Print
The great greenbriers gall mystery resolved? New species of Aprostocetus Westwood (Hymenoptera, Eulophidae) gall inducer and two new parasitoids (Hymenoptera, Eurytomidae) associated with Smilax L. in southern Florida, USA
expand article infoMichael W. Gates, Y. Miles Zhang, Matthew L. Buffington
‡ National Museum of Natural History, Washington DC, United States of America
Open Access

Abstract

Aprostocetus smilax Gates & Zhang, sp. nov., is described from stem and leaf galls on Smilax havanensis Jacq. in southern Florida, USA. It is the third species of Aprostocetus Westwood known to induce plant galls. Two parasitoids of A. smilax are also described: Phylloxeroxenus smilax Gates & Zhang sp. nov. and Sycophila smilax Gates & Zhang, sp. nov. We conclude that A. smilax is the true gall inducer on Smilax L., and thus the host records of Diastrophus smilacis Ashmead and its inquiline Periclistus smilacis Ashmead, both from Smilax, are erroneous.

Keywords

Chalcidoidea, Cynipidae, Diastrophus, Sycophila, Phylloxeroxenus, Periclistus

Introduction

Gall induction in Chalcidoidea was summarized by La Salle (2005) wherein he noted its occurrence in six families, representing at least 15 independent origins. Within Eulophidae he reported 11 genera across two subfamilies, Opheliminae (two genera) and Tetrastichinae (nine genera), with documented gall induction behaviors. Since then, an additional six genera have been added to the list of gall inducers (Fisher et al. 2014; Kim et al. 2004; Kim and La Salle 2008; Kim et al. 2005; Mendel et al. 2004; Rasplus et al. 2011), including serious invasive pests of Eucalyptus L'Hér worldwide. In Tetrastichinae, the Neotropical gall associates and inducers tend to have heavier sclerotization and be larger in size than other members of the subfamily (La Salle 2005).

Aprostocetus Westwood is the largest genus within the subfamily Tetrastichinae, containing >800 species distributed worldwide that are most frequently associated with insect galls induced by four insect orders and Acari as parasitoids or inquilines (Graham 1987; La Salle 1994). Gall induction is somewhat rare in Aprostocetus, with only five documented cases worldwide: (1) A. colliguayae (Philippi) in flower buds of Colliguaja Molina (Euphorbiaceae) in Chile (Martinez et al. 1992); (2) A. gallicolus Nieves-Aldrey & Askew on stems of Hedyarum boveanum Bunge ex Basiner (Fabaceae) in Spain (Nieves-Aldrey and Askew 2011); (3) Aprostocetus monacoi Viggiani described from stem galls in Melilotus indicus L. (Fabaceae) from Italy; (4) Aprostocetus sp. on leaf midribs of Corymbia citriodora (Hook.) (Myrtaceae) reported from California and Hawaii (Beardsley and Perreira 2000); and (5) Aprostocetus sp. on stems of Melilotus Mill. infested with wound tumor virus Aureogenus Black in the US (Teitelbaum and Black 1954). In this paper we describe Aprostocetus smilax, sp. nov. (Hymenoptera: Eulophidae), a gall inducer on Smilax havanensis Jacq. and the second recorded case of gall induction for the genus in North America. We also describe two parasitoids of A. smilax, Phylloxeroxenus smilax, sp. nov., and Sycophila smilax, sp. nov. (Fig. 1).

Figure 1. 

Illustration of the stem gall on Smilax havanensis induced by Aprostocetus smilax (top right), with the inset showing the internal structure and an egg. Two eurytomid parasitoids, Phylloxeroxenus smilax (middle right), and Sycophila smilax (bottom right) are included. Illustration by Taina Litwak.

Smilax L. are monocots in the family Smilaceae, with ~350 species found mostly in tropical and subtropical regions of the world (Ferrufino-Acosta 2014). A variety of gall midges (Diptera: Cecidomyiidae) and fungi in the genus Synchytrium de Bary & Woronin are known to induce galls on Smilax (Cook 1951; Uechi et al. 2012; Urso-Guimarães and Scareli-Santos 2006). The only record of a Smilax gall induced by Hymenoptera in North America is Diastrophus smilacis Ashmead (Cynipidae: Diastrophini), collected from Illinois and Florida (Ashmead 1896a). This host record is somewhat puzzling, as other members of Diastrophus Hartig exclusively induce galls on Rosaceae (Fragaria L., Rubus L., and Potentilla L.), and all other known cynipids have been recorded from dicots (Ronquist and Liljeblad 2001; Schick et al. 2003).

Methods

Dissection

Freshly collected stem and leaf galls of S. havanensis were dissected during field work in the Miami area in 2010 by MWG and MLB. A Nikon 20× Mini Field stereoscope, fine forceps, and GEM Blue Star Super Single Edge razors were used. Slices of galls were successively removed, gradually exposing individual locules. We dissected ~20 multilocular galls and notes were made about the contents of each locule in terms of its condition and occupant prior to each occupant being assigned a code and preserved in 80% ethanol. We noted six ectoparasitoid specimens. Pertinent taxon-specific notes are included in results below.

Imaging

Ethanol-preserved specimens were dehydrated through increasing concentrations of ethanol, and transferred to hexamethyldisilazane (HMDS) (Heraty and Hawks 1998) before point-mounting. MWG identified parasitoids using a Leica M205C stereomicroscope with 10X oculars and a Leica LED ring light source for point-mounted specimen observation. We took scanning electron microscope (SEM) images with a Hitachi TM3000 (Tungsten source). Body parts of disarticulated specimens were adhered to a 12.7 × 3.2 mm Leica/Cambridge aluminum SEM stub by a carbon adhesive tab (Electron Microscopy Sciences, #77825-12). Stub-mounted specimens were sputter coated with gold-palladium using a Cressington Scientific 108 Auto from multiple angles to ensure complete coverage (~20–30 nm coating). Habitus images were obtained using a Visionary Digital imaging system. The system consists of a Canon EOS 5D Mark II digital SLR camera with a 65 mm macro lens. A Dynalite MP8 power pack and lights provided illumination. Image capture software was Visionary Digital’s proprietary application with images saved as TIF with the RAW conversion occurring in Canon Digital Photo Professional software. Image stacks were montaged with Helicon Focus 6.2.2. Image editing was done in Adobe Photoshop and plate layout in Adobe Illustrator. The painting was made from pinned and live insect specimens, plant herbarium sheets and photographs. Additional structural details of the insects were obtained from SEM photographs. The final image was painted using Adobe Photoshop.

