidae, while Orussidae and Vespidae have the ancestral state. If the W-Y-C state found in these different families had a common origin, it would conflict with a recent phylogenetic reconstruction of Hymenoptera [52]. However, a common origin seems unlikely as the ancestral state is conserved in the non-cyclostome C. vestalis, while a different derived state C-W-Y was found in S. agrili. As the sequence of this region has not been reported for the remaining 26 Braconidae included in our study, it is presently impossible to determine if the W-Y-C state found in Psyttalia is unique within the family. Remarkably, all Psyttalia have a unique rearrangement: the swapping of positions between P and T, which are situated between ND4L and ND6. This arrangement is not present in D. longicaudata; therefore, the feature could be specific to the genus Psyttalia.

3.5. Phylogenetic Position of Psyttalia within Braconidae

Phylogenetic analyses were performed primarily to determine the position of Psyttalia within Braconidae, and to obtain an estimate of the divergence times between the three species. The reconstruction was based on the set of 15 partitions from 11 PCGs used by Li et al. (2016) [37]. The Psyttalia species clustered with D. longicaudata, the only (partially) sequenced member of the subfamily Opiinae (Figure 4). The placement of Psyttalia was robust, as it was insensitive to the use of different datasets and partition-clustering schemes. The divergence between Psyttalia and D. longicaudata was estimated at ~55 MYA (95% HPD: 34–83 MYA). Psyttalia lounsburyi occupied the most basal position among the three Psyttalia, having diverged from the other two species ~11 MYA (95% HPD: 6–17 MYA). Psyttalia concolor and P. humilis were recovered as sister species, having diverged ~2 MYA (95% HPD: 1.1–3.6 MYA). These results support the taxonomic classification of P. concolor and P. humilis as distinct species, despite their high morphological similarity. These divergence times are also interesting from the perspective of gene rearrangements. Indeed, either a single (swapping of positions between the P and T tRNA genes) or two (if the swapping of positions between Y and C tRNAs is restricted to Psyttalia) rearrangements occurred between the divergence of D. longicaudata and the ancestor of Psyttalia, and none in the 11 MY after the divergence of the three species, suggesting that the timescale for gene rearrangements in Braconidae is in the order of tens of millions of years. Clarification of this matter will require a denser taxonomic coverage of mitogenomes of Braconidae.
Li et al. (2006) reported topological variations depending on the data matrices and analytical methods used in their analyses that included all available Braconidae mitogenomes. As such variations could potentially affect the phylogenetic position of Psyttalia, we conducted additional analyses using a data matrix containing 24 gene and codon partitions and/or different partition-clustering schemes. Some of the alternative analyses resulted in topological alterations, but none involved any of the Opiinae species (Figure S1). Surprisingly, the monophyly of the cyclostomes was sensitive to partition-clustering but not to the data matrix. Aphidius gifuensis was recovered as the most basal subfamily within cyclostomes when using the scheme selected by PartitionFinder, but it occupied the most basal position among all Braconidae in the trees obtained with the alternative partition-clustering schemes. Our analyses confirmed that some of the relationships among subfamilies of Braconidae are not recovered robustly, as previously reported [37]. The monophyly of cyclostomes was not robust, as the single available representative of Aphidiinae (A. gifuensis) was recovered as either basal within cyclostomes or basal within Braconidae, depending on the partition-clustering scheme used. This inconsistency and the smaller instabilities observed in the placement of other subfamilies should not be interpreted as evidence for the paraphyly of cyclostomes. Most probably, they reflect the difficulty in resolving phylogenies involving large time scales (almost 120 MYA) using mitochondrial sequences when only a small number of taxa is available. This limitation highlights the importance of increasing the mitogenomic coverage of Braconidae, particularly in subfamilies such as Aphidiinae, for which a robust phylogenetic position could not be determined at this point.

Supplementary Materials

The following are available online at https://www.mdpi.com/2075-4450/11/12/854/s1, Figure S1: Bayesian phylogenetic reconstruction among Braconidae obtained with different datasets and partition clustering schemes., Table S1: Primers and cycling protocols used for polymerase chain reaction (PCR) amplification and Sanger sequencing (Seq) of the complete mitochondrial genome of Psyttalia concolor.

Author Contributions

B.v.A. and L.T.d.C. conceived and designed the study. V.C. performed sample collection and morphological identification of specimens. L.T.d.C., C.P. and C.R. performed laboratory work and analyzed the data. All authors contributed to the interpretation of the results and the writing of the manuscript. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by Stellenbosch University (Staff Research Grant and SU Open Access Fund).

