Synopeas maximum Awad &amp; Talamas (Hymenoptera, Platygastridae): a new species of parasitoid associated with soybean gall midge, Resseliella maxima Gagné (Diptera, Cecidomyiidae)

Gloria Melotto; Jessica Awad; Elijah J. Talamas; Robert L. Koch; Amelia R. I. Lindsey

doi:10.3897/jhr.96.102865

Abstract

Synopeas maximum Awad & Talamas, sp. nov., the first reported parasitoid associated with the soybean gall midge, Resseliella maxima Gagné, is described based on morphological and molecular data. Parasitoids were reared from soybean stems infested by R. maxima in Minnesota. A phylogenetic reconstruction of the genus Synopeas Förster was performed with COI sequences (n=2412) available on the Barcode of Life Data System (BOLD). Phylogenetic and barcode gap analyses suggest 279 Synopeas species in this dataset, with S. maximum sequences forming a monophyletic clade that is distinct from relatives. The Synopeas maximum clade was close to specimens from Canada and the United States, suggesting it is native to North America. We present a taxonomic treatment of S. maximum to facilitate its identification, including comparison to morphologically similar species. This project provides baseline data for further ecological study of R. maxima parasitism, and its management in soybean.

Keywords

gall midge, parasitism, phylogenetic reconstruction, soybean

Introduction

Gall midges (Diptera: Cecidomyiidae) are a hyper-diverse lineage, representing 30% of dipteran diversity in some ecosystems (Huang et al. 2022). Similarly diverse are parasitoids in the subfamily Platygastrinae (Hymenoptera: Platygastridae), (Chen et al. 2021). Platygastrines are larval or egg-larval parasitoids of Cecidomyiidae, and development of the parasitoid is often suspended until the host is in the last instar or prepupal stage (Kim et al. 2011; Abram et al. 2012; Chen et al. 2021). For both Cecidomyiidae and Platygastrinae, the diversity of species far exceeds what has been described (Srivathsan et al. 2022). However, a recent treatment of the genus Synopeas from Papua New Guinea, using morphology, DNA barcoding, and available data on host associations, has set a new standard for making inroads into both the taxonomy and ecology of gall midge parasitoids (Awad et al. 2021). The diversity of species and trophic interactions between Cecidomyiidae and Platygastrinae are large and complex. However, agricultural ecosystems can provide excellent opportunities to investigate these relationships within much simpler systems.

In 2018, larvae of an unknown species of cecidomyiid were found associated with dying soybean plants, Glycine max (L.) Merr., in the midwestern United States (Gagné et al. 2019). In 2019, this species of unknown origin was described as Resseliella maxima Gagné, the soybean gall midge (Gagné et al. 2019). Soybean plants become susceptible to R. maxima attack when the plants have two or three expanded trifoliate leaves, which is when the plant stems present natural fissures below the cotyledonary node (McMechan et al. 2021). Adult female midges oviposit in these fissures and the larvae feed within the stem at the base of the plant, resulting in necrotic lesions that can cause lodged, wilted, and dead plants (McMechan et al. 2021; Helton et al. 2022).

Efforts to reduce R. maxima injury to soybeans have focused primarily on chemical control, but have so far provided insufficient protection of soybean plants (Hodgson and Helton 2021; McMechan 2021). Therefore, additional tactics for more effective and sustainable management are needed. One potential management approach is biological control, which has yet to be examined for R. maxima. Other pestiferous Cecidomyiidae have been successfully managed using biological control agents, including Aprostocetus epicharmus (Walker) (Hymenoptera: Eulophidae) that parasitized up to 38% of raspberry cane midge, Resseliella theobaldi (Barnes) (Vétek et al. 2006), and Synopeas myles (Walker) (Hymenoptera: Platygastridae) that parasitized up to 28% of swede midge, Contarinia nasturtii (Kieffer) (Abram et al. 2012b; Ferland 2020).

The present work investigates potential parasitism of R. maxima by rearing parasitoids from field-collected soybean stems. We present a taxonomic and molecular description of the R. maxima-associated parasitoid, Synopeas maximum Awad & Talamas, sp. nov., and a phylogenetic analysis of Synopeas sequences available on the Barcode of Life Data System (BOLD).

Methods

Field collection and emergence cages

Soybean stems presenting symptoms of infestation by R. maxima (i.e., darkened swollen lesions at the base of the stems) were collected during the summer of 2021 in two fields on one farm near the city of Luverne (Rock County), Minnesota, USA. Field collection started on 30 June 2021, when soybean plants started to show symptoms of infestation, and continued every other week until R. maxima larval infestation was no longer detected on 01 September 2021. On each sampling date, 10 randomly selected symptomatic plants were collected per eight sampling locations per field by pulling the entire plant from the soil. These plants were trimmed above the first pair of unifoliate leaves, placed in zipper-locking plastic bags (17.7 × 18.8 cm, Ziploc), and held in coolers until brought to the laboratory (approximately five hours).

In the laboratory, the stems were prepared for placement in emergence cages. The cut end of each stem was wrapped with a small piece of PARAFILM to slow plant dehydration. Soybean roots were trimmed to a length of five centimeters. Ten trimmed stems were placed together in one emergence cage per location. Emergence cages consisted of 5-liter clear plastic paint-mixing buckets with lids (TCP Global Corporation, Lakeside, California, USA). A 6-cm diameter hole was cut in the side of each bucket approximately 6 cm from the bottom of the bucket. A white fine-mesh (0.02 cm mesh size, 100% polyester, Quest Outfitters, Sarasota, Florida, USA) sleeve 30 cm long was attached to the hole with hot glue to allow access to the contents of the cages. The sleeves were tied to prevent insects from escaping. In each cage, the stems were placed vertically into a 3 cm deep layer of potting soil (BM2 Seed Germination and Propagation Mix, Berger, Saint-Modeste, Quebec, CA). The emergence cages were maintained at room temperature in 16h light:8h dark, watered as needed to maintain soil moisture, and checked daily for emergence of insect adults. Adult insects were collected manually into microcentrifuge tubes, freeze-killed in -20 °C for 24 hours and preserved in 95% ethanol for taxonomic and molecular identification.

DNA extraction

Non-destructive DNA extraction from individual specimens followed a modified HotSHOT protocol (Truett et al. 2000). Each specimen was placed in a 0.2 mL PCR tube (Olympus plastic, Cat# 27-125) with 100 μL of the lysis reagent (25 mM NaOH: 0.2 mM disodium EDTA) and incubated at 95 °C for 30 minutes on a Mastercycler nexus PCR cycler (Eppendorf). Samples were cooled to 4 °C and 100 μL of neutralizing reagent (40 mM Tris-HCl) was added to each sample (a final volume of 200 μL). The aqueous solution containing DNA was moved to a fresh tube and 95% Ethanol was added to the specimen for preservation.