We used several species keys to determine whether our material belonged to any described species (Balduf 1932; Graham 1987) with details below under each specific treatment. Where possible, all species identifications were corroborated by comparison with authoritatively identified specimens in the Smithsonian National Museum of Natural History.

Terminologies used for surface sculptures follow Harris (1979), while the morphology follows Gibson (1997), La Salle (1994), Lotfalizadeh et al. (2007), and Gates and Pérez-Lachaud (2012). Abbreviations for museums are: ABS, Archbold Biological Station, Archbold, FL, USA; FSCA, Florida State Collection of Arthropods; USNM, United States National Museum of Natural History, Washington, D.C., USA.

Molecular protocol

Specimens were extracted, amplified, and sequenced at USDA Beltsville Agricultural Research Center (BARC) using the DNeasyTM Tissue Kit protocol (Qiagen, Valencia, CA, USA). Specimens were digested for circa three hours using 20 μL of 20 mg/mL Proteinase K at 55 °C. The DNA was resuspended with 150 μL of Qiagen elution buffer. Fragments of mtDNA COI (620 bp) were amplified using LCO1490 5’-GGTCAACAAATCATAAAGATATTGG-3’ and HCO2198 5’-TAAACTTCAGGGTGACCAAAAAATCA-3’ (Folmer et al. 1994). Amplifications for rDNA 28S (820 bp) used 28S_D1F 5’-ACCCGCTGAATTTAAGCATAT-3’ (Harry et al. 1996) and 28S_D2R 5’-TTGGTCCGTGTTTCAAGACGGG-3’ (Campbell et al. 1994). All PCRs were performed using approximately 2 μL DNA extract, 1.25 μL 10× Buffer, 1 μL dNTP, 1 μL of each primer, 1 unit of Taq DNA polymerase (TaKaRa Bio, Mountain View, CA, USA), and purified water for a final volume of 25 μL. Amplicons of COI were generated with an initial denaturation of 1 min at 95 °C, followed by 35 cycles of 95 °C for 15 s, 49 °C for 15 s and 72 °C for 45 s, and a final elongation period of 4 min at 72 °C. The thermocycler setting for 28S is similar to COI, with the exception of annealing temperature being at 55 °C. Sequencing was conducted using a ABI 3730xl DNA sequencer following manufacturer’s instructions. Contigs were assembled and edited using Sequencher version 4.5 (Gene Codes). DNA sequences were then compared with available sequences in the Barcode of Life Database (BOLD, Ratnasingham and Hebert 2007) and the Basic Local Alignment Search Tool (BLAST) for nucleotides in GenBank. All sequences are uploaded onto GenBank (see Table 1).

Table 1.

Voucher identification and associated GenBank accession numbers.

ID Voucher COI 28S
Aprostocetus smilax G0006A MT576085 MT560740
G0008A N/A MT560741
G0008B MT576086 MT560742
G0009 MT576087 MT560743
G0010A MT576088 MT560744
G0010C MT576089 MT560745
G0010E MT576090 MT560746
G0010F MT576091 MT560747
G0013A MT576092 MT560748
G0014 MT576093 MT560749
Phylloxeroxenus smilax G0015 MT576094 MT560750
Aprostocetus smilax G0016 MT576095 N/A
G0017 MT576096 MT560751
G0018 MT576097 MT560752
G0019 MT576098 MT560753
G0021 MT576099 MT560754
G0022 MT576100 MT560755
G0023 MT576101 MT560756
G0024 MT576102 MT560757
G0025 MT576103 MT560758
G0026 MT576104 MT560759
G0030 MT576105 MT560760
G0031 MT576106 MT560761
G0034 MT576107 MT560762
G0036 MT576108 MT560763
G0041 MT576109 MT560764
Tetrastichinae sp. G0042 MT576110 MT560765
Aprostocetus smilax G0043 MT576111 MT560766
G0044 MT576112 MT560767
Sycophila smilax G0045 N/A MT560768
Aprostocetus smilax G0046 MT576113 MT560769
G0047 MT576114 MT560770
G0049 MT576115 MT560771
G0050 MT576116 N/A
Phylloxeroxenus smilax G0051 N/A MT560772
Brasema sp. G0052 MT576117 MT560773
Sycophila smilax G0053 N/A MT560774
Phylloxeroxenus smilax G0054 MT576118 MT560775
Sycophila smilax G0055 N/A MT560776
G0056 MT576119 MT560777
Phylloxeroxenus smilax G0057 N/A MT560778
G0058 N/A MT560779
Aprostocetus smilax G0059 MT576120 MT560780
G0061 MT576121 MT560781
G0063 MT576122 MT560782
G0064 MT576123 MT560783
G0065 N/A MT560784
G0066 MT576124 MT560785
G0070 MT576125 MT560786
Phylloxeroxenus smilax G0071 N/A MT560787
Sycophila smilax G0076 MT576126 MT560788
G0077 MT576127 MT560789
Aprostocetus smilax G0078 MT576128 MT560790
G0080 MT576129 MT560791
Phylloxeroxenus smilax G0082 N/A MT560792

Phylogenetic analysis

COI was aligned using default MAFFT v7.45 settings (Katoh et al. 2002) and checked by eye, for 28S the Q-INS-I strategy (Katoh and Toh 2008) was implemented to account for secondary RNA structures. Each gene was analyzed separately, and concatenated using SequenceMatrix (Vaidya et al. 2011) in IQ-TREE v2.0.5 (Minh et al. 2020). Best models of evolution were determined using ModelFinder (Kalyaanamoorthy et al. 2017) implemented in IQ-TREE, with 1000 ultrafast bootstrap pseudoreplicate support (Hoang et al. 2017). The output trees were visualized in R v4.0 (R Core Team 2020) using the packages ggtree v2.2.0 (Yu et al. 2017) and treeio v1.12.0 (Wang et al. 2020).