Acknowledgments

The authors are grateful to olive farmers and landowners for allowing the collection of samples on their properties.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Shi, M.; Chen, X.X.; Van Achterberg, C. Phylogenetic relationships among the Braconidae (Hymenoptera: Ichneumonoidea) inferred from partial 16S rDNA, 28S rDNA D2, 18S rDNA gene sequences and morphological characters. Mol. Phylogenet. Evol. 2005, 37, 104–116. [Google Scholar] [CrossRef]
  2. Yu, D.S.; Van Achterberg, C.; Horstmann, K. World Ichneumonoidea 2011. In Taxonomy, Biology, Morphology and Distribution (Braconidae). Taxapad (Scientific Names for Information Management) Interactive Catalogue on DVD/CDROM; Vancouver, BC, Canada; 2012. [Google Scholar]
  3. Dowton, M. Relationships among the cyclostome braconid (Hymenoptera: Braconidae) subfamilies inferred from a mitochondrial tRNA gene rearrangement. Mol. Phylogenet. Evol. 1999. [Google Scholar] [CrossRef] [PubMed]
  4. Dowton, M.; Belshaw, R.; Austin, A.D.; Quicke, D.L.J. Simultaneous molecular and morphological analysis of braconid relationships (Insecta: Hymenoptera: Braconidae) indicates independent mt-tRNA gene inversions within a single wasp family. J. Mol. Evol. 2002, 54, 210–226. [Google Scholar] [CrossRef] [PubMed]
  5. Achterberg, C. van Apanteles (Choeras) gielisi spec. nov. (Hymenoptera: Braconidae: Microgastrinae) from The Netherlands and the first report of Trichoptera as host of Braconidae. Zool. Meded. 2002, 76, 53–60. [Google Scholar]
  6. Wharton, R.A.; Shaw, S.R.; Sharkey, M.J.; Wahl, D.B.; Woolley, J.B.; Whitfield, J.B.; Marsh, P.M.; Johnson, W. Phylogeny of the subfamilies of the family Braconidae (Hymenoptera: Ichneumonoidea): A reassessment. Cladistics 1992. [Google Scholar] [CrossRef]
  7. Gauld, I.D. Evolutionary patterns of host utilization by ichneumonoid parasitoids (Hymenoptera: Ichneumonidae and Braconidae). Biol. J. Linn. Soc. 1988, 35, 351–377. [Google Scholar] [CrossRef]
  8. Zaldivar-Riverón, A.; Mori, M.; Quicke, D.L.J. Systematics of the cyclostome subfamilies of braconid parasitic wasps (Hymenoptera: Ichneumonoidea): A simultaneous molecular and morphological Bayesian approach. Mol. Phylogenet. Evol. 2006, 38, 130–145. [Google Scholar] [CrossRef]
  9. Belshaw, R.; Fitton, M.; Herniou, E.; Gimeno, C.; Quicke, D.L.J. A phylogenetic reconstruction of the Ichneumonoidea (Hymenoptera) based on the D2 variable region of 28S ribosomal RNA. Syst. Entomol. 1998, 23, 109–123. [Google Scholar] [CrossRef]
  10. Belshaw, R.; Quicke, D.L.J. A Molecular Phylogeny of the Aphidiinae (Hymenoptera:Braconidae). Mol. Phylogenet. Evol. 1997, 7, 281–293. [Google Scholar] [CrossRef]
  11. Dowton, M.; Austin, A.D.; Antolin, M.F. Evolutionary relationships among the Braconidae (Hymenoptera: Ichneumonoidea) inferred from partial 16S rDNA gene sequences. Insect Mol. Biol. 1998, 7, 129–150. [Google Scholar] [CrossRef]
  12. Rice, R.E.; Phillips, P.A.; Stewart-leslie, J.; Sibbett, G.S. Olive fruit fly populations measured in Central and Southern California. Calif. Agric. 2003, 57, 122–127. [Google Scholar] [CrossRef]
  13. Hladnik, M. A review of plant protection against the olive fly (Bactrocera oleae (Rossi, 1790) Gmelin) and molecular methods to monitor the insecticide resistance alleles. Acta Agric. Slov. 2017, 109, 135–146. [Google Scholar] [CrossRef]
  14. Vontas, J.G.; Hejazi, M.J.; Hawkes, N.J.; Cosmidis, N.; Loukas, M.; Hemingway, J. Resistance-associated point mutations of organophosphate insensitive acetylcholinesterase, in the olive fruit fly Bactrocera oleae. Insect Mol. Biol. 2002. [Google Scholar] [CrossRef] [PubMed]
  15. Kakani, E.G.; Ioannides, I.M.; Margaritopoulos, J.T.; Seraphides, N.A.; Skouras, P.J.; Tsitsipis, J.A.; Mathiopoulos, K.D. A small deletion in the olive fly acetylcholinesterase gene associated with high levels of organophosphate resistance. Insect Biochem. Mol. Biol. 2008. [Google Scholar] [CrossRef] [PubMed]
  16. Vontas, J.G.; Cosmidis, N.; Loukas, M.; Tsakas, S.; Hejazi, M.J.; Ayoutanti, A.; Hemingway, J. Altered acetylcholinesterase confers organophosphate resistance in the olive fruit fly Bactrocera oleae. Pestic. Biochem. Physiol. 2001. [Google Scholar] [CrossRef]
  17. Pereira-Castro, I.; Van Asch, B.; Trinidade Rei, F.; Texeira Da Costa, L. Bactrocera oleae (Diptera: Tephritidae) organophosphate resistance alleles in Iberia: Recent expansion and variable frequencies. Eur. J. Entomol. 2014, 112, 20–26. [Google Scholar] [CrossRef]
  18. Daane, K.M.; Johnson, M.W. Olive Fruit Fly: Managing an Ancient Pest in Modern Times Olive Fruit Fly: Managing an Ancient Pest in Modern Times. Annu. Rev. Entomol. 2010, 55, 157–169. [Google Scholar] [CrossRef]
  19. Silvestri, F. Report of an Expedition to Africa in Search of the Natural Enemies of Fruit Flies (Trypaneidae) with Descriptions, Observations and Biological Notes. Hawaii Board Agric. For. Div. Entomol. Bull. 1914, 3, 1–146. [Google Scholar]
