Microhoriini Bonadona, 1974

Kejval, Zbyněk & Chandler, Donald S., 2020, Generic revision of the Microhoriini with new species and synonymies from the Palaearctic Region (Coleoptera: Anthicidae), Acta Entomologica Musei Nationalis Pragae 60 (1), pp. 95-154 : 105-109

Microhoriini Bonadona, 1974

Microhorini Bonadona, 1974: 106 , 108, 110, Figs 9 –15.

Microhorini : BONADONA (1991): 12, 14 (key to genera, France); BUCCI- ARELLI (1980): 48, 175 (characters, key to genera); BOCÁK (1993): 114 (checklist); HEMP (1994): 114 (characters, relationships, phylogeny, canthariphily); DETTNER (1997): 126 (canthariphily); KUBISZ & SZWAŁKO (1998): 9, 32 (checklist); NARDI (2003): 53 (note, synonymy).

Microhoriini : DE MARZO (1996): 150, 160 (internal reproductive structures); CHANDLER (2002): 550, 555 (biology, note); NARDI (2003): 53 (note); BOUCHARD et al. (2011): 448 (checklist); ZAHRADNÍK (2017): 319 (checklist).

Liparoderini Bonadona, 1990a: 20.

Liparoderini: BONADONA (1991): 123 (characters, key to genera and species, France); BONADONA (2013): 11, 82 (ditto); NARDI (2003): 53 (synonymy).

Diagnosis. Antennal insertion exposed and clearly visible; mandibles always with two apical teeth, distinct mola, cutting edge on right mandible, and well-developed prostheca ( Figs 51, 52 ); maxillary galea rounded and simply densely setose apically; apical maxillary palpomere small and more or less elongate to subtriangular (never conspicuously triangular/securiform); labial palpi always trimerous; latero-basal pronotal foveae usually distinct (varying in distinctness); mesepisterna usually narrowly separate medially at anterior margin (except Liparoderus , Fig. 32 ); procoxal rests with extension of anterior margin laterally; intercoxal process of mesoventrite always fully developed; mesepimera usually deeply excavate and conspicuously setose in cavity and along its ventral margin (except some apterous Aulacoderus and all Falsophilus); mesepimeral foveae always present, varying in prominence (extremely developed in some Aulacoderus ); metendosternite with well-developed lamina ( Fig. 53 , simplified in apterous species only); mesoscutellum usually rounded apically, rarely bluntly pointed (apterous species only); sutural striae of elytra absent or at most indicated subapically and thus indistinct; elytral apices in males almost always modified, with channel/pores for cantharidin gland ( Figs 41–46 , except Neocrohoria and some species of Microhoria ); metacoxae with complete posterior carina (except Liparoderus ); terminal spurs of tibiae simple; abdominal sternite VIII formed by single or paired sclerites (sometimes conspicuously modified, Figs 90–92 , 99 ); abdominal segment IX (spiculum) Y-shaped; tegmen of aedeagus at least partly tubular (except monotypic Neocrohoria); endophallus of aedeagus with various sclerotized inner structures (spines, projections, longitudinal sclerite/-s), and slender free apodeme with basal cup/plate (both sometimes strongly reduced); ovipositor with well-developed, slender coxites and styli.

Remarks. BONADONA (1974) proposed Microhoriini to include those groups with a unique form of the aedeagus that he described as cuculliform (= like a cap or hood). His description of this form (here modified using the terminology of LAWRENCE et al. 2011) stated that the elongate, tubular phallobase formed a ‘sleeve’, with the fused parameres forming a triangular apical parameral plate that was angled over the phallobase like a cap, and with the endophallus typically containing some large hooks/ spines/teeth. This form of the aedeagus is indeed unique in the Anthicidae . The tegmen (the combined phallobase and parameral plate) forms a circular, ventrally closed structure, within which the endophallus moves. Slight lateral indentations usually indicate the separation point of the parameral plate and phallobase, but for numerous species this is not apparent. A distinct penis (median lobe) is absent. At the basal end of the tegmen is typically a distinct, lightly sclerotized ‘cup’ ( VAN HILLE 1984: 6, Figs 7–9 ) that can move freely within the tubular tegmen. The ‘cup’ has its basal portion usually distinctly sclerotized, with the fused baculi (struts) extending apically as an apodeme that can be minute or conspicuous and may extend through the length of the endophallus. The cup can be easily pulled out and disassociated from the aedeagus. While the function of the ‘cup’ is unknown, its apodeme possibly serves to provide rigidity to the endophallus when everted during mating. The primary gonopore is typically found in the apical half near the apex of the endophallus, and is usually visible as a broad, lightly sclerotized ring.