Results

Taxonomy

Eulophidae

Aprostocetus smilax Gates & Zhang, sp. nov.

Figs 2–3, 4–11, 12–18

Diagnosis

This species keys to Aprostocetus subgenus Aprostocetus, couplet 103 in Schauff et al. (1997) and 53 in La Salle (1997). This is the most biologically diverse and speciose of the five Aprostocetus subgenera, often associated with insects inhabiting plant galls such as Diptera (Cecidomyiidae), Hymenoptera (Cynipoidea), Hemiptera (Coccoidea), Coleoptera, and eriophyid mites (La Salle 2005). Burks (1967) published a key to 13 North American species, which is dated, and a comparative diagnosis of all 58 species is beyond the scope of this paper. Nevertheless, this species keys to couplet 2 of Burks’ key, and differs from the two species with coriaceous mesoscutum (A. coelioxydis Burks and A. granulatus Ashmead) which are both metallic blue/black in coloration. Recent phylogenomic study of Eulophidae has shown Aprostocetus to be paraphyletic (Rasplus et al. 2020), and some of these subgenera might be elevated to genus level in the future.

Material examined

Holotype , female: USA • FL: Dade Co.: SE Miami, Rockdale Pineland, Ex Smilax havanensis stem gall; 19.Dec.2001, C. Rodriguez & T. Smith leg.; USNMENT01735185 (deposited at USNM). Paratypes (4♀, 7♂): Same information as holotype; USNMENT01735186, 01735187 (1♀, 1♂, USNM). FL: Dade Co.: SE Miami, Rockdale Pineland, Ex Smilax havanensis stem gall; 18.Apr. 2010; M. Gates & M. Buffington leg.; USNMENT01735188–01735196 (3♀, 6♂, USNM).

Description

Female. Body length 1.7 mm (Fig. 2).

Color. Mostly whitish-yellow, pedicel, flagellomeres, clava, axillula, and marginal vein, sides of gastral tergites brown. Fore and midlegs white (Fig. 2).

Figure 2–3. 

Aprostocetus smilax 2 female habitus 3 male habitus.

Head. Squareish in dorsal view, 1.2× as wide as long in dorsal view (Fig. 4). Lower face coriaceous, clypeus bilobed, mandible tridentate (Fig. 5). Malar sulcus present, malar space 0.7× eye height. Genal carina absent. Toruli positioned slightly below median of compound eyes, diameter of torulus equal that of the intertorular space. Frons striate, scrobal depression converging towards the clypeus with a row of setae along depression (Fig. 4). Vertex coriaceous, ratios of POL:OOL:LOL equal to 2.8:2.1:1 (Fig. 6). Ratio of scape (minus radicle):pedicel:A1:A2: F1:F2: F3:club as 72:33:3:1:53:40:35:68; pedicel conical expanding distally; funicle cylindrical; anellus two segmented, funicular segments with single row of longitudinal sensilla and one whorl of setae, shorter than its bearing segment; clava trisegmented (Fig. 8). Head posteriorly coriaceous with a ring of setae around the outer edge, smooth with two setae. Postgenal bridge ornamentation narrow. Postgenal sulci, postgenal groove, and hypostomal bridge absent (Fig. 7). Labium square-shaped.

Forewing. Three setae on submarginal vein, 7 setae on marginal vein. Ratio of marginal vein:postmarginal vein:stigmal vein as 22.5:1:6.

Mesosoma. Mesosoma coriaceous, 1.14× as long as broad (Fig. 9); notauli complete, shallow. With 2 adnotaular seta on the midlobe of mesoscutum, and two setae on the lateral lobes (Fig. 10). Scutellum with two setae on each side, submedian groove deep, complete. Lateral panel of axilla strigate, axillula coriaceous dorsally and strigate ventrally. Prepectus coriaceous. Mesopleuron coriaceous, dorsally delimited from femoral depression raised ridge. Epicnemium flat and ventral shelf not projected forward (Fig. 11). Propodeum coriaceous and divided by median carina that diverges into raised, scalloped ridges posteriorly. Spiracle within a depression. Callus with a single seta, raised and partly overhanging outer rim of conspicuous spiracle (Fig. 10).

Figure 4–11. 

Aprostocetus smilax 4 frontal view of head 5 frontal view of lower face 6 dorsal view of head 7 posterior view of head 8 female antenna 9 lateral view of mesosoma 10 dorsal view of mesosoma 11 ventral view of mesosoma.

Metasoma. Metasoma smooth, Gt1 and Gt2 dorsally glabrous (Fig. 12), subsequent tergites each with a ring of setae (Fig. 13). Cercus with 1 seta distinctly longer (>1.5×) than others (Fig. 14).

Male. 1.1 mm. Color and sculpture as described for female (Fig. 3). Antennae with setae >1.5× as long as width of segment (Fig. 15). Gt7 curves up to form genital opening (Figs 16, 17), with a pair of long and three pairs of shorter cercal setae (Fig. 18).

Figure 12–18. 

Aprostocetus smilax 12 dorsal view of female metasoma 13 ventral view of female metasoma 14 lateral view of female metasoma 15 male antenna 16 lateral view of male metasoma 17 ventral view of male metasoma 18 closeup of male genital opening.

Variation

Size ranges from 1.6–1.8 mm for females, and 1.1–1.2 mm for males. The number of setae on marginal vein ranges from 6–8.

Biology

It induces round galls on the stems of Smilax havanensis, often coalescing to form irregularly rounded, polythalamous swellings. Individual galls can also be found on the edge of leaves. Green when fresh and of a pithy structure (Figs 1, 63 inset), tissues around the emergence hole often form a black ring.