  20. Powell, C.; Caleca, V.; Sinno, M.; van Staden, M.; van Noort, S.; Rhode, C.; Allsopp, E.; van Asch, B. Barcoding of parasitoid wasps (Braconidae and Chalcidoidea) associated with wild and cultivated olives in the Western Cape of South Africa. Genome 2019. [Google Scholar] [CrossRef]
  21. Rugman-Jones, P.F.; Wharton, R.; van Noort, T.; Stouthamer, R. Molecular differentiation of the Psyttalia concolor (Szépligeti) species complex (Hymenoptera: Braconidae) associated with olive fly, Bactrocera oleae (Rossi) (Diptera: Tephritidae), in Africa. Biol. Control 2009, 49, 17–26. [Google Scholar] [CrossRef]
  22. Copeland, R.S. A new species of Munromyia Bezzi (Diptera: Tephritidae) reared from Chionanthus battiscombei (Oleaceae) in Northern Kenya. J. Nat. Hist. 2009. [Google Scholar] [CrossRef]
  23. Teixeira da Costa, L.; Powell, C.; van Noort, S.; Costa, C.; Sinno, M.; Caleca, V.; Rhode, C.; Kennedy, R.J.; van Staden, M.; van Asch, B. The complete mitochondrial genome of Bactrocera biguttula (Bezzi) (Diptera: Tephritidae) and phylogenetic relationships with other Dacini. Int. J. Biol. Macromol. 2019. [Google Scholar] [CrossRef] [PubMed]
  24. Mkize, N.; Hoelmer, K.A.; Villet, M.H. A survey of fruit-feeding insects and their parasitoids occurring on wild olives, Olea europaea ssp. cuspidata, in the Eastern Cape of South Africa. Biocontrol Sci. Technol. 2008, 18, 991–1004. [Google Scholar] [CrossRef]
  25. Copeland, R.; White, I.; Okumu, M.; Machera, P.; Wharton, R. Insects Associated with Fruits of the Oleaceae (Asteridae, Lamiales) in Kenya, with Special Reference to the Tephritidae (Diptera). Bish. Museum Bull. Entomol. 2004, 12, 135–164. [Google Scholar]
  26. Wharton, R.; Gilstrap, F. Key to and Status of Opiine Braconid (Hymenoptera) Parasitoids Used in Biological Control of Ceratitis and Dacus s. l. (Diptera: Tephritidae). Ann. Entomol. Soc. Am. 1983, 76, 721–742. [Google Scholar] [CrossRef]
  27. Silvestri, F. La lotta biologica contro le mosche dei frutti della famiglia Trypetidae. In Proceedings of the VII International Congress of Entomology, Berlin, Germany, 17–23 August 1939; pp. 2396–2418. [Google Scholar]
  28. Caleca, V.; Giacalone, C.; Maltese, M.; Tortorici, F. Contenimento naturale di Bactrocera oleae (Rossi): Clima o parassitoidi? Confronto tra Western Cape (Sud Africa) e Sicilia. Atti Acc. Naz. Ital. Entomol. 2017, 64, 99–105. [Google Scholar]
  29. Raspi, A.; Loni, A.; Canovai, R. Entomophages of olive pests in corsica, coastal tuscany and islands of Tuscan Archipelago. Frustula Entomol. 2007, 30, 187–194. [Google Scholar]
  30. Debouzie, D.; Mazih, A. Argan (Sapotaceae) trees as reservoirs for Mediterranean fruit fly (Diptera: Tephritidae) in Morocco. Environ. Entomol. 1999. [Google Scholar] [CrossRef]
  31. Monastero, S.; Delanoue, P. Lute biologique expérimentale contre la Mouche de l’olive (Dacus OleaeGmel.) au moyen D’Opius concolorSzepl.SiculusMon. dans les iles éoliennes (sicile) en 1965. Entomophaga 1966. [Google Scholar] [CrossRef]
  32. Liaropoulus, C.; Mavraganis, V.G.; Broumas, T.; Ragoussis, N. Field tests on the combination of mass trapping with the release parasite Opius concolor (Hymenoptera: Braconidae), for the control of the olive fruit fly Bactrocera oleae (Diptera: Tephritidae). IOBC/WPRS Bull. 2005, 28, 77–81. [Google Scholar]
  33. Daane, K.M.; Sime, K.R.; Wang, X.; Nadel, H.; Johnson, M.W.; Walton, V.M.; Kirk, A.; Pickett, C.H. Psyttalia lounsburyi (Hymenoptera: Braconidae), potential biological control agent for the olive fruit fly in California. Biol. Control 2008, 44, 79–89. [Google Scholar] [CrossRef]
  34. Daane, K.M.; Johnson, M.W.; Pickett, C.H.; Sime, K.R.; Wang, X.G.; Nadel, H.; Andrews, J.W.; Hoelmer, K.A. Biological controls investigated to aid management of olive fruit fly in California. Calif. Agric. 2011, 65, 21–28. [Google Scholar] [CrossRef]
  35. Cameron, S.L. Insect mitochondrial genomics: Implications for evolution and phylogeny. Annu. Rev. Entomol. 2014, 59, 95–117. [Google Scholar] [CrossRef] [PubMed]
  36. Wei, S.-j.; Shi, M.; Sharkey, M.J.; van Achterberg, C.; Chen, X.-x. Comparative mitogenomics of Braconidae (Insecta: Hymenoptera) and the phylogenetic utility of mitochondrial genomes with special reference to Holometabolous insects. BMC Genom. 2010, 11, 371. [Google Scholar] [CrossRef]
  37. Li, Q.; Wei, S.J.; Tang, P.; Wu, Q.; Shi, M.; Sharkey, M.J.; Chen, X.X. Multiple lines of evidence Frommitochondrial genomes resolve phylogenetic relationships of parasitic wasps in braconidae. Genome Biol. Evol. 2016, 8, 2651–2662. [Google Scholar] [CrossRef]
  38. Wei, S.J.; Shi, M.; Chen, X.X.; Sharkey, M.J.; van Achterberg, C.; Ye, G.Y.; He, J.H. New views on strand asymmetry in insect mitochondrial genomes. PLoS ONE 2010, 5, e12708. [Google Scholar] [CrossRef]
  39. Dowton, M.; Cameron, S.L.; Dowavic, J.I.; Austin, A.D.; Whiting, M.F. Characterization of 67 mitochondrial tRNA gene rearrangements in the hymenoptera suggests that mitochondrial tRNA gene position is selectively neutral. Mol. Biol. Evol. 2009. [Google Scholar] [CrossRef]