DNA barcoding

We barcoded all Synopeas (n=16) as well as six adult specimens of R. maxima randomly selected from the emergence cages. The cytochrome oxidase subunit I (COI) gene was amplified alongside negative controls using the universal primer pair LCO-1490/HCO-2198 for S. maximum and COIA/J-1718 for R. maxima (Folmer et al. 1994; Simon et al. 1994; Funk et al. 1995; Gagné et al. 2019). PCR reactions were prepared in a final volume of 20 μL with 1 μL of DNA template, Q5 Hot Start High-Fidelity 2X Master Mix (New England BioLabs), and 500 nM of each primer. Thermalcycling was performed on a Mastercycler nexus PCR cycler (Eppendorf) with an initial denaturation of 2 min at 98 °C, followed by 40 cycles of amplification (10 s at 98 °C, 30 s at 60 °C, and 20 s at 72 °C), and a final elongation of 2 min at 72 °C. The annealing temperature was determined using NEB Tm calculator (version 1.15.0, https://tmcalculator.neb.com/). PCR-products were separated by gel electrophoresis on a 1% agarose gel and imaged under ultraviolet light after staining with GelRed 3X in water (Biotium). PCR-products were cleaned with the Exo-CIP Rapid PCR Cleanup Kit (New England BioLabs) according to manufacturer’s instructions and Sanger sequenced in both directions at the University of Minnesota Genomics Center (Saint Paul, Minnesota, USA). Sequences were inspected for peak quality, aligned, and trimmed of priming regions in SnapGene (SnapGene5.3.2).

Synopeas maximum-specific primer design and DNA barcoding

We designed a Synopeas maximum-specific forward primer after finding that (1) not all Synopeas samples were compatible with the LCO-1490/HCO-2198 primer set, and (2) Synopeas samples that were amplified with LCO-1490/HCO-2198 primers typically generated low sequence quality when sequencing from the LCO-1490 primer (these sequences were ~50% of the amplicon length and were low quality, with average quality scores around 20, rather than a typical mean of ~40). Using the preliminary data from the reverse reads, we selected a new forward primer targeting a region that was conserved across our samples (SYN_F: 5’-CGATTAGAAGTTGGAACTCC-3’) and generated a 550-bp amplicon when combined with the HCO-2198 reverse primer. A new PCR reaction mix was prepared for all wasps (n=16) as described in the section above, using the new primer pair (SYN_F/HCO-2198). Thermal cycling was performed on a Mastercycler nexus PCR cycler (Eppendorf) with an initial denaturation of 2 min at 98 °C, followed by 35 cycles of amplification (10 s at 98 °C, 30 s at 61 °C, and 20 s at 72 °C), and a final elongation of 2 min at 72 °C.New PCR-products were separated, purified, and sequenced as described above. Resulting sequences were uploaded to BOLD and are listed in Table 1.

Table 1.

Download as

CSV

XLSX

Specimens of Synopeas maximum reared from emergence cages.

Lab code†	Collecting unit identifier	Sex	BOLD ID
PG01	FSCA 00095876	Male	SYMAX005-23
PG02	FSCA 00095877	Male	SYMAX006-23
PG03	FSCA 00095878	Male	SYMAX011-23
PG04	FSCA 00095879	Male	SYMAX010-23
PG05	FSCA 00095880	Female	SYMAX009-23
PG06	FSCA 00095881	Female	SYMAX001-23
PG07	FSCA 00095882	Male	SYMAX002-23
PG08	FSCA 00095883	Female	SYMAX003-23
PG10	FSCA 00095885	Female	SYMAX012-23
PG11	FSCA 00060750	Male	SYMAX013-23
PG12	FSCA 00060752	Female	SYMAX014-23
PG13	FSCA 00060751	Male	SYMAX004-23
PG17	FSCA 00060754	Female	SYMAX016-23
PG18	FSCA 00060755	Male	SYMAX007-23
PG20	FSCA 00060756	Male	SYMAX008-23
PG21	FSCA 00060757	Female	SYMAX017-23

Phylogenetic reconstruction

A phylogenetic analysis was performed with Synopeas sequences available on BOLD (Suppl. material 1) along with S. maximum sequences, and selected outgroups (n=2412). Amino acid sequences were aligned with MAFFT version 7.475 (Katoh and Standley 2013) using default parameters for downstream use in phylogenetic analyses. The alignment was trimmed in SnapGene (SnapGene5.3.2) to remove sequences from outside of the SYN_F/HCO-2198 amplicon. A reduced version of the alignment, which excluded identical sequences (n=1459, 518 bp in length), was submitted to maximum likelihood analysis in RAxML version 8.2.11 using a GTRGAMMA substitution model and 1000 bootstrap replicates (Stamatakis 2014). Tree topology was rooted and visualized in FigTree version 1.4.4 and annotated in Inkscape version 1.2.1. Species delimitation was performed with the ASAP web server and default settings (https://bioinfo.mnhn.fr/abi/public/asap/) including the full aligned FASTA file containing all sequences (n=2412) and a JC69 Jukes-Cantor substitution model (Puillandre et al. 2021). The best scoring partition was selected for downstream processing as per ASAP guidance.

Imaging

Photography was performed using a Macropod microphotography system (Macroscopic Solutions) using 10X and 20X Mitutoyo objective lenses, with image stacks rendered in Helicon Focus. Images of primary types were deposited in Zenodo (Table 2), and images of molecular voucher specimens were deposited in BOLD.

Table 2.

Download as

CSV

XLSX

Synopeas species in the rhanis species group.

Species	Type repository	Type locality	Images
S. basipubens Buhl, 2014	NHMD	Togo
S. bialowiezaensis Buhl, 2005	MZH	Poland
S. bouceki Buhl, 2007	NHMUK	India
S. convexum Thomson, 1859	MZLU	Sweden	https://www.flickr.com/search/?tags=MZLUTYPE02851
S. cynipsiphilum (Ashmead, 1887)	USNM	USA	https://zenodo.org/record/7662443#.Y_Up1HbMJaQ
S. decurvatum (Nees von Esenbeck, 1834)	type lost	Germany
S. dentiscutum (Szabo, 1981)	HMNH	Hungary	https://doi.org/10.5281/zenodo.7585486
S. epigeios Buhl, 2006	NHMD	Denmark
S. flavicorne (Ashmead, 1893)	USNM	USA	https://zenodo.org/record/7662438#.Y_Uoy3bMJaQ
S. gibberosum Buhl, 1997	ZMUN	Norway	https://zenodo.org/record/7591619#.Y9mCmnbMJaQ
S. hyllus (Walker, 1835)	NMINH	Ireland	https://zenodo.org/record/7591590#.Y9l_AHbMJaQ
S. kimi Choi & Buhl, 2006	NIAS	South Korea
S. maximum Awad & Talamas, 2023	FSCA	USA	https://zenodo.org/deposit/7662387
S. oleae Buhl & Viggiani, 2008	MEFS, paratypes in NHMD	Italy
S. politiventre Buhl, 2015	NHMD	Chile
S. prospectum Förster, 1861	NHMW	Switzerland	https://zenodo.org/record/7591572#.Y9l9cnbMJaQ
S. protuberatus Buhl, 2009	RMNH	Vietnam
S. rhanis (Walker, 1835)	NMINH	England	https://zenodo.org/record/7442827#.Y9l-M3bMJaQ
S. sheldrakei Buhl, 2014	MZLU	Sri Lanka
S. subtilis Buhl, 2004	HMNH	Mongolia
S. talhouki Vlug, 1976	Vlug collection, paratypes in USNM, HMNH, NHMW	Lebanon	https://zenodo.org/record/7662450#.Y_UxlnbMJaQ
S. thailandicum Buhl, 2007	OOLL	Thailand
S. triangulatum Buhl, 2014	NHMD	Australia
S. tuberosum Sundholm, 1970	MZLU	South Africa
S. veenakumariae Buhl, 2014	MZLU	Sri Lanka
S. vietnamianus Buhl, 2009	RMNH	Vietnam