Distribution

Southern Florida, USA.

Eurytomidae

Phylloxeroxenus smilax Gates & Zhang, sp. nov.

Figs 19–20, 21–28, 29–34

Diagnosis

Phylloxeroxenus smilax can be easily distinguished from the only other known North American species, Phylloxeroxenus phylloxerae (Ashmead), which is suspected to be a parasitoid of the cecidomyiid inquiline within Phylloxera Boyer de Fonscolombe galls on hickory (Carya Nutt.) (Ashmead 1881). The lower face is strigose and the ventral half of the body is yellow in P. smilax, while in P. phylloxerae the lower face is striate and the body is completely black. There are at least 50 undescribed species in at least three species groups for the Neotropical region that exhibit a range of variation in diagnostic generic characters such as the propodeum in lateral view forming a 90° angle with mesosoma; long/short petiole and resultant effect on striate part of S1 (Fig. 30), with the striae on S1 being a reliable diagnostic though expressed to varying degrees; and lower face with/without striae (Gates, unpublished data).

Material examined

Holotype , female: USA • FL: Dade Co.: SE Miami, Rockdale Pineland, Ex Smilax havanensis stem gall; 18.Apr. 2010; M. Gates & M. Buffington leg.; USNMENT01735174 (deposited at USNM). Paratypes (5♀, 6♂): Same information as holotype; USNMENT01735175–01735178 (3♀, 1♂, USNM). FL: Dade Co.: SE Miami, Rockdale Pineland, Ex Smilax havanensis stem gall; 19.Dec.2001, C. Rodriguez & T. Smith leg.; USNMENT01735179–01735184 (2♀, 5♂, USNM). Additional material: FL: Dade Co.: Coral Gables, Deering Estate Pineland, Ex Smilax havanensis stem gall; 23.Feb.1995, G. Melika leg.; (3♀, 4♂, ABS).

Description

Female. Body length 1.88 mm (Fig. 19).

Color. Orange-yellow; antennal segments light brown; edges of ocelli, scutellum, metasoma mediodorsally with black band, eyes pinkish red (Fig. 19).

Figure 19–20. 

Phylloxeroxenus smilax 19 female habitus 20 male habitus.

Head. Rounded in dorsal view, 1.3× as wide as long in dorsal view, umbilicate with appressed setae (Fig. 21). Lower face strigose, clypeus emarginate, mandible tridentate and step-like, supraclypeal area smooth, glabrous, slightly raised, and extending to the toruli (Fig. 22). Malar sulcus present, malar space 0.7× eye height. Genal carina present. Toruli positioned slightly above lower ocular line, diameter of torulus 4.4× that of the intertorular space. Scrobal depression carinate laterally, fading apically. Vertex imbricate, ratios of POL:OOL:LOL equal to 2.5:1:1 (Fig. 23). Ratio of scape (minus radicle):pedicel:anellus: F1:F2: F3:F4:F5:club as 19:7.3:1:7:6.6:6.6:6.4:6:18; pedicel chalice-shaped; funicle fusiform; funicular segments with single row of longitudinal sensilla and one whorl of setae, as long as its bearing segment; clava bisegmented (Fig. 25). Head posteriorly lacking postgenal lamina, postgenal groove straight and not converging in their lower part, extending ventrally to lower margin of eyes. Postgenal bridge ornamentation narrow and delicate (Fig. 24). Postgenal sulci small.

Forewing. Eight submarginal setae, 3 on parastigma. Ratio of marginal vein:postmarginal vein:stigmal vein as 2:1:1.

Mesosoma. Mesosoma umbilicate, 1.45× as long as broad; notauli complete, shallow (Fig. 27); lateral surface of prepectus triangular, smooth, ventral surface of prepectus without median tooth, subventral carina diverging strongly (Fig. 26). Mesopleuron reticulate ventrally, dorsally delimited from femoral depression by fine carina. Epicnemium flat and ventral shelf not projected forward. Propodeum in lateral view forming a 90° angle with mesosoma, broadly flattened and apically arcuate, with numerous carinae forming irregular asetose cells, these bordered laterally by setose cells; cluster of setae anterolaterad nucha (Fig. 29). Metaplural-precoxal carina complete (Fig. 28).

Figure 21–28. 

Phylloxeroxenus smilax 21 frontal view of head 22 frontal view of lower face 23 dorsal view of head 24 posterior view of head 25 female antenna 26 lateral view of mesosoma 27 dorsal view of mesosoma 28 ventral view of mesosoma.

Metasoma. Metasoma smooth, Gt4–syntergum setose, Gt6 and syntergum microreticulate; petiole 0.78× as long as broad in dorsal view, with ventral anterior groove and carina (Fig. 31); gaster S-shaped in lateral view, ovipositor angled at about 30° dorsad horizontal axis (Fig. 30); Gt4 emarginate posteriorly in dorsal view.

Male. 1.51 mm. Color and sculpture as described for female (Fig. 20). Antennal with funicular segments pedicellate, each with 2 or more rows of erect setae and about 1.5× as long as width of segment. Four funicular segments and a trisegmented clava (Fig. 32). Gastral petiole in lateral view cylindrical with projecting lateral teeth and mediodorsal prong (Fig. 34), in dorsal view length about 2.5× as long as greatest width, 1.6× as long as the length to metacoxa; evenly reticulate dorsally and ventrally (Fig. 33), smooth laterally.

Figure 29–34. 

Phylloxeroxenus smilax 29 dorsal view of propodeum 30 lateral view of female metasoma 31 ventral view of female petiole 32 male antenna 33 ventral view of male metasoma 34 lateral view of male metasoma.

Variation

Size ranges from 1.76–1.91 mm for females, and 1.45–1.52 mm for males. The coloration on the body can range from almost completely yellow, to mostly black on the dorsolateral surfaces, particularly in males.

Biology

Associated with galls of Aprostocetus smilax, likely a parasitoid of the gall inducer.

Distribution

Southern Florida, USA.

Sycophila smilax Gates & Zhang, sp. nov.