  40. Xiao, J.H.; Jia, J.G.; Murphy, R.W.; Huang, D.W. Rapid evolution of the mitochondrial genome in chalcidoid wasps (hymenoptera: Chalcidoidea) driven by parasitic lifestyles. PLoS ONE 2011, 6, e26645. [Google Scholar] [CrossRef]
  41. Silvestri, F. Viaggio in Eritrea per Cercare Parassiti della Mosca delle Olive. Boll. Lab. Zool. Gen. Agrar. 1913, 9, 186–226. [Google Scholar]
  42. Kimura, M. A simple method for estimating evolutionary rates of base substitutions through comparative studies of nucleotide sequences. J. Mol. Evol. 1980, 16, 111–120. [Google Scholar] [CrossRef]
  43. Kumar, S.; Stecher, G.; Li, M.; Knyaz, C.; Tamura, K. MEGA X: Molecular evolutionary genetics analysis across computing platforms. Mol. Biol. Evol. 2018, 35, 1547–1549. [Google Scholar] [CrossRef] [PubMed]
  44. Folmer, O.; BLACK, M.; HOEH, W.; Lutz, R.; Vrijenhoek, R. DNA primers for amplification of mitochondrial cytochrome c oxidase subunit I from diverse metazoan invertebrates. Mol. Mar. Biol. Biotechnol. 1994, 3, 294–299. [Google Scholar] [CrossRef] [PubMed]
  45. Smith, D.R. The past, present and future of mitochondrial genomics: Have we sequenced enough mtDNAs? Brief. Funct. Genom. 2016, 15, 47–54. [Google Scholar] [CrossRef] [PubMed]
  46. Laslett, D.; Canbäck, B. ARWEN: A program to detect tRNA genes in metazoan mitochondrial nucleotide sequences. Bioinformatics 2008, 24, 172–175. [Google Scholar] [CrossRef] [PubMed]
  47. Abascal, F.; Zardoya, R.; Telford, M.J. TranslatorX: Multiple alignment of nucleotide sequences guided by amino acid translations. Nucleic Acids Res. 2010, 38. [Google Scholar] [CrossRef] [PubMed]
  48. Kumar, S.; Stecher, G.; Tamura, K. MEGA7: Molecular Evolutionary Genetics Analysis Version 7.0 for Bigger Datasets. Mol. Biol. Evol. 2016, 33, 1870–1874. [Google Scholar] [CrossRef]
  49. Lanfear, R.; Frandsen, P.B.; Wright, A.M.; Senfeld, T.; Calcott, B. Partitionfinder 2: New methods for selecting partitioned models of evolution for molecular and morphological phylogenetic analyses. Mol. Biol. Evol. 2017, 34, 772–773. [Google Scholar] [CrossRef]
  50. Miller, M.A.; Pfeiffer, W.; Schwartz, T. Creating the CIPRES Science Gateway for inference of large phylogenetic trees. In Proceedings of the 2010 Gateway Computing Environments Workshop (GCE), New Orleans, LA, USA, 14 November 2010; pp. 1–8. [Google Scholar]
  51. Drummond, A.J.; Rambaut, A. BEAST: Bayesian evolutionary analysis by sampling trees. BMC Evol. Biol. 2007, 7. [Google Scholar] [CrossRef]
  52. Branstetter, M.G.; Danforth, B.N.; Pitts, J.P.; Faircloth, B.C.; Ward, P.S.; Buffington, M.L.; Gates, M.W.; Kula, R.R.; Brady, S.G. Phylogenomic Insights into the Evolution of Stinging Wasps and the Origins of Ants and Bees. Curr. Biol. 2017. [Google Scholar] [CrossRef]
  53. Nardi, F.; Carapelli, A.; Boore, J.L.; Roderick, G.K.; Dallai, R.; Frati, F. Domestication of olive fly through a multi-regional host shift to cultivated olives: Comparative dating using complete mitochondrial genomes. Mol. Phylogenet. Evol. 2010, 57, 678–686. [Google Scholar] [CrossRef]
  54. Darriba, D.; Taboada, G.L.; Doallo, R.; Posada, D. jModelTest 2: More models, new heuristics and parallel computing. Nat. Methods 2012, 9, 772. [Google Scholar] [CrossRef] [PubMed]
  55. Coates, B.S. Assembly and annotation of full mitochondrial genomes for the corn rootworm species, Diabrotica virgifera virgifera and Diabrotica barberi (Insecta: Coleoptera: Chrysomelidae), using Next Generation Sequence data. Gene 2014, 542, 190–197. [Google Scholar] [CrossRef] [PubMed]
  56. Castro, L.R.; Ruberu, K.; Dowton, M. Mitochondrial genomes of Vanhornia eucnemidarum (Apocrita: Vanhorniidae) and Primeuchroeus spp. (Aculeata: Chrysididae): Evidence of rearranged mitochondrial genomes within the Apocrita (Insecta: Hymenoptera). Genome 2006, 49, 752–766. [Google Scholar] [CrossRef] [PubMed]
  57. Castro, L.R.; Dowton, M. The position of the Hymenoptera within the Holometabola as inferred from the mitochondrial genome of Perga condei (Hymenoptera: Symphyta: Pergidae). Mol. Phylogenet. Evol. 2005, 34, 469–479. [Google Scholar] [CrossRef] [PubMed]
  58. Cameron, S.L.; Dowton, M.; Castro, L.R.; Ruberu, K.; Whiting, M.F.; Austin, A.D.; Diement, K.; Stevens, J. Mitochondrial genome organization and phylogeny of two vespid wasps. Genome 2008, 51, 800–808. [Google Scholar] [CrossRef]
  59. Hassanin, A.; Léger, N.; Deutsch, J. Evidence for multiple reversals of asymmetric mutational constraints during the evolution of the mitochondrial genome of metazoa, and consequences for phylogenetic inferences. Syst. Biol. 2005, 54, 277–298. [Google Scholar] [CrossRef]
  60. Kilpert, F.; Podsiadlowski, L. The complete mitochondrial genome of the common sea slater, Ligia oceanica (Crustacea, Isopoda) bears a novel gene order and unusual control region features. BMC Genom. 2006, 7, 241. [Google Scholar] [CrossRef]