DE MARZO (1996) produced the only paper dealing explicitly with the internal female and male reproductive structures of Anthicidae . He covered 23 European species that were members of the Notoxinae and Anthicinae . Two species of Microhoriini were examined ( Clavicomus paganettii and Microhora raveli), one species of Formicomini , 17 species of Anthicini from 8 genera including Endomia (all preceding are Anthicinae ), and three species of Notoxus ( Notoxinae – N. monoceros species-group). He found that males of Microhoriini and Notoxinae species lack an ejaculatory ampulla (present in all other groups examined), and that for the females only the Microhoriini had a large thick-walled bursa copulatrix ( Fig. 2 ) adjacent to the spermatheca in which he found 1–6 spermatophores in the Microhoria species and one in the single Clavicomus species examined. The members of Notoxus and other genera had a flexible expandable spermathecal receptacle situated at some distance from the vagina, with that of the Notoxinae comparably quite large. These genera lacked spermatophores, and the sperm were deposited freely mixed with secretions of the male accessory glands ( DE MARZO 1996). Analogous similarities of Notoxinae and Microhoriini are that both have large sperm storage receptacles and that males could potentially share a large amount of cantharidin-laden secretions with females when mating, potentially much more than would be shared with non-canthariphilous species. In consideration of the female large seminal fluid storage organs and lack of a male ejaculatory ampulla ( DE MARZO 1996), the very large primary gonopore suggests the ability to move large amounts of seminal fluid under a lower pressure, one of the issues necessary to address in transfer of cantharidin-laden fluids. Members of the European Anthicini genera, including Endomia, typically have a long, narrow ejaculatory duct coupled to a muscular ejaculatory ampulla that can force the free, non-motile sperm quickly through their narrow conduit ( DE MARZO 1996).

The unique form of the male genitalia, with its basal cup, lack of a penis, and tubular form, along with the presence of modified male elytra associated with canthariphily provide strong support for recogniting Microhoriini as a monophyletic group, and the shared form of the mesothoracic sclerites ( Figs 11–30 ) provides similarly strong support that Microhoria sensu novo is a monophyletic group. DE MARZO (1996) examined 17 species from eight European genera of Anthicini and two species of Microhoria placed in different species-groups, and provided additional, if limited, evidence for the monophyly of the Microhoriini in noting that the female internal genitalia of Microhoria are different from those of Anthicini by possession of a large bursa copulatrix, presumably enlarged to receive poten- tially large amounts of cantharidin-rich spermatic fluids. This coupled with the quite large primary gonopore can allow the freer and quicker passage of male fluids during mating (the primary gonopore being comparatively very small in the Anthicini studied).

Biology and collecting information. Larvae of Microhoriini are unknown. Limited information on biology of the adults at best generally indicates where or how they were collected. In northern India adults of Microhoria have been most commonly taken by beating the vegetation of trees and shrubs (Z. Kejval, pers. observ.). Others have stated that beating/sweeping of oak, hazel, alder, and cherry shrubs is productive, particularly when they are flowering, as well as by sweeping trees and herbaceous plants in grasslands and meadows ( BUCCIARELLI 1980, TEZCAN et al. 2002, BONADONA 2013, KEJVAL 2017). TAKADA et al. (2006) found Microhoria fugiens (as Clavicollis) to be commonly taken by sweeping the shrub layer in a Japanese forest. NARDI & MIFSUD (2003) reported the collection of Microhoria velox velox (as Tenuicomus) on Foeniculum vulgare (Apiaceae) in Malta, while this species was common on low whitish walls separating crops. Also Aulacoderus sulcithorax melitensis (Pic, 1903) was taken mainly by sweeping flowers, and in 2002 was collected primarily on mature shrubs of Lonicera implexa (Caprifoliaceae) . Microhoria species have also been taken by hand-collecting from the ground where they crawl in or beneath plant debris, or very occasionally have been found beneath stones or on sand in riparian areas, though this is not a typical habitat for members of the genus.

Baits are especially effective for collecting many species of this tribe. The odor of cantharidin is quite attractive to many species of Microhoriini . The odor can be produced from the cantharidin present in living or dead meloid beetles, or may be generated by using ‘cantharidin traps’, Petri dishes with filter paper impregnated with a solution containing cantharidin dissolved in acetone and then dried to recrystallize the compound in the paper, or by following a similar process with cantharidin infused ethanol from vials that held meloids ( VAN HILLE 1954, ABDULLAH 1965a, CHANDLER 1976, YOUNG 1984b, KEJVAL 2017). Other attractants for some Microhoria species are rancid lard or fats from sausage or ham ( BUCCIARELLI 1980), while meat, banana, or feces may attract many species of Aulacoderus ( VAN HILLE 1985).