Figs 35–36, 37–44, 45–53

Diagnosis

This species is recognized by its small size, pale yellow coloration and small/faint stigmal band. It keys to couplet 9 of Balduf (1932)’s key of North American Sycophila, but differs from the other mostly yellow species Sycophila mimosae Balduf by the lack of a constricted marginal vein. The Central and South American Sycophila fauna is poorly known, and no current key exists.

Material examined

Holotype , female: FL: Dade Co.: SE Miami, Rockdale Pineland, Ex Smilax havanensis stem gall; 19.Dec.2001, C. Rodriguez & T. Smith leg.; USNMENT01735197 (deposited at USNM). Paratypes (36♀, 8♂): Same information as holotype; USNMENT01735198–01735206 (6♀, 2♂, USNM). FL: Dade Co.: SE Miami, Rockdale Pineland, Ex Smilax havanensis stem gall; 18.Apr. 2010; M. Gates & M. Buffington leg.; USNMENT01735207–01735238 (27♀, 5♂, USNM). FL: Dade Co.: South Miami, Quail Roost Pineland, Em 1.VI.2000 from galls of Smilax sp.; 8.V.2000; USNMENT01735239–01735242 (3♀, 1♂, USNM). Additional material: FL: Dade Co.: Coral Gables, Deering Estate Pineland, Ex Smilax havanensis stem gall; 23.Feb.1995, G. Melika leg. (3♀, 2♂, ABS). FL: Dade Co.: Coral Gables, Ex. Diastrophus smilacis on Smilax havanensis; 8.Nov.1977, R. Schimmel leg. (1♀, 1♂, FSCA).

Description

Female. Body length 1.8 mm (Fig. 35).

Color. Mostly pale yellow; antennal segments dark yellow; edges of ocelli, scutellum, hindtibia laterally, tarsal claw, tip of ovipositor black, pterostigma dark brown, wing band light brown, eyes pinkish red (Fig. 35).

Figure 35–36. 

Sycophila smilax 35 female habitus 36 male habitus.

Head. Rounded in dorsal view, 1.22× as wide as long in dorsal view, umbilicate with appressed setae (Fig. 37). Lower face strigose, clypeus bilobate, mandible tridentate with supraclypeal area smooth, glabrous, slightly raised and extending to toruli (Fig. 38). Malar sulcus present, malar space 0.59× eye height. Genal carina absent. Toruli positioned on lower ocular line, diameter of torulus 1.2× that of the intertorular space. Interantennal projection narrow, 1.5× that of the diameter of torulus. Scrobal depression carinate laterally, slightly diverging basally. Vertex imbricate, ratios of POL:OOL:LOL equal to 2.7:1:1 (Fig. 39). Ratio of scape (minus radicle):pedicel:anellus: F1:F2: F3:F4:F5:club as 17:6.7:1:5:5:4.7:4.7:4.7:13; pedicel chalice-shaped; funicle fusiform; funicular segments with single row of longitudinal sensilla and two whorls of setae, as long as its bearing segment; clava bisegmented (Fig. 41). Head posteriorly lacking postgenal lamina, postgenal groove faint, straight and not converging in their lower part, extending ventrally to ⅘ the lower margin of eyes (Fig. 40). Postgenal sulci small.

Forewing. Dark brown band on the wing about the same width as pterostigma and does not reach uncus, faint, reaching about ½ down the wing width, 8 submarginal setae, 3 on parastigma, 1 in basal cell, surrounded by basal and costal setal lines. Pterostigma covering marginal, postmarginal, and stigmal vein.

Mesosoma. Mesosoma umbilicate, 1.52× as long as broad; notauli complete, shallow (Fig. 43); lateral surface of prepectus triangular, smooth, ventral surface of prepectus without median tooth (Fig. 42). Mesopleuron reticulate ventrally, dorsally delimited from femoral depression by fine carina. Epicnemium flat and ventral shelf not projected forward. Propodeum with median furrow bordered mediolaterally by numerous carinae forming irregular asetose cells, these bordered laterally by setose cells (Fig. 44). Metaplural-precoxal carina interrupted by rugose carinae (Fig. 45).

Figure 37–44. 

Sycophila smilax 37 frontal view of head 38 frontal view of lower face 39 dorsal view of head 40 posterior view of head 41 female antenna 42 lateral view of mesosoma 43 dorsal view of mesosoma 44 dorsal view of propodeum.

Metasoma. Metasoma smooth, ovipositor sheath microreticulate (Figs 46, 49); petiole 2.3× as long as broad in dorsal view, with ventral anterior groove, carina, and mediodorsal prong (Figs 47, 48); gaster teardrop-shaped in lateral view, ovipositor angled at about 30° dorsad horizontal axis (Fig. 49).

Male. 1.88 mm. Mediodorsal of Gt3–5 black, wing band very faint. Otherwise color and sculpture as described for female (Fig. 36). Antenna with four funicular segments (Fig. 50). Gaster cylindrical, petiole 3× as long as wide (Figs 51, 52). Gt4 emarginate posteriorly in dorsal view (Fig. 53).

Figure 45–53. 

Sycophila smilax 45 ventral view of mesosoma 46 lateral view of female metasoma 47 ventral view of female petiole 48 lateral view of female petiole 49 closeup of female ovipositor 50 male antenna 51 dorsal view of male petiole 52 ventral view of male petiole 53 lateral view of male metasoma.

Variation

Body ranges 1.7–1.8 mm for females, 1.65–1.88 mm for males. The wing band can range from very faint, mesosoma and metasoma dorsally can be yellow or with a tinge of black.

Biology

Associated with galls of Aprostocetus smilax, likely a parasitoid of the gall inducer.

Distribution

Southern Florida, USA.