  61. Masta, S.E.; Longhorn, S.J.; Boore, J.L. Arachnid relationships based on mitochondrial genomes: Asymmetric nucleotide and amino acid bias affects phylogenetic analyses. Mol. Phylogenet. Evol. 2009, 50, 117–128. [Google Scholar] [CrossRef]
  62. Hassanin, A. Phylogeny of Arthropoda inferred from mitochondrial sequences: Strategies for limiting the misleading effects of multiple changes in pattern and rates of substitution. Mol. Phylogenet. Evol. 2006, 38, 100–116. [Google Scholar] [CrossRef]
  63. Jühling, F.; Pütz, J.; Bernt, M.; Donath, A.; Middendorf, M.; Florentz, C.; Stadler, P.F. Improved systematic tRNA gene annotation allows new insights into the evolution of mitochondrial tRNA structures and into the mechanisms of mitochondrial genome rearrangements. Nucleic Acids Res. 2012, 40, 2833–2845. [Google Scholar] [CrossRef]
  64. Wei, S.J.; Shi, M.; He, J.H.; Sharkey, M.; Chen, X.X. The complete mitochondrial genome of Diadegma semiclausum (Hymenoptera: Ichneumonidae) indicates extensive independent evolutionary events. Genome 2009, 52, 308–319. [Google Scholar] [CrossRef] [PubMed]
  65. Boore, J.; Collins, T.; Stanton, D.; Daehler, L.; Brown, W. Deducing the pattern of arthropod phylogeny from mitochondrial DNA rearrangements. Nature 1995, 376, 163. [Google Scholar] [CrossRef] [PubMed]
  66. Moritz, C.; Dowling, T.E.; Brown, W.M. Evolution of animal mitochondrial DNA: Relevance for population biology and systematics. Annu. Rev. Ecol. Syst. 1987, 18, 269–292. [Google Scholar] [CrossRef]
  67. Rokas, A.; Holland, P.W.H. Rare genomic changes as a tool for phylogenetics. Trends Ecol. Evol. 2000, 15, 454–459. [Google Scholar] [CrossRef]
  68. Boore, J.L. Animal mitochondrial genomes. Nucleic Acids Res. 1999, 27, 1767–1780. [Google Scholar] [CrossRef] [PubMed]
Figure 1. Adult specimens representative of three Psyttalia (Hymenoptera: Braconidae) species. (A) Psyttalia concolor male (scale bar = 1 mm); (B) Psyttalia humilis male (scale bar = 1 mm); and (C) Psyttalia lounsburyi female (scale bar = 100 μm).
Figure 1. Adult specimens representative of three Psyttalia (Hymenoptera: Braconidae) species. (A) Psyttalia concolor male (scale bar = 1 mm); (B) Psyttalia humilis male (scale bar = 1 mm); and (C) Psyttalia lounsburyi female (scale bar = 100 μm).
Insects 11 00854 g001
Figure 2. Organization of the mitochondrial genomes of Psyttalia concolor, P. humilis and P. lounsburyi (Hymenoptera: Braconidae). Transfer RNA genes are designated by the single-letter amino acid code. Arrows indicate the direction of gene transcription. AT-rich *-putative location of the control region.
Figure 2. Organization of the mitochondrial genomes of Psyttalia concolor, P. humilis and P. lounsburyi (Hymenoptera: Braconidae). Transfer RNA genes are designated by the single-letter amino acid code. Arrows indicate the direction of gene transcription. AT-rich *-putative location of the control region.
Insects 11 00854 g002
Figure 3. Gene arrangements in the mitochondrial genomes of 13 species of cyclostome wasps (Hymenoptera: Braconidae). Genes involved in rearrangements relative to the ancestral are shown in red, underlined. The species are ordered following the cyclostome tree presented in Figure 4.
Figure 3. Gene arrangements in the mitochondrial genomes of 13 species of cyclostome wasps (Hymenoptera: Braconidae). Genes involved in rearrangements relative to the ancestral are shown in red, underlined. The species are ordered following the cyclostome tree presented in Figure 4.
Insects 11 00854 g003
Figure 4. Bayesian reconstruction of the phylogenetic relationships among 31 species in the family Braconidae, with Diadegma semiclausum and Enicosphilus sp. (Ichneumonidae) as outgroups. Nodal support is given as Bayesian posterior probability (only values < 1 are shown). Scale bar represents 20 million years.
Figure 4. Bayesian reconstruction of the phylogenetic relationships among 31 species in the family Braconidae, with Diadegma semiclausum and Enicosphilus sp. (Ichneumonidae) as outgroups. Nodal support is given as Bayesian posterior probability (only values < 1 are shown). Scale bar represents 20 million years.
Insects 11 00854 g004
Table 1. List of the partial and complete mitochondrial sequences used in phylogenetic reconstruction of the family Braconidae for the inference of the evolutionary relationships between Psyttalia concolor, P. humilis and P. lounsburyi and 28 other species of cyclostome and non-cyclostome wasps (Hymenoptera: Braconidae). Diadegma semiclausum and Enicospilus sp. (Hymenoptera: Ichneumonidea) were used as outgroups.
Table 1. List of the partial and complete mitochondrial sequences used in phylogenetic reconstruction of the family Braconidae for the inference of the evolutionary relationships between Psyttalia concolor, P. humilis and P. lounsburyi and 28 other species of cyclostome and non-cyclostome wasps (Hymenoptera: Braconidae). Diadegma semiclausum and Enicospilus sp. (Hymenoptera: Ichneumonidea) were used as outgroups.