Institutional abbreviations

Specimens examined during this study are deposited in the following institutions and abbreviated as follows:

CNCI Canadian National Collection of Insects, Ottawa, Canada;

FSCA Florida State Collection of Arthropods, Gainesville, Florida, USA;

HMNH Hungarian Museum of Natural History, Budapest, Hungary;

MEFS Museo Entomologico Filippo Silvestri, Portici, Italy;

MZH Finnish Museum of Natural History, Helsinki, Finland;

MZLU Biological Museum Lund University, Lund, Sweden;

NHMD Natural History Museum Denmark, Copenhagen, Denmark;

NHMO Zoological Museum, University of Oslo, Oslo, Norway;

NHMUK Natural History Museum, London, UK;

NHMW Naturhistorisches Museum Wien, Vienna, Austria;

NIAS National Institute of Agricultural Sciences, Jeonju, South Korea;

NMINH National Museum of Ireland, Dublin, Ireland;

OOLL Oberösterreichische Landesmuseum Linz, Austria;

RMNH Naturalis Biodiversity Center, Leiden, Netherlands;

UMSP University of Minnesota, St. Paul, Minnesota, USA;

USNM United States National Museum, Washington, DC, USA;

ZMUN Natural History Museum, University of Oslo, Norway.

Results

Emergence cages

We collected 2221 adults of R. maxima. Other cecidomyiids collected from the cages included two individuals of Lestodiplosis spp. Two taxa of parasitoids were collected from the cages, including 16 individuals of S. maximum and 4 individuals of Aphanogmus sp. (Ceraphronidae).

DNA Barcoding

We performed DNA barcoding on each of the 16 Synopeas adults recovered from emergence cages (Table 1). COI sequences from these specimens had greater than 98% sequence similarity to each other but no close matches on GenBank. The most similar was a specimen identified as Platygastrinae sp., (GenBank ID MG501619.1, 97% query cover, 89.6% nucleotide identity), collected in Banff National Park, Alberta, Canada. Unfortunately, there are no photographic records of this specimen and a morphological comparison was not possible. We also verified the identity of randomly selected gall midges that emerged from the cages. The specimens were morphologically identified as R. maxima, had identical nucleotide sequences to each other, and a 97.67% sequence similarity to a R. maxima specimen from Nebraska, USA (accession number LC437340.1).

Systematics of Synopeas

The generic concept of Synopeas is rather straightforward, and it can be separated from other platygastrines by the fusion of T1–T2 and S1–S2 (Jackson 1969; Awad et al. 2021). However, taxonomic structure within the genus is essentially non-existent. Given that Synopeas includes over 350 species, this presents a significant challenge for species identification (Awad et al. 2021). This situation is exacerbated by the spread of numerous species by human activities, requiring revision of the world fauna to be certain that a species was not previously described. Our ability to ascertain if S. maximum is adventive is presently limited to the use of DNA barcode libraries, and comparison to type specimens that are in the morphological vicinity of S. maximum. So far, neither of these approaches have provided a match. Our efforts are not exhaustive, nor is it currently feasible to examine type material for all described species of Synopeas. Fortunately, S. maximum has a distinctive characteristic shared by a minority of species: a deep scuto-scutellar sulcus (i.e., deep divide between the mesoscutum and mesoscutellum) (Fig. 6C, D). This eliminates the need for comparison to the vast majority of described Synopeas. Within the Nearctic region, primary type images provided by Talamas et al. (2017) yielded only two species, S. cynipsiphilum (Ashmead) and S. flavicorne (Ashmead), with the mesosomal structure found in S. maximum. Given that the soybean gall midge is an emerging pest, we also considered it possible that S. maximum represents an adventive population, derived from a distribution outside of the United States. Our literature search and examination of type specimens found other species with the deeply divided mesosoma, but not a species-level match. Because the mesosomal divide creates such a distinctive “hunchbacked” appearance, we consider that it has high value as a diagnostic character and use it to define the rhanis-group of Synopeas, named for S. rhanis (Walker, 1835), which was the earliest described species with this character. The rhanis-group includes 26 species (Table 2) that we consider to be worth comparing to S. maximum. We selected five species from this group for closer examination and comparison, based on morphological similarity and occurrence in cold temperate climates. However, all species of the rhanis-group were considered and compared to the new species using type images or published descriptions.

Synopeas cynipsiphilum (Ashmead)

Fig. 1

Leptacis cynipsiphila Ashmead, 1887: 129 (original description); Fouts 1924: 117, 128 (description, lectotype designation, keyed).

Leptacis cynipsiphilus Ashmead, 1893: 271, 274 (description, spelling error, keyed).

Leptacis cynipiphila Ashmead: Kieffer 1926: 637, 646 (description, spelling error, keyed).

Synopeas cynipsiphilum (Ashmead): Masner 1967: 303 (generic transfer); Masner and Muesebeck 1968: 99 (type information); Vlug 1995: 77 (cataloged, type information).

Synopeas flavicorne (Ashmead)

Leptacis flavicornis Ashmead, 1893: 275 (original description); Fouts 1924: 117, 128 (description, type designation, keyed).

Amblyaspis flavicornis (Ashmead): Brues 1916: 532, 533 (generic transfer, description, keyed).

Synopeas flavicorne (Ashmead): Masner and Muesebeck 1968: 99 (generic transfer, type information, emendation).

Synopeas gibberosum Buhl

Fig. 2

Synopeas gibberosus Buhl, 1997: 439 (original description); Buhl 1999: 39 (keyed); Buhl and Choi 2006: 133 (keyed).

Synopeas gibberosum Buhl: Buhl and Notton 2009: 1696 (cataloged, distribution, emendation); Buhl and O’Connor 2010: 4 (distribution).

Synopeas prospectum Förster

Fig. 3

Synopeas prospectus Förster, 1861: 41 (original description); Dalla Torre 1885: 78 (reprint of Förster (1861)); Kieffer 1926: 668, 680 (description, keyed); Vlug 1973: 179 (lectotype designation); Vlug 1995: 81 (cataloged, type information); Buhl 1997: 27 (description).