Molecular analyses

A total of 55 individuals had both or at least one of the two genes sequenced. BLAST and BOLD search results confirmed the family and sometimes genus level identification, but did not return any hits at the species level. This Smilax gall contains 3 different families of chalcidoids: the majority of the gall inhabitants consisted of the suspected gall inducer Aprostocetus smilax (n = 40), and two eurytomid parasitoids Phylloxeroxenus smilax (n = 7) and Sycophila smilax (n = 6) (Fig. 63). Specimen G0042 was identified as an unknown tetrastichine eulophid that was destructively sampled, while G0052 was identified as Brasema Cameron (Eupelmidae) (Fig. 63). This Brasema specimen was never reared as an adult from this system, we noted it encircling another larva, presumably the gall inducer, and characterized by large mandibles and erect setae.

Validity of Cynipidae associated with Smilax

As the result of this study, the validity of Diastrophus smilacis (Figs 54, 55) inducing galls on Smilax was also investigated. The resulting fieldwork revealed Aprostocetus smilax is the true gall inducer in Florida, after some 400 galls never yielded any cynipids. Further, dissections of the galls from which the type specimen of D. smilacis was reared from (collected in Illinois) revealed vascular tissue patterns consistent with dicots and not monocots (Fig. 56). As no additional material of D. smilacis has been found since its original description, despite extensive searches in Illinois (Zhiwei Liu, pers. comm.) and other parts of North America (Weld 1959), we can safely conclude Smilax is not the host of Diastrophus smilacis.

Working with the type material of both D. smilacis and Periclistus smilacis Ashmead (Figs 59, 62, the putative inquilline of D. smilacis) revealed additional curiosities that require mentioning here. Ashmead (1896a) reports specimens of D. smilacis were apparently sent to C.V. Riley from Chicago, Illinois (Figs 57, 58), and that Ashmead intended to describe them but the publication of the manuscript was delayed due to C.V. Riley’s untimely death. Ergo, time passed, and in the same year (1896), in two different publications, we find the descriptions of D. smilacis (Ashmead 1896a) and P. smilacis (Ashmead 1896b). While this is not an entirely foreign set of circumstances, the specimens referred to in these two publications are quite confusing.

Figure 54–58. 

Diastrophus smilacis 54 lateral habitus of holotype 55 dorsal habitus of holotype 56 gall of holotype 57 label of holotype 58 label of other specimens collected by C.V. Riley.

Ashmead (1896a) reports 13 specimens (females) for the description of D. smilacis, but the taxon is only known from the type specimen in the USNM and there is no record of additional specimens being loaned out; one cotype of this taxon is in AMNH, for a total of two specimens. The gall with the same type specimen number as the holotype wasp in the USNM (No. 3096, Fig. 57) has the label ‘86x’ affixed to the pin, and it is mentioned in Ashmead (1896a) that a gall was collected for this species in Florida, but no wasp. Hence, Illinois is the origin of all material associated with D. smilacis. Ashmead (1896b) describes P. smilacis from 17 specimens and goes on to say the collection data for 13 specimens (same number as D. smilacis, above) is labeled ‘No. 864, reared April 28, 1871 and four numbered 1010, reared February 4, 1884, from Diastrophus smilacis’. However, the type specimen of P. smilacis (Fig. 61) in the USNM has label data consistent with the label data of D. smilacis, suggesting Ashmead (1896b) erroneously read ‘86x’ as ‘864’ and that the same gall that yielded the type specimens of D. smilacis yielded the type specimens for P. smilacis; there is no date on the ‘86x’ specimens and it is not clear how the collection date in Ashmead (1896b) was obtained. The four specimens labeled ‘1010’ cannot be located and are presumed either lost or in another, unreported museum.

Adding to this confusing picture is that it appears A. Ritchie intended to include P. smilacis in his dissertation work on Periclistus in 1984, and even went so far as to designate a lectotype for this species (Fig. 61). The series of specimens seen in Fig. 60 is the source of the specimen that Ritchie intended as the lectotype, making the total number of specimens for P. smilacis, in the USNM, 11 specimens. When we consider D. smilacis is represented by two specimens (holotype in USNM, cotype in AMNH), we have a grand total of 13 specimens. Our conclusion from all of this is that the original 13 specimens mentioned in Ashmead (1896a) for D. smilacis turned out to be a mixture of gall inducer and inquilline, and further, the host plant for this gall was mis-identified in the field as Smilax rotundifolia L. The US Forest Service Fire Effects Information System indicates S. rotundifolia and Rubus spp. co-occur in old fields throughout the range of Smilax and it is possible that the galls of Diastrophus smilacis are actually collected from a Rubus, and the two host plants were confused when the original collection was made.

The original collections made in Florida in 2010 that led to the chalcidoids described herein were also focused on the (now) erroneous records of D. smilacis on Smilax havanensis mentioned in Beutenmüller (1909) collected around Miami by Dr. E. Bessey. When looking closely at the D. smilacis gall figured in Beutenmüller (1909), it is clear that gall matches exactly what was collected in this project and illustrated in Fig. 1. No gall material from S. havanensis is in the cynipid gall collection, and indeed, there are no galls in this collection that look like the one figured in Beutenmüller (1909). As no cynipids apparently emerged from the Miami gall reported and figured in Beutenmüller (1909), we consider this an erroneous host record as well.

Lastly, the USNM has a specimen labeled as lectotype for Periclistus smilacis, yet this taxon lacks a published lectotype designation. We presume the team of Ritchie and Shorthouse, whose names appear on the purported lectotype labels, planned to publish these designations (as mentioned above), but were not able to. In order to stabilize the name of Periclistus smilacis, we hereby designate USNMENT00802336, type number 3287, as lectotype of this taxon, deposited in the USNM (Figs 59, 61, 62).

Figure 59–62. 

Periclistus smilacis 59 lateral habitus of lectotype 60 series of specimens and gall of lectotype 61 label of lectotype 62 dorsal habitus of lectotype.

Figure 63. 

Concatenated 28S and COI phylogram of the Smilax gall inhabitants estimated using Maximum Likelihood framework in IQ-TREE2. Black dots at the nodes indicate ≥90% ultrafast bootstrap support. Inset images in counterclockwise order: Stem gall induced by Aprostocetus smilax on Smilax havanensis, with emergence holes (photo by MWG); A. smilax, Sycophila smilax, Phylloxeroxenus smilax.