Species FamilySubfamilyLineageGenBankReferenceSize (bp)Status
Acanthormius sp.BraconidaeLysiterminaeCyclostomeKF385867.1Li et al., 201613,051Partial
Afrocampsis griseosetosus van Achterberg et Quicke, 1990BraconidaeAcampsohelconinaeNon-cyclostomeKJ412474.1Li et al., 201610,104Partial
Aphidius gifuensis Ashmead, 1906 BraconidaeAphidiinaeCyclostomeGU097658.2Wei et al., 201011,970Partial
Capitonius sp. BraconidaeCenocoeliinaeNon-cyclostomeKF385869.1Li et al., 201613,077Partial
Cardiochiles fuscipennis Szépligeti, 1900BraconidaeCardiochilinaeNon-cyclostomeKF385870.1Li et al., 201614,390Partial
Cotesia vestalis (Haliday, 1834)BraconidaeMicrogastinaeNon-cyclostomeNC_014272.1Wei et al., 201015,543Complete
Diachasmimorpha longicaudata (Ashmead, 1905)BraconidaeOpiinaeCyclostomeGU097655.1Wei et al., 201013,850Partial
Diadegma semiclausum (Hellén, 1949)IchneumonidaeCampopleginae-NC_012708.1Wei et al., 200918,728Complete
Elasmosoma sp. BraconidaeEuphorinaeNon-cyclostomeKJ412470.1Li et al., 201612,326Partial
Enicospilus sp. IchneumonidaeOphioninae-FJ478177.1Dowton et al., 200915,300Partial
Eumacrocentrus sp. BraconidaeHelconinaeNon-cyclostomeKF385872.1Li et al., 201614,080Partial
Euurobracon breviterebrae Watanabe, 1934BraconidaeBraconinae CyclostomeKF385871.1Li et al., 201612,957Partial
Histeromerus sp. BraconidaeHisterominaeCyclostomeKF418765.1Li et al., 201613,168Partial
Homolobus sp. BraconidaeHomolobinaeNon-cyclostomeKF385873.1Li et al., 201613,927Partial
Ichneutes sp. BraconidaeIchneutinaeNon-cyclostomeKF385874.1Li et al., 201613,092Partial
Macrocentrus camphoraphilus He et Chen, 2008BraconidaeMacrocentrinaeNon-cyclostomeGU097656.1Wei et al., 201015,801Partial
Meteorus pulchricornis (Wesmael, 1835)BraconidaeEuphorinaeNon-cyclostomeGU097657.1Wei et al., 201010,186Partial
Mirax sp. BraconidaeMiracinaeNon-cyclostomeKJ412471.1Li et al., 201613,664Partial
Pambolus sp. BraconidaePambolinaeCyclostomeKF385875.1Li et al., 201613,175Partial
Paroligoneurus sp. BraconidaeIchneutinaeNon-cyclostomeKJ412472.1Li et al., 201613,413Partial
Phaenocarpa sp.BraconidaeAlysiinaeCyclostomeKJ412475.1Li et al., 20169981Partial
Phanerotoma flava Ashmead, 1906BraconidaeCheloninaeNon-cyclostomeGU097654.1Wei et al., 201010,171Partial
Proterops sp. BraconidaeIchneutinaeNon-cyclostomeKJ412477.1Li et al., 201612,883Partial
Pselaphanus sp.BraconidaePselaphaninaeNon-cyclostomeKF385876.1Li et al., 201613,204Partial
Pseudognaptodon sp. BraconidaeGnamptodontinaeCyclostomeKJ412473.1Li et al., 201613,190Partial
Psyttalia concolor (Szépligeti, 1910)BraconidaeOpiinaeCyclostomeMW279212This study15,308Partial
Psyttalia humilis (Silvestri, 1913)BraconidaeOpiinaeCyclostomeMW279213This study15,311Partial
Psyttalia lounsburyi (Silvestri, 1913)BraconidaeOpiinaeCyclostomeMW279214This study14,982Partial
Sigalphus bicolor (Cresson, 1880)BraconidaeSigalphinaeNon-cyclostomeKF385878.1Li et al., 201612,744Partial
Spathius agrili Yang, 2005 BraconidaeDoryctinaeCyclostomeNC_014278.1Wei et al., 201015,425Complete
Therophilus festivus Muesebeck, 1953BraconidaeAgathidinaeNon-cyclostomeKF385868.1Li et al., 201614,216Partial
Triraphis sp.BraconidaeRogadinaeCyclostomeKF385877.1Li et al., 201613,162Partial
Xiphozele sp.BraconidaeXiphozelinaeNon-cyclostomeKJ412476.1Li et al., 20169160Partial
Table 2. Main features of the mitochondrial genomes of Psyttalia concolor, P. humilis and P. lounsburyi (Hymenoptera: Braconidae). J-strand–majority strand, N-strand–minority strand; AC–anticodon; IGN–intergenic regions (+) and overlapping nucleotides (−). n. a.—not applicable.
Table 2. Main features of the mitochondrial genomes of Psyttalia concolor, P. humilis and P. lounsburyi (Hymenoptera: Braconidae). J-strand–majority strand, N-strand–minority strand; AC–anticodon; IGN–intergenic regions (+) and overlapping nucleotides (−). n. a.—not applicable.