Synopeas Prospectus Förster: Kieffer 1914: 419 (description).

Synopeas (Synopeas) prospectus Förster: Kieffer 1916: 554, 555 (description, subgeneric assignment, keyed).

Synopeas rhanis (Walker)

Fig. 4

Platygaster Rhanis Walker, 1836: 225 (original description).

Platygaster Acco Platygaster Acco Walker, 1836: 229 (original description. Synonymized by Vlug (1985)); Vlug 1985: 209 (junior synonym of Synopeas rhanis (Walker)).

Synopeas Acco (Walker): Marshall 1873: 19 (generic transfer); Kieffer 1914: 427 (description).

Synopeas Rhanis (Walker): Marshall 1873: 19 (generic transfer); Kieffer 1914: 341 (description of adult and immature stages).

Synopeas rhanis (Walker): Kieffer 1906: 134 (description); Kieffer 1926: 668, 676 (description, keyed); Masner 1965: 142 (type information); Vlug and Graham 1984: 130 (lectotype designation); Vlug 1985: 209 (description of type, keyed); Vlug 1995: 81 (cataloged, type information); Buhl 1999: 39 (keyed); MacGown and Evans 2003: 6 (description); O’Connor, Nash, Notton and Fergusson 2004: 24 (catalog of Irish species); Buhl and Notton 2009: 1698 (cataloged, distribution); Buhl and O’Connor 2010: 14 (distribution).

Synopeas acco (Walker): Kieffer 1926: 669, 683 (description, keyed); Vlug and Graham 1984: 129 (lectotype designation); Vlug 1985: 209 (description of type, keyed); Vlug 1995: 75 (type information); O’Connor, Nash, Notton and Fergusson 2004: 24 (type information, catalog of Irish species).

Synopeas ?rhanis (Walker): Buhl and Choi 2006: 133, 135 (keyed, distribution in Korea).

Synopeas maximum Awad & Talamas, sp. nov.

https://zoobank.org/74487E47-EB76-4899-8A22-F91906954842

Figs 5, 6

Description

Females. Body length: 1.4–1.7 mm. Body color: black. Color of legs: coxae dark brown, otherwise yellow to dark brown. Color of mesoscutellar spine: concolorous with mesoscutellar disc.

Head. Shape of head in anterior view: ovoid. Central keel: absent; present only between toruli. Sculpture on frons: reticulate microsculpture. Epitorular sculpture: reticulate microsculpture; minute rugulae. Number of clypeal setae: 4. Length of median pair of clypeal setae: longer than lateral pair. Arrangement of clypeal setae: evenly spaced. Shape of mandible: bidentate. Distance between lateral ocellus and compound eye (OOL): approximately 1 ocellar diameter. OOL: LOL: 1:2. Lateral ocellar depression: present posterolaterally. Hyperoccipital carina: present between lateral ocelli. Hyperoccipital carina strength: fine, laterally weakened. Distance between lateral ocellus and hyperoccipital carina: approximately 1 ocellar diameter. Female antenna with 3 clavomeres, claval formula 1-1-1.

Figure 1A–E.

Synopeas cynipsiphilum, female (FSCA 00097487) A head, anterior view B head and mesosoma, dorsal view C head and mesosoma, ventrolateral view D head and mesosoma, lateral view E mesosoma and metasoma, dorsal view.

Figure 2A–E.

Synopeas gibberosum, holotype female (NHMO 0001) A head, anterior view B head and mesosoma, dorsal view C head and mesosoma, ventrolateral view D head and mesosoma, lateral view E habitus, dorsolateral view.

Figure 3A–D.

Synopeas prospectum, lectotype female (NHMW-HYM #0005306) A head, anterolateral view B head and mesosoma, dorsolateral view C habitus, lateral view D habitus, dorsal view.

Figure 4A–E.

Synopeas rhanis, female (FSCA 00034191) A head, anterior view B head and mesosoma, dorsal view C mesosoma, dorsolateral view D mesosoma, lateral view E mesosoma and metasoma, dorsal view.

Mesosoma. Epomial carina: present, complete, or nearly so. Microsculpture of lateral pronotum: present anterodorsally, absent posteroventrally. Lateral pronotal sculpture coverage: more than ¾. Setation of lateral pronotum: anteroventrally glabrous, otherwise uniformly sparse (Figs 5, 6D). Mesoscutellar spine: short and pointed. Mesoscutellar spine in lateral view: pointing posteriorly; slightly upcurved at tip. Posterior margin of propodeal carina in lateral view: rounded. Mesosomal dorsum in lateral view: slightly convex. Scuto-scutellar sulcus: deep, causing mesoscutum to be elevated relative to mesoscutellum. Notauli: unmarked or faintly indicated. Parapsidal line: very faint. Setation of mesoscutum: sparse (Fig. 6B). Mesoscutal lamella: broad and rounded. Setation of mesoscutellum: anteromedially absent, posterolaterally dense.

Figure 5.

Synopeas maximum Awad & Talamas, sp. nov., holotype, female (FSCA 00095883), lateral habitus.

Figure 6A–E.

Synopeas maximum, female A head, anterior view (FSCA 00095883) B head and mesosoma (FSCA 00095881) C mesosoma, dorsolateral view (FSCA 00095883) D mesosoma, lateral view (FSCA 00095883) E metasoma, dorsal view (FSCA 00095881).

Metasoma. Microsculpture of S2: absent; faint narrow bands in lateral portion of posterior margin. Shape of S2: medioventrally expanded. Sculpture of T2: absent. Length of T2: approximately as long as mesosoma. Sculpture of S3 to S5: reticulate. Sculpture of S6: entirely reticulate. Sculpture of T6: entirely reticulate. Shape of T6: triangular, longer than wide.

Wing. Length of setae on disc of fore wing: shorter than distance between setal bases. Density of setae on disc of fore wing: moderately dense. Arrangement of setae on disc of fore wing: uniformly setose distally, proximally glabrous with linea setosa. Fore wing marginal setae: uniformly very short.

Males. Body length: 1.1 to 1.3 mm. Identical to females except for metasoma and antenna.

Antenna. Setation: A1 and A2 with few scattered setae, A3 to A10 with long, uniformly dense setae. A2 in lateral view: slightly longer than wide, distally widened forming a “teardrop” shape. A3: round, about half the size of A2 or A4. A4: roughly cylindrical, about twice as long as wide. A5 in lateral view: about half as long as A4, proximally widened. A6 to A9: roughly ovoid, wider in lateral view than in anterior view, A6 slightly smaller than following antennomeres. A10: about twice as long as wide.

Metasoma. Microsculpture of S2: narrow band at posterior margin. Sculpture of T2: absent; narrow transverse band of microsculpture at posterior margin. Length of T2: approximately as long as mesosoma, or shorter.