Conclusion

Here we describe the new eulophid species Aprostocetus smilax, the second recorded case of gall induction by Aprostocetus in North America. This new species is the true gall inducer on Smilax, and previous records of cynipid species Diastrophus smilacis and the inquiline Periclistus smilacis associated with this host plant are erroneous. Additionally, we described two eurytomid parasitoids associated with this Smilax gall. The distribution of all three new species is on the southern tip of mainland USA, but it is likely that they are also found in the Caribbean region in which the host plant S. havanensis is found (Ferrufino-Acosta 2014). A comprehensive taxonomic revision of these incredibly diverse but understudied minute wasps will undoubtedly reveal additional ecological associations and new species.

Acknowledgements

We thank the following museum collections and curators for providing loans: Archbold Biological Station (Mark Deyrup), Florida State Collection of Arthropods (Elijah Talamas). Staff at the Charles Deering Estate permitted access for collecting galls and staff from Miami-Dade County (C. Rodriguez and T. Smith) provided specimens and access to Rockdale Pineland for gall collecting. We would also like to thank Gary Oullette for performing all DNA extractions and amplifications, Taina Litwak for the illustration and image editing, and Zhiwei Liu for discussion on the validity of Diastrophus smilacis. Finally, we would like to thank Paul Hanson and an anonymous reviewer that have provided comments that improved the manuscript. Mention of trade names or commercial products in this publication is solely for the purpose of providing specific information and does not imply recommendation or endorsement by the USDA. USDA is an equal opportunity provider and employer.