Psyttalia concolorPsyttalia humilisPsyttalia lounsburyi
Gene/RegionStrandCoordinatesSizeACStartStopIGNCoordinatesSizeACStartStopIGNCoordinatesSizeACStart StopIGN
tRNAGlnN1–7474TTG--0n.a.n.a n.a --n.a.n.a.n.a n.a --n.a
ND2J78–10981021-ATA T--377–10971021-ATAT--n.a.19–10391021-ATAT--n.a.
tRNATrpJ1099–116668TCA--01097–116569TCA--−11039–110769TCA--−1
tRNATyrN1163–122866GTA--−41162–122766GTA--−41105–116864GTA--−3
tRNACysN1228–129164GCA--−11227–129064GCA--−11170–123364GCA--1
COX1J1292–28251534-ATGT--01291–28291539-ATGTAA01234–27721539-ATGTAA0
tRNALeu1J2826–289368TAA--02824–289168TAA--−62767–283569TAA--−6
COX2J2902–3558657-ATA TAA82900–3558657-ATATAA82844–3500657-ATATAA8
tRNAAspN3722–379170GTC--1633726–379570GTA--1683663–373674GTC--162
tRNAHisJ3791–385868GTG--−13795–386268GTG--−13736–380469CAC--−1
tRNALysJ3858–392871TTT--−13862–393271TTT--−13804–387471TTT--−1
ATP8J3929–4084156-ATA TAA03933–4088156-ATATAA03875–4030156-ATTTAA0
ATP6J4063–4752690-ATTTAA−224067–4756690-ATTTAA−224009–4698690-ATTTAA−22
COX3J4762–5550789-ATGTAA94766–5554789-ATGTAA94702–5490789-ATGTAA3
tRNAGlyJ5551–561565GGA--05555–561864TCC--05491–555666TCC--0
ND3J5630–6016387-ATTTAG145633–6019387-ATTTAG145571–5957387-ATTTAG14
tRNAAlaJ6015–607662TGC--−26017–607963TGC--−35955–601763TGC--−3
tRNAArgJ6076–614267TCG---16079–614567ACG--−16017–608367ACG--−1
tRNAAsnJ6136–620267GTT--−76139–620567AAC--−76077–614367GTT--−7
tRNASer1J6200–626667AGA--−36203–626967AGA--−36141–620767AGA--−3
tRNAGluJ6266–633065TTC--−16269–633365TTC--−16207–627165GAA--−1
tRNAPheN6329–639264GAA--−26332–639665GAA--−26720–633465GAA--−2
ND5N6393–80521660-ATA T--06396–80551660-ATAT--−16334–79931660-ATAT--−1
ND4N8079–94011323-ATGTAA268082–94041323-ATGTAA268017–93421326-ATGTAA23
ND4LN9395–9691297-ATTTAA−79398–9694297-ATTTAA79336–9632297-ATTTAA−7
tRNAProN9699–976567TGG--79702–976867TGG--79641–970868TGG--8
tRNAThrJ9766–982964TGT--09769–983264TGT--09709–977264TGT--0
ND6J9841–10,406565-ATGT--119844–10408565-ATGT--119784–10,350567-ATGTAA11
CYTBJ10,407–11,5371131-ATGTAA110,410–11,5401131-ATGTAA110,353–11,4831131-ATGTAA2
tRNASer2J11,536–11,60267TGA--−211,539–11,60567TGA--−211,482–11,54968TGA--−2
ND1N11,601–12,560960-ATTTAA−211,604–12,563960-ATTTAA−211,548–12,507960-ATTTAA−2
tRNALeu2N12,561–12,62666TAG--012,568–12,62962TAG--412,508–12,57467TAG--0
16s RNAN12,627–13,9141288---012,630–13,9181289---012575–13,8521278---0
tRNAValN13,915–13,98066GTA--013,919–13,98466GTA--013,852–13,91665TAC--−1
12s RNAN13,981–14,727747---013,985–14,729745---013,918–14,671754---1
tRNAIleN14,728–14,79164GAT--014,730–14,79364GAT--014,672–14,73564ATC--0
tRNAMetN14,794–14,85966CAT--214,797–14,86266CAT--314,740–14,80667CAT--4
AT-rich region-148,69–15,308449---0n.a.n.a.---0n.a.n.a.----
Table 3. Nucleotide composition of the mitochondrial genomes of Psyttalia concolor, P. humilis and P. lounsburyi (Hymenoptera: Braconidae). PCGs–protein-coding genes. AT-skew = (A − T)/(A + T); GC-skew = (G − C)/(G + C).
Table 3. Nucleotide composition of the mitochondrial genomes of Psyttalia concolor, P. humilis and P. lounsburyi (Hymenoptera: Braconidae). PCGs–protein-coding genes. AT-skew = (A − T)/(A + T); GC-skew = (G − C)/(G + C).