Material examined

Holotype : USA • ♀; Minnesota, Luverne; 43.605889°N, 96.275111°W; 30.VI–30.VII.2021; Gloria Melotto leg.; Resseliella maxima on soybean; FSCA 00095883 (FSCA).

Paratypes : USA • 1♀1♂; same collection data as for preceding; VI–VII.2021; FSCA 00095881 to 00095882 (CNCI) • 3♀3♂; same collection data as for preceding; VI–VIII.2021; FSCA 00060750 to 00060754 (FSCA) • 1♀3♂; same collection data as for preceding; VI–VII.2021; FSCA 00095876 to 00095878, 00095885 (UMSP) • 1♀2♂; same collection data as for preceding; VIII–XII.2021; FSCA 00060755 to 00060757 (UMSP) • 1♀1♂; same collection data as for preceding; VI–VII.2021; FSCA 00095879 to 00095880 (USNM).

Etymology

The species epithet refers to the ecological association with soybean gall midge, Resseliella maxima Gagné, and soybean, Glycine max (L.) Merr.

Diagnosis

Synopeas maximum can be separated from other species in the rhanis group by the following combination of characters: scuto-scutellar sulcus deep, causing mesoscutum to be elevated relative to mesoscutellum; hyperoccipital carina present between lateral ocelli, laterally weakened; mesoscutellar spine short, pointing posteriorly, sometimes with a slight upturn at the tip, but always originating from below the dorsal apex of the mesoscutellum (separating it from S. gibberosum, S. prospectum, and S. rhanis); female S2 expanded ventromedially, with microsculpture absent or very faint; female S6 and T6 entirely sculptured, triangular, about 2 times as long as wide. The latter character is very useful for separating S. maximum from S. cynipsiphilum and S. flavicorne, in which female T6 is wider than long.

Phylogenetic reconstruction

The tree in Fig. 7 is a simplified version of the full consensus tree (Suppl. materials 3, 4) that comprises all the COI sequences derived from putative Synopeas sp. available in BOLD (excluding duplicates) wherein clades not relevant to species-level treatment of S. maximum were collapsed. While the backbone largely has lower bootstrap support (<50%), many species or putative species groups are strongly supported. ASAP analysis for species delimitation defined a total of 279 putative Synopeas species with 0.024 threshold distance (Suppl. material 5). Additionally, all S. maximum sequences were clustered into a single putative species by ASAP (Suppl. material 2).

Figure 7.

Simplified phylogenetic tree of the genus Synopeas. Maximum likelihood analyses were used to reconstruct a Synopeas phylogeny. This reduced version focuses on S. maximum (red box) and its closest relatives in clade A (gray box). Node circles are color coded to indicate bootstrap support. Specimens are named with sequence ID and taxon originated from BOLD. The full tree can be found in Suppl. material 3.

We used the phylogenetic reconstruction (Fig. 7) to infer S. maximum relationships and origin. Specimens in clade A are all from Canada (n=37) and the northern United States (n=7), suggesting that S. maximum is also native to North America. However, not all specimens in clade A belong to the rhanis group, and some representatives of the rhanis group are outside of clade A. Our analysis does not support the monophyly of the rhanis group, although the deep scuto-scutellar sulcus is still useful for identification.

Discussion

The goal of the emergence cages was to obtain R. maxima and potential parasitoids of cecidomyiid agricultural pests. In addition to R. maxima, the only other cecidiomyiids collected were two individuals identified as Lestodiplosis spp., which are known to be predaceous (Gagné and Jaschhof 2017). Although we detected emergence of two hymenopteran parasitoid species (i.e., 4 individuals of Aphanogmus sp. and 16 individuals of S. maximum) in our emergence cages, we focused our investigations only on S. maximum for several reasons. Species from the genus Aphanogmus are primarily hyperparasitoids (Polaszek and Dessart 1996; Jaramillo and Vega 2009; Hofsvang et al. 2014; Pérez-Rodríguez et al. 2019) or parasitoids of predaceous cecidomyiids (Gilkeson et al. 1993; Matsuo et al. 2016), while Synopeas is a genus that is known to parasitize only cecidomyiids (Awad et al. 2021). However, the role of Aphanogmus sp. in this system needs further investigation.

Platygastrinae are important natural enemies of cecidomyiids (Austin 1984; Kim et al. 2011; Johnson et al. 2013; Chavalle et al. 2015). The recent emergence and spread of R. maxima and several other cecidomyiids, such as Contarinia nasturtii Kieffer (Philips et al. 2017), Contarinia brassicola Rondani (Mori et al. 2019) and Chilophaga virgati Gagné (Calles Torrez et al. 2014), poses a threat to agriculture in the northern U.S. and Canada. To improve management for these and other cecidomyiids that may emerge as pests in the future, further work on the ecological relationships and taxonomy of Synopeas and other Platygastrinae is required.

Here, morphological assessments grouped S. maximum with the rhanis-group (i.e., “hunchbacked” appearance) which facilitates its identification. Although the rhanis-group did not form a monophyletic clade in molecular analyses, this feature appears to be useful for diagnostics. Additionally, we discovered that S. maximum clustered with other putative Synopeas species collected from Canada and the United States, suggesting that S. maximum may be native to North America.

Different methods for assessing parasitism of midges have been described, including rearing of field-collected hosts in the laboratory (Abram et al. 2012b), PCR-based molecular methods (Greenstone 2006; Magagnoli et al. 2022), and host dissections (Roubos and Liburd 2013). Here, we opted to rear out putative natural enemies of R. maxima because host dissections would not provide us with adults for taxonomic identification, and PCR-based molecular methods were impractical since we did not want to make assumptions about putative parasitoids that might be present. Our results suggest that S. maximum is likely a primary parasitoid of R. maxima because (1) it was reared out of emergence cages with field-collected soybean stems heavily infested with R. maxima and (2) the genus Synopeas is known to exclusively parasitize Cecidomyiidae. However, additional research is needed to confirm that S. maximum is indeed a parasitoid of R. maxima. Furthermore, the potential impact of S. maximum as a biological control agent of R. maxima is still unknown and more research needs to be performed in this area.

With the known geographic range of R. maxima expanding (McMechan et al. 2021), sustainable methods to manage this pest, such as biological control, need to be explored. As taxonomic work is foundational to the introduction, conservation, and augmentation of natural enemies (de Moraes 1987), the description of S. maximum will facilitate future research on the biological control of R. maxima.

Acknowledgements

We thank Bruce Potter (University of Minnesota) for assistance collecting field samples, Zoltán Vas (HMNH), Lars Ove Hansen (ZMUN), Rune Bygebjerg (MZLU), Dominique Zimmermann (NHMW), Joe Monks and Andy Polaszek (NHMUK) for providing access to type specimens, and István Mikó (University of New Hampshire) for identifying Aphanogmus specimens. G.M. was supported by a grant secured by R.L.K and A.R.I.L. from the Minnesota Rapid Agricultural Response Fund. J.A. was supported by the Bundesministerium für Bildung und Forschung, Berlin, Germany, project “German Barcode of Life III: Dark Taxa” (FKZ 16LI1901C).