References

  • Ashmead WH (1881) Studies on the North American Chalcididae with descriptions of new species from Florida. Transactions of the American Entomological Society 13: 125–135. https://doi.org/10.2307/25076470
  • Ashmead WH (1896b) Descriptions of new parasitic Hymenoptera. Transactions of the American Entomological Society 23: 179–234.
  • Beardsley J, Perreira W (2000) Aprostocetus sp. (Hymenoptera: Eulophidae: Tetrastichinae), a gall wasp new to Hawaii. Proceedings Hawaiian Entomological Society 34: 183.
  • Beutenmüller W (1909) North American species of Diastrophus and their galls. Bulletin of the American Museum of Natural History 26: 135–145.
  • Burks BD (1967) The North American species of Aprostocetus Westwood (Hymenoptera: Eulophidae). Annals of the American Entomological Society 60: 756–760. https://doi.org/10.1093/aesa/60.4.756
  • Campbell B, Steffen‐Campbell J, Werren J (1994) Phylogeny of the Nasonia species complex (Hymenoptera: Pteromalidae) inferred from an internal transcribed spacer (ITS2) and 28S rDNA sequences. Insect Molecular Biology 2: 225–237. https://doi.org/10.1111/j.1365-2583.1994.tb00142.x
  • Ferrufino-Acosta L (2014) Taxonomic revision of the genus Smilax (Smilacaceae) in Central America and the Caribbean Islands. Willdenowia 40: 227–280. https://doi.org/10.3372/wi.40.40208
  • Fisher N, Moore A, Brown B, Purcell M, Taylor GS, La Salle J (2014) Two new species of Selitrichodes (Hymenoptera: Eulophidae: Tetrastichinae) inducing galls on Casuarina (Casuarinaceae). Zootaxa 3790: 534–542. https://doi.org/10.11646/zootaxa.3790.4.2
  • Folmer O, Black M, 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.
  • Gates MW, Pérez-Lachaud G (2012) Description of Camponotophilus delvarei, gen. n. and sp. n. (Hymenoptera: Chalcidoidea: Eurytomidae), with discussion of diagnostic characters. Proceedings of the Entomological Society of Washington 114: 111–124. https://doi.org/10.4289/0013-8797.114.1.111
  • Gibson GA (1997) Morphology and Terminology. In: Gibson GA, Huber JT, Woolley JB (Eds) Annotated keys to the genera of Nearctic Chalcidoidea (Hymenoptera). NRC Research Press, Ottawa, 16–44.
  • Graham MWRdV (1987) A reclassification of the European Tetrastichinae (Hymenoptera: Eulophidae), with a revision of certain genera. Bulletin of the British Museum 55: 1–392.
  • Harris RA (1979) Glossary of surface sculpturing. Occasional Papers in Entomology 28: 1–31.
  • Harry M, Solignac M, Lachaise D (1996) Adaptive radiation in the Afrotropical region of the Paleotropical genus Lissocephala (Drosophilidae) on the pantropical genus Ficus (Moraceae). Journal of Biogeography 23: 543–552. https://doi.org/10.1111/j.1365-2699.1996.tb00016.x
  • Heraty J, Hawks D (1998) Hexamethyldisilazane: A chemical alternative for drying insects. Entomological News 109: 369–374.
  • Hoang DT, Chernomor O, von Haeseler A, Minh BQ, Le SV (2017) UFBoot2: Improving the ultrafast bootstrap approximation. Molecular Biology and Evolution: msx281. https://doi.org/10.1101/153916
  • Kalyaanamoorthy S, Minh BQ, Wong TK, von Haeseler A, Jermiin LS (2017) ModelFinder: fast model selection for accurate phylogenetic estimates. Nature Methods 14: 587–589. https://doi.org/10.1038/nmeth.4285
  • Katoh K, Misawa K, Kuma Ki, Miyata T (2002) MAFFT: a novel method for rapid multiple sequence alignment based on fast Fourier transform. Nucleic Acids Research 30: 3059–3066. https://doi.org/10.1093/nar/gkf436
  • Katoh K, Toh H (2008) Recent developments in the MAFFT multiple sequence alignment program. Briefings in Bioinformatics 9: 286–298. https://doi.org/10.1093/bib/bbn013
  • Kim I-K, Delvare G, La Salle J (2004) A new species of Quadrastichus (Hymenoptera: Eulophidae): a gall-inducing pest on Erythrina (Fabaceae). Journal of Hymenoptera Research 13: 243–249.
  • Kim I-K, La Salle J (2008) A new genus and species of Tetrastichinae (Hymenoptera: Eulophidae) inducing galls in seed capsules of Eucalyptus. Zootaxa 1745: 63–68. https://doi.org/10.11646/zootaxa.1745.1.6
  • Kim I-K, McDonald M, La Salle J (2005) Moona, a new genus of tetrastichine gall inducers (Hymenoptera: Eulophidae) on seeds of Corymbia (Myrtaceae) in Australia. Zootaxa 989: 1–10. https://doi.org/10.11646/zootaxa.989.1.1
  • La Salle J (2005) Biology of gall inducers and evolution of gall induction in Chalcidoidea (Hymenoptera: Eulophidae, Eurytomidae, Pteromalidae, Tanaostigmatidae, Torymidae). In: Raman A, Schaefer C, Withers T (Eds) Biology, ecology, and evolution of gall-inducing arthropods. Science Publishers Inc, Enfield, 507–537.
  • Lotfalizadeh H, Delvare G, Rasplus J-Y (2007) Phylogenetic analysis of Eurytominae (Chalcidoidea: Eurytomidae) based on morphological characters. Zoological Journal of the Linnean Society 151: 441–510. https://doi.org/10.1111/j.1096-3642.2007.00308.x
  • Martinez E, Montenegro G, Elgueta M (1992) Distribution and abundance of two gall-makers on the euphorbiaceous shrub Colliguaja odorifera. Revista Chilena de Historia Natural 65: 75–82.
  • Mendel Z, Protasov A, Fisher N, La Salle J (2004) Taxonomy and biology of Leptocybe invasa gen. & sp. n. (Hymenoptera: Eulophidae), an invasive gall inducer on Eucalyptus. Australian Journal of Entomology 43: 101–113. https://doi.org/10.1111/j.1440-6055.2003.00393.x
  • Minh BQ, Schmidt HA, Chernomor O, Schrempf D, Woodhams MD, Von Haeseler A, Lanfear R (2020) IQ-TREE 2: New models and efficient methods for phylogenetic inference in the genomic era. Molecular Biology and Evolution 37: 1530–1534. https://doi.org/10.1093/molbev/msaa015
  • Nieves-Aldrey JL, Askew R (2011) Two new species of Tetrastichinae (Hymenoptera: Eulophidae) from Spain, the first known native European gall inducing tetrastichine and its parasitoid. Annales de la Société entomologique de France 47: 154–161. https://doi.org/10.1080/00379271.2011.10697707
  • R Core Team (2020) R: A language and environment for statistical computing.
  • Rasplus J-Y, La Salle J, Delvare G, Mckey D, Webber BL (2011) A new Afrotropical genus and species of Tetrastichinae (Hymenoptera: Eulophidae) inducing galls on Bikinia (Fabaceae: Caesalpinioideae) and a new species of Ormyrus (Hymenoptera: Ormyridae) associated with the gall. Zootaxa 2907: 51–59.
  • Rasplus J-Y, Blaimer BB, Brady SG, Burks RA, Delvare G, Fisher N, Gates M, Gauthier NA, Gumovsky AV, Hansson C, Heraty JM, Fusu L, Nidelet S, Pereira RAS, Sauné L, Ubaidillah R, Cruaud A (2020) A first phylogenomic hypothesis for Eulophidae (Hymenoptera, Chalcidoidea). Journal of Natural History 54: 597–609. https://doi.org/10.1080/00222933.2020.1762941
  • Schauff ME, La Salle J, Coote LD (1997) Eulophidae. In: Gibson GA, Huber JT, Woolley JB (Eds) Annotated keys to the genera of Nearctic Chalcidoidea (Hymenoptera). NRC Research Press, Ottawa, 327–429.
  • Schick K, Liu Z, Goldstein P (2003) Phylogeny, historical biogeography, and macroevolution of host use among Diastrophus gall wasps (Hymenoptera: Cynipidae). Proceedings of the Entomological Society of Washington 105: 715–732.
  • Teitelbaum SS, Black L (1954) The effect of a phytophagous species of Tetrastichus, new to the United States, on sweet clover infected with wound-tumor virus. Phytopathology 44: 548–550.
  • Uechi N, Yukawa J, Usuba S, Gyoutoku N, Mitamura T (2012) Findings of new cecidomyiid galls induced by Asphondylia segregates (Diptera: Cecidomyiidae) in Japan. Esakia: 51–57.
  • Urso-Guimarães M, Scareli-Santos C (2006) Galls and gall makers in plants from the Pé-de-Gigante Cerrado reserve, Santa Rita do Passa Quatro, SP, Brazil. Brazilian Journal of Biology 66: 357–369. https://doi.org/10.1590/S1519-69842006000200018
  • Vaidya G, Lohman DJ, Meier R (2011) SequenceMatrix: concatenation software for the fast assembly of multi‐gene datasets with character set and codon information. Cladistics 27: 171–180. https://doi.org/10.1111/j.1096-0031.2010.00329.x
  • Viggiani G, Monaco R (2014) Description of a new gall-inducing species of Aprostocetus (Hymenoptera: Eulophidae) on Melilotus indicus from Southern Italy. Journal of Entomological and Acarological Research 46: 27–29. https://doi.org/10.4081/jear.2014.1782
  • Wang L-G, Lam TT-Y, Xu S, Dai Z, Zhou L, Feng T, Guo P, Dunn CW, Jones BR, Bradley T (2020) treeio: an R package for phylogenetic tree input and output with richly annotated and associated data. Molecular Biology and Evolution 37: 599–603. https://doi.org/10.1093/molbev/msz240
  • Weld LH (1959) Cynipid Galls of Eastern United States. Privately Printed, Ann Arbor, 124 pp.
  • Yu G, Smith DK, Zhu H, Guan Y, Lam TTY (2017) ggtree: an R package for visualization and annotation of phylogenetic trees with their covariates and other associated data. Methods in Ecology and Evolution 8: 28–36. https://doi.org/10.1111/2041-210X.12628