Psyttalia concolorPsyttalia humilisPsyttalia lounsburyi
RegionStrandA%C%G%T%A+T%G+C%AT-SkewGC-SkewA%C%G%T%A+T%G+C%AT-SkewGC-SkewA%C%G%T%A+T%G+C%AT-SkewGC-Skew
COX1J30.29.815.844.074.425.6−0.20.230.69.415.944.174.725.3−0.20.330.710.215.143.974.725.3−0.20.2
COX2J36.17.312.044.480.619.4−0.10.235.37.012.645.180.419.6−0.10.335.67.611.345.581.118.9−0.10.2
ATP8J39.15.15.850.089.110.9−0.10.137.84.55.152.690.49.6−0.20.136.55.14.553.890.49.6−0.2−0.1
ATP6J33.57.88.350.483.916.1−0.20.034.18.07.850.184.215.8−0.20.032.28.68.750.682.817.2−0.20.0
COX3J31.38.614.745.476.723.3−0.20.330.78.715.645.075.724.3−0.20.330.48.615.745.275.724.3−0.20.3
ND2J36.23.27.652.989.110.9−0.20.437.03.47.552.089.011.0−0.20.437.13.47.452.089.110.9−0.20.4
ND3J32.73.910.153.286.014.0−0.20.433.13.910.153.086.014.0−0.20.433.14.810.651.684.715.3−0.20.4
ND5N45.46.29.538.984.315.70.10.244.86.59.739.183.916.10.10.244.76.310.039.083.716.30.10.2
ND4N44.87.010.138.182.917.10.10.244.96.99.838.483.316.70.10.244.47.39.938.482.817.20.10.2
ND4LN49.56.46.138.087.512.50.10.050.26.45.737.787.912.10.1−0.149.26.76.437.786.913.10.10.0
ND6J38.73.76.451.289.910.1−0.10.338.34.26.950.688.911.1−0.10.239.04.26.949.988.911.1−0.10.2
CytBJ33.28.711.646.579.720.3−0.20.133.28.812.245.879.021.0−0.20.233.59.411.945.278.721.3−0.10.1
ND1N45.09.19.936.081.019.00.10.044.39.110.436.380.519.50.10.144.39.310.236.380.519.50.10.0
12s rRNAN39.94.35.250.690.59.5−0.10.139.54.35.650.390.19.9−0.10.139.54.64.950.890.49.6−0.10.0
16s rRNAN41.45.46.247.088.411.6−0.10.141.25.46.147.288.411.6−0.10.142.35.26.943.187.612.40.00.1
All PCGsN+J38.17.110.744.182.217.8−0.10.238.07.110.844.082.018.0−0.10.237.97.510.843.981.818.2−0.10.2
All tRNAs 42.95.97.244.087.013.00.00.142.65.87.143.687.212.80.00.142.85.47.943.886.713.30.00.2
All rRNAs 40.85.05.848.389.110.9−0.10.140.65.05.948.489.011.0−0.10.141.35.06.246.088.711.3−0.10.1
Complete mtDNA 39.46.59.544.684.016.0−0.10.239.36.59.644.583.916.1−0.10.239.06.89.844.483.416.6−0.10.2
Table 4. AT-and GC-skews in the protein-coding genes (PCGs), tRNAs, rRNAs and the AT-rich region in the partial and complete mitochondrial genomes of 31 species in the family Braconidae. AT-skew = (A − T)/(A + T); GC-skew = (G − C)/(G + C).
Table 4. AT-and GC-skews in the protein-coding genes (PCGs), tRNAs, rRNAs and the AT-rich region in the partial and complete mitochondrial genomes of 31 species in the family Braconidae. AT-skew = (A − T)/(A + T); GC-skew = (G − C)/(G + C).
Total SequencePCGsJ-Strand PCGstRNAsrRNAs
SpeciesAT-SkewGC-SkewAT-SkewGC-SkewAT-SkewGC-SkewAT-SkewGC-SkewAT-SkewGC-Skew
Acanthormius sp.−0.110.19−0.110.20−0.230.25−0.020.13−0.140.04
Afrocampsis griseosetosus0.440.31−0.110.34−0.180.34−0.040.350.050.05
Aphidius gifuensis−0.060.05−0.060.07−0.190.12−0.020.00−0.07−0.10
Capitonius sp. −0.070.19−0.070.22−0.200.25−0.010.14−0.09−0.09
Cardiochiles fuscipennis−0.070.18−0.080.22−0.200.25−0.050.10−0.060.01
Cotesia vestalis−0.090.10−0.110.12−0.180.09−0.030.11−0.03−0.11
Diachasmimorpha longicaudata0.090.19−0.100.21−0.220.22−0.030.17−0.020.05
Elasmosoma sp. −0.120.38−0.140.40−0.280.420.050.24−0.070.16
Eumacrocentrus sp. −0.010.05−0.020.07−0.130.100.010.070.03−0.17
Euurobracon breviterebrae−0.110.37−0.130.39−0.280.37−0.030.32−0.090.17
Histeromerus sp. −0.060.16−0.060.19−0.180.19−0.010.04−0.09−0.02
Homolobus sp. −0.060.10−0.060.10−0.180.160.000.09−0.06−0.06
Ichneutes sp. −0.060.21−0.060.24−0.080.25−0.030.10−0.09−0.01
Macrocentrus camphoraphilus−0.050.10−0.060.13−0.170.17−0.010.02−0.04−0.11
Meteorus pulchricornis−0.060.14−0.060.16−0.200.20−0.020.15--
Mirax sp. −0.070.19−0.070.02−0.190.26−0.040.22−0.09−0.06
Pambolus sp. −0.090.16−0.100.17−0.220.20−0.050.09−0.080.00
Paroligoneurus sp. −0.120.22−0.120.25−0.240.25−0.040.08−0.150.00
Phaenocarpa sp.−0.090.11−0.100.14−0.190.170.000.17−0.08−0.08
Phanerotoma flava−0.070.28−0.070.29−0.180.30−0.010.15--
Proterops sp. 0.06−0.150.07−0.14−0.04−0.100.030.000.06−0.15
Pselaphanus sp.−0.030.04−0.030.08−0.150.120.020.01−0.05−0.17
Pseudognaptodon sp. −0.020.03−0.020.04−0.160.12−0.030.11−0.03−0.18
Psyttalia concolor−0.060.19−0.070.20−0.170.23−0.010.10−0.080.07
Psyttalia humilis−0.060.19−0.070.21−0.170.24−0.010.10−0.090.08
Psyttalia lounsburyi−0.060.18−0.070.18−0.170.20−0.010.19−0.050.11
Therophilus festivus−0.030.02−0.020.05−0.150.090.010.01−0.06−0.13
Triraphis sp.−0.120.19−0.120.21−0.250.19−0.060.18−0.17−0.11
Sigalphus bicolor−0.030.00−0.020.02−0.150.08−0.020.07−0.04−0.18
Spathius agrili−0.070.19−0.070.20−0.180.23−0.040.13−0.120.01
Xiphozele sp.−0.01−0.05−0.02−0.02−0.090.010.02−0.020.03−0.27
Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Back to TopTop