References

Abram PK, Haye T, Mason PG, Cappuccino N, Boivin G, Kuhlmann U (2012a) Biology of Synopeas myles, a parasitoid of the swede midge, Contarinia nasturtii, in Europe. BioControl 57: 789–800. https://doi.org/10.1007/s10526-012-9459-x

Abram PK, Haye T, Mason PG, Cappuccino N, Boivin G, Kuhlmann U (2012b) Identity, distribution, and seasonal phenology of parasitoids of the swede midge, Contarinia nasturtii (Kieffer) (Diptera: Cecidomyiidae) in Europe. Biological Control 62: 197–205. https://doi.org/10.1016/j.biocontrol.2012.04.003

Ashmead W (1887) Studies of the North American Proctotrupidae, with descriptions of new species from Florida. Canadian Entomologist 19: 125–132. https://doi.org/10.4039/Ent19125-7

Ashmead W (1893) Monograph of the North American Proctotrypidae. Bulletin of the United States National Museum 45: 1–472. https://doi.org/10.5479/si.03629236.45.1

Austin AD (1984) New species of Platygastridae (Hymenoptera) from India which parasitise pests of mango, particularly Procontarinia spp. (Diptera: Cecidomyiidae). Bulletin of Entomological Research 74: 549–557. https://doi.org/10.1017/S0007485300013924

Awad J, Bremer JS, Butterill PT, Moore MR, Talamas EJ (2021) A taxonomic treatment of Synopeas Förster (Platygastridae, Platygastrinae) from the island of New Guinea. Journal of Hymenoptera Research 87: 5–65. https://doi.org/10.3897/jhr.87.65563

Buhl P (1997a) On some new or little known species of Platygastrinae (Hymenoptera, Platygastridae). Entomofauna 18: 429–467.

Buhl P (1997b) Revision of some types of Platygastrinae described by A. Förster (Hymenoptera, Platygastridae). Zeitschrift der Arbeitsgemeinschaft Österreicher Entomologen: 21–28.

Buhl P (1999) A synopsis of the Platygastridae of Fennoscandia and Denmark (Hymenoptera, Platygastroidea). Entomofauna 20: 17–52.

Buhl P (2004) Platygastridae (Hymenoptera) from Mongolia. Annales Historico-Naturales Musei Nationalis Hungarici 96: 115–152.

Buhl P (2005) New or little known Palaerctic species of Platygastrinae (Hymenoptera: Platygastridae). II. Entomologica Fennica 16: 74–82. https://doi.org/10.33338/ef.84240

Buhl P (2006) Taxonomical and distributional notes on new and known Palaearctic platygastrid species (Hymenoptera: Platygastridae). Acta Zoologica Academiae Scientiarum Hungaricae 52: 287–311.

Buhl P (2007) New or little known exotic Platygastrinae (Hymenoptera, Platygastridae). Entomofauna 28: 285–292.

Buhl P (2008) On twenty-six new or little known species of Platygastridae (Hymenoptera, Platygastroidea). Entomofauna 29: 385–416.

Buhl P (2009) New species of Platygastridae from Vietnam (Hymenoptera: Platygastroidea). Zoologische Mededelingen 83: 877–918.

Buhl P (2014a) New species and new records of Platygastrinae and Sceliotrachelinae from Togo (Hymenoptera: Platygastridae). African Invertebrates 55: 67–117. https://doi.org/10.5733/afin.055.0106

Buhl P (2014b) New species of Platygastrinae and Sceliotrachelinae (Hymenoptera, Platygastroidea: Platygastridae) from Australia. International Journal of Environmental Studies 71: 425–448. https://doi.org/10.1080/00207233.2014.939436

Buhl P (2014c) New species of Platygastrinae from Sri Lanka (Hymenoptera, Platygastrinae). Entomofauna 35: 149–168.

Buhl P (2015) Further new or little known Neotropical species of Platygastrinae (Hymenoptera: Platygastridae). International Journal of Environmental Studies 72: 316–330. https://doi.org/10.1080/00207233.2014.994275

Buhl P, Choi J-Y (2006) Taxonomic review of the family Platygastridae (Hymenoptera: Platygastroidea) from the Korean peninsula. Journal of Asia-Pacific Entomology 9: 121–137. https://doi.org/10.1016/S1226-8615(08)60283-6

Buhl P, Viggiani G (2008) Description of Synopeas oleae sp. n. (Hymenoptera: Platygastridae), parasitoid of Lasioptera berlesiana Paoli (Diptera: Cecidomyiidae), with biological notes. Bollettino det Laboratorio di Entomologia Agraria “Filippo Silvestri” Portici 62: 33–37.

Buhl P, Notton G (2009) A revised catalogue of the Platygastridae of the British Isles (Hymenoptera: Platygastroidea). Journal of Natural History 43: 1651–1703. https://doi.org/10.1080/00222930902993732

Buhl P, O’Connor J (2010) Distributional records of Irish Platygastridae with corrections and additions to the Irish list. Bulletin of the Irish Biogeographical Society 34: 3–16.

Calles Torrez V, Johnson PJ, Boe A (2014) The switchgrass gall midge (Chilophaga virgati Gagné) in the northern great plains. BioEnergy Research 7: 417–423. https://doi.org/10.1007/s12155-013-9386-4

Chavalle S, Buhl PN, Censier F, De Proft M (2015) Comparative emergence phenology of the orange wheat blossom midge, Sitodiplosis mosellana (Géhin) (Diptera: Cecidomyiidae) and its parasitoids (Hymenoptera: Pteromalidae and Platygastridae) under controlled conditions. Crop Protection 76: 114–120. https://doi.org/10.1016/j.cropro.2015.06.016

Chen H, Lahey Z, Talamas EJ, Valerio AA, Popovici OA, Musetti L, Klompen H, Polaszek A, Masner L, Austin AD, Johnson NF (2021) An integrated phylogenetic reassessment of the parasitoid superfamily Platygastroidea (Hymenoptera: Proctotrupomorpha) results in a revised familial classification. Systematic Entomology 46: 1088–1113. https://doi.org/10.1111/syen.12511

Dalla Torre C (1885) Die hymenopterologischen Arbeiten Prof. Dr. Arn. Försters. Jahresbericht der Naturforschenden gesellschaft Graubünden 28: 44–82.

Ferland C-É (2020) Distribution, abundance, and biological control potential of Synopeas myles, a parasitoid of the swede midge. Master thesis, 111 pp. https://atrium.lib.uoguelph.ca/xmlui/bitstream/handle/10214/17823/Ferland_CharlesE-tienne_202003_Msc.pdf?sequence=1&isAllowed=y [August 22, 2022]

Folmer O, Hoeh W, Lutz R, Vrijenhoek R (1994) DNA primers for amplification of mitochondrial cytochrome c oxidase subunit I from diverse metazoan invertebrates. Molecular Marine Biology and Biotechnology 3: 294–299.

Förster A (1861) Ein Tag in den hoch Alpen. J.J. Beaufort, Aachen.

Fouts R (1924) Revision of the North American wasps of the subfamily Platygasterinae. Proceedings of the United States National Museum 63: 1–145. https://doi.org/10.5479/si.00963801.63-2484.1

Funk DJ, Futuyma DJ, Ortí G, Meyer A (1995) Mitochondrial DNA sequences and multiple data sets: a phylogenetic study of phytophagous beetles (Chrysomelidae: Ophraella). Molecular Biology and Evolution. https://doi.org/10.1093/oxfordjournals.molbev.a040242

Gagné RJ, Jaschhof MA (2017) Catalog of the Cecidomyiidae (Diptera) of the World (4^th Edn.). U.S. Department of Agriculture, Systematic Entomology Laboratory, Washington, 762 pp.

Gagné RJ, Yukawa J, Elsayed AK, McMechan AJ (2019) A new pest species of Resseliella (Diptera: Cecidomyiidae) on soybean (Fabaceae) in North America, with a description of the genus. Proceedings of the Entomological Society of Washington 121: e168. https://doi.org/10.4289/0013-8797.121.2.168

Gilkeson L, McLean J, Dessart P (1993) Aphanogmus fulmeki Ashmead (Hymenoptera: Ceraphronidae), a parasitoid of Aphidoletes aphidimyza Rondani (Diptera: Cecidomyiidae). The Canadian Entomologist 125: 161–162. https://doi.org/10.4039/Ent125161-1

Greenstone MH (2006) Molecular methods for assessing insect parasitism. Bulletin of Entomological Research 96: 1–13. https://doi.org/10.1079/BER2005402

Helton ML, Tinsley NA, McMechan AJ, Hodgson EW (2022) Developing an injury severity to yield loss relationship for soybean gall midge (Diptera: Cecidomyiidae). Journal of Economic Entomology 115: 767–772. https://doi.org/10.1093/jee/toac038

Hodgson EW, Helton M (2021) Soybean gall midge efficacy. Arthropod Management Tests 46: tsab010. https://doi.org/10.1093/amt/tsab010

Hofsvang T, Godonou I, Tepa-Yotto GT, Sæthre M-G (2014) The native hyperparasitoid complex of the invasive aphid parasitoid Lysiphlebus testaceipes (Hymenoptera: Braconidae: Aphidiinae) in Benin, West Africa. International Journal of Tropical Insect Science 34: 9–13. https://doi.org/10.1017/S1742758413000428

Huang J, Miao X, Wang Q, Menzel F, Tang P, Yang D, Wu H, Vogler AP (2022) Metabarcoding reveals massive species diversity of Diptera in a subtropical ecosystem. Ecology and Evolution 12(1): e8535. https://doi.org/10.1002/ece3.8535

Jackson DL (1969) New characters for generic separation in the Synopeas-Leptacis complex. Proceedings of the Entomological Society of Washington 71: 400–404. https://doi.org/10.5281/ZENODO.24369

Jaramillo J, Vega FE (2009) Aphanogmus sp. (Hymenoptera: Ceraphronidae): a hyperparasitoid of the coffee berry borer parasitoid Prorops nasuta (Hymenoptera: Bethylidae) in Kenya. Biocontrol Science and Technology 19: 113–116. https://doi.org/10.1080/09583150802509181

Johnson PJ, Torrez VC, Buhl PN (2013) A new species of Platygaster Latreille (Hymenoptera: Platygastridae) parasitizing Chilophaga virgati Gagné (Diptera: Cecidomyiidae). Zootaxa 3630: 184–190. https://doi.org/10.11646/zootaxa.3630.1.8

Katoh K, Standley DM (2013) MAFFT Multiple sequence alignment software version 7: improvements in performance and usability. Molecular Biology and Evolution 30: 772–780. https://doi.org/10.1093/molbev/mst010

Kieffer J (1906) Description de nouveaux Hyménoptères. Annales de la Société Scientifique de Bruxelles 30: 113–178.

Kieffer J (1914) Proctotrypidae (Partie 3). Species des Hyménoptères dʼEurope et dʼAlgérie. A. Hermann et Fils, Paris.

Kieffer J (1916) Beitrag zur Kenntnis der Platygasterinae und ihrer Lebensweise. Centralblatt für Bakteriologie, Parasitenkunde und Infekstionskrankheiten 46: 547–592.

Kieffer J (1926) Hymenoptera, Proctotrupoidea, Scelionidae (48^th edn.). Walter de Gruyter, Berlin/Leipzig. https://doi.org/10.1515/9783111432939

Kim I-K, Park J-D, Shin S-C, Park I-K (2011) Prolonged embryonic stage and synchronized life-history of Platygaster robiniae (Hymenoptera: Platygastridae), a parasitoid of Obolodiplosis robiniae (Diptera: Cecidomyiidae). Biological Control 57: 24–30. https://doi.org/10.1016/j.biocontrol.2010.12.007

Macgown MW, Evans GA (2003) A new platygastrid wasp from Florida (Hymenoptera: Platygastridae). Zootaxa 320: 1–1. https://doi.org/10.11646/zootaxa.320.1.1

Magagnoli S, Tondini E, Ratti C, Burgio G, Petacchi R (2022) A new PCR based molecular method for early and precise quantification of parasitization in the emerging olive pest Dasineura oleae. Pest Management Science 78: 1842–1849. https://doi.org/10.1002/ps.6802

Marshall T (1873) A Catalogue of British Hymenoptera; Oxyura. Entomological Society of London, London.

Masner L (1965) The types of Proctotrupoidea (Hymenoptera) in the British Museum (Natural History) and in the Hope Department of Entomology, Oxford. Bulletin of the British Museum of Natural History, Entomology Supplement 1: 3–154. https://doi.org/10.5962/p.97756

Masner L, Muesebeck C (1968) The types of Proctotrupoidea (Hymenoptera) in the United States National Museum. Bulletin of the United States National Museum 270: 1–143. https://doi.org/10.5479/si.03629236.270

Matsuo K, Uechi N, Tokuda M, Maeto K, Yukawa J (2016) Host range of braconid species (Hymenoptera: Braconidae) that attack Asphondyliini (Diptera: Cecidomyiidae) in Japan: Host range of braconid species. Entomological Science 19: 3–8. https://doi.org/10.1111/ens.12167

McMechan AJ (2021) Evaluation of at-planting soil treatment Thimet against soybean gall midge. Arthropod Management Tests 46: tsab062. https://doi.org/10.1093/amt/tsab062

McMechan AJ, Hodgson EW, Varenhorst AJ, Hunt T, Wright R, Potter B (2021) Soybean gall midge (Diptera: Cecidomyiidae), a new species causing injury to soybean in the United States. Journal of Integrated Pest Management 12: 1–8. https://doi.org/10.1093/jipm/pmab001

de Moraes GJ (1987) Importance of taxonomy in biological control. International Journal of Tropical Insect Science 8: 841–844. https://doi.org/10.1017/S1742758400023031

Mori BA, Andreassen L, Heal JD, Dupuis JR, Soroka JJ, Sinclair BJ (2019) A new species of Contarinia Rondani (Diptera: Cecidomyiidae) that induces flower galls on canola (Brassicaceae) in the Canadian prairies. The Canadian Entomologist 151: 131–148. https://doi.org/10.4039/tce.2018.63

Nees von Esenbeck C (1834) 2 Hymenopterorum Ichneumonibus affinium, Monographiae, genera Europaea et species illustrantes. JG Cottae, Stuttgart. https://doi.org/10.5962/bhl.title.26555

O’Connor J, Nash R, Notton D, Fergusson N (2004) A catalogue of the Irish Platygastroidea and Proctotrupoidea (Hymenoptera). Occasional Publications of the Irish Biogeographical Society 7: 1–110.

Pérez-Rodríguez J, Krüger K, Pérez-Hedo M, Ruíz-Rivero O, Urbaneja A, Tena A (2019) Classical biological control of the African citrus psyllid Trioza erytreae, a major threat to the European citrus industry. Scientific Reports 9: e9440. https://doi.org/10.1038/s41598-019-45294-w

Philips C, Ambourn A, Christianson L (2017) First detections of swede midge (Diptera: Cecidomyiidae) in Minnesota. Journal of Entomological Science 52: 297–300. https://doi.org/10.18474/JES17-32.1

Polaszek A, Dessart P (1996) Taxonomic problems in the Aphanogmus hakonensis species complex; (Hymenoptera: Ceraphronidae) common hyperparasitoids in biocontrol programmes against lepidopterous pests in the tropics. Bulletin of Entomological Research 86: 419–422. https://doi.org/10.1017/S0007485300035008

Potter BD, Koch RL, Melotto G, Lisak S (2022) Soybean gall midge – not just for soybean anymore. https://blog-crop-news.extension.umn.edu/2022/10/soybean-gall-midge-not-just-for.html [February 12, 2023]

Puillandre N, Brouillet S, Achaz G (2021) ASAP: assemble species by automatic partitioning. Molecular Ecology Resources 21: 609–620. https://doi.org/10.1111/1755-0998.13281

Roubos CR, Liburd OE (2013) Parasitism of Dasineura oxycoccana (Diptera: Cecidomyiidae) in North Central Florida. Environmental Entomology 42: 424–429. https://doi.org/10.1603/EN12307

Simon C, Frati F, Beckenbach A, Crespi B, Liu H, Flook P (1994) Evolution, weighting, and phylogenetic utility of mitochondrial gene sequences and a compilation of conserved polymerase chain reaction primers. Annals of the Entomological Society of America 87: 651–701. https://doi.org/10.1093/aesa/87.6.651

Srivathsan A, Ang Y, Heraty JM, Hwang WS, Jusoh WFA, Kutty SN, Puniamoorthy J, Yeo D, Roslin T, Meier R (2022) Global convergence of dominance and neglect in flying insect diversity. Ecology [preprint]. https://doi.org/10.1101/2022.08.02.502512

Stamatakis A (2014) RAxML version 8: a tool for phylogenetic analysis and post-analysis of large phylogenies. Bioinformatics 30: 1312–1313. https://doi.org/10.1093/bioinformatics/btu033

Sundholm A (1970) Chapter XVI. Hymenoptera: Proctotrupoidea. South African Animal Life: Results of the Lund University Expedition in 1950–1951. Almqvist & Wiksell, Stockholm, 305–401.

Szabo J (1981) Contributions to the platygasterid wasps of the Hortobagy National Park (Hymenoptera, Platygasteridae). Natural History of the National Parks of Hungary 1: 285–297.

Talamas EJ, Thompson J, Cutler A, Fitzsimmons Schoenberger S, Cuminale A, Jung T, Johnson NF, Valerio AA, Smith AB, Haltermann V, Alvarez E, Schwantes C, Blewer C, Bodenreider C, Salzberg A, Luo P, Meislin D, Buffington ML (2017) An online photographic catalog of primary types of Platygastroidea (Hymenoptera) in the National Museum of Natural History, Smithsonian Institution. Journal of Hymenoptera Research 56: 187–224. https://doi.org/10.3897/jhr.56.10774

Thomson C (1859) Sveriges Proctotruper. Tribus VII. Platygastrini. Öfversigt af Kongliga Ventenskaps-Akademiens Förhandlingar 16: 69–87.

Truett GE, Heeger P, Mynatt RL, Truett AA, Walker JA, Warman ML (2000) Preparation of PCR-quality mouse genomic DNA with hot sodium hydroxide and tris (HotSHOT). BioTechniques 29: 52–54. https://doi.org/10.2144/00291bm09

Vétek G, Thuróczy C, Pénzes B (2006) Interrelationship between the raspberry cane midge, Resseliella theobaldi (Diptera: Cecidomyiidae) and its parasitoid, Aprostocetus epicharmus (Hymenoptera: Eulophidae). Bulletin of Entomological Research 96: 367–372. https://doi.org/10.1079/BER2006439

Vlug H (1973) The species of Platygastridae (Hymenoptera, Proctotrupoidea) in the collection of the Naturhistorisches Museum in Vienna with special reference to the types of Förster. Entomophaga 18: 175–184. https://doi.org/10.1007/BF02372027

Vlug H (1976) Synopeas talhouki n.sp. (Hym. Platygastridae), a parasite of Odinadiplosis amygdali (Anagnostopoulos) (Dipt. Cecidomyidae) with notes on its distribution and biology. Zeitschrift für Angewandte Entomologie 80: 262–266. https://doi.org/10.1111/j.1439-0418.1976.tb03323.x

Vlug H (1985) The types of Platygastridae (Hymenoptera, Scelionoidea) described by Haliday and Walker and preserved in the National Museum of Ireland and the British Museum (Natural History). 2. Keys to species, redescriptions, synonymy. Tijdschrift voor Entomologie 127: 179–224.

Vlug H (1995) Catalogue of the Platygastridae (Platygastroidea) of the world (Insecta: Hymenoptera). Hymenopterorum Catalogus 19: 1–168.

Vlug H, Graham M (1984) The types of Platygastridae (Hymenoptera, Scelionoidea) described by Haliday and Walker and preserved in the National Museum of Ireland and the British Museum (Natural History). Designation of lectotypes. Tijdschrift voor Entomologie 127: 115–135.

Walker F (1835) On the species of Platygaster, &c. The Entomological Magazine 3: 217–274.

Supplementary materials

Supplementary material 1

CSV file with BOLD bins for putative Synopeas sequences