AUDISIO & TAGLIANTI (2010) listed three taxa of Microhoria (as Tenuicollis) as occurring in the marine littoral zone of Italy, and COLOMBINI et al. (1991) indicated that Microhoria dejeani was present in the littoral zone from the vegetated area of the foredunes to the area of the back dunes with its mesophytic vegetation and ground cover. Non-overlapping succession over the course of a year was documented for six species of Aulacoderus at one site in Botswana by FORCHHAMMER (1986).

ELMALI (1997) reported that Microhoria unicolor was an effective predator of the aphid Diuraphis noxia (Kurdjumov, 1913) in Turkish wheat fields, though since there are numerous externally similar species occurring in Turkey and no identification keys, the accuracy of the species identification is open to question. Activity of Anthicidae as micropredators of small arthropods in crops has been noted by others ( CHANDLER 2010, with references).

Distribution. Microhoriini is almost exclusively an Old World group, with the exception of the monotypic Neo- crohoria from Chile. Biogeographically the two genera ( Aulacoderus and Neocrohoria) with straight lateral mesoventral margins have a Gondwanan distribution, while the Palaearctic lineage is essentially Laurasian, holding Liparoderus and Microhoria . The tribe is most speciose in subtropical areas. Many Aulacoderus species are present in southern Africa, but no records are known from Madagascar. In the Oriental Region, they appear to be restricted to higher elevations of the Asian mainland, with southernmost records originating from the northern provinces of Laos, Thailand, and Vietnam, and southeastern Myanmar (about same latitude as northern Thailand). Microhoriini species are known from Japan and Taiwan, but they are absent from the islands of the Philippines and Indonesia. Similarly, many species occur in the Himalaya, but no reliable records are known from the Indian subcontinent. The records of Microhoria and Aulacoderus from Australia by UHMANN (2000, 2007) actually belong to the anthicine genera Sahulanthicus Telnov, 2018 and Sapintus Casey, 1895 ( KEJVAL 2017, TELNOV 2018b).

Figures

Figs 7–14. Mesothorax in ventral (left) and lateral (right) view: 7, 8 – Neocrohoria melanura (Fairmaire & Germain, 1863), Chile, Malleco prov., 4 km W Victoria (DCDC); 9, 10 – Aulacoderus mutatus (Gemminger, 1870); 11, 12 – Liparoderus venator (Dufour, 1849); 13, 14 – Microhoria oedipus (Chevrolat, 1860). Scale bars: 100 μm (Figs 7–10); 200 μm (Figs 11–14).

Figs 47–53. 47, 48 – Neocrohoria melanura (Fairmaire & Germain, 1863), O’Higgins prov., Cuesta Chada (ZKDC): 47 – aedeagus in lateral (left) and ventral (right) view; 48 – fused baculi of endophallus. 49, 50 – Prothorax in ventro-cranial view: 49 – Aulacoderus mutatus (Gemminger, 1870); 50 – Liparoderus venator (Dufour, 1849). 51, 52 – Right mandible: 51 – Microhoria oedipus (Chevrolat, 1860); 52 – Liparoderus insignis (Lucas, 1843). 53 – M. oedipus, metendosternite. Scale bars: 0.2 mm – A (Figs 51, 52); 0.5 mm – B (Fig. 50), C (Fig. 53), D (Figs 47, 48, 49).

Figs 31–38. 31 – Neocrohoria melanura (Fairmaire & Germain, 1863), Chile, Malleco prov., 4 km W Victoria (DCDC), setae of metepisterna.32 – Liparoderus insignis (Lucas, 1843), antero-median margin of mesothorax. 33–36 – Mesothorax in lateral view (detail): 33 – Microhoria oedipus (Chevrolat, 1860); 34 – M. longiceps (LaFerté-Sénectère, 1849) comb. nov.; 35 – M. caeruleicolor (Pic, 1906) comb. nov.; 36 – M. gigas (Pic, 1899). comb. nov. 37, 38 – Falsophilus minutus (Pic, 1894), after KEJVAL (2015): 37 – prothorax in ventro-lateral view; 38 – intercoxal process of abdomen. Scale bars: 100 μm.

Figs 39–46. 39, 40 – Metacoxa: 39 – Liparoderus venator (Dufour, 1849); 40 – Microhoria oedipus (Chevrolat, 1860). 41–46 – Apex of male elytron: 41 – Microhoria fasciata (Chevrolat, 1834); 42 – M. paupercula (LaFerté-Sénectère, 1849) comb. nov.; 43 – M. oedipus (Chevrolat, 1860); 44 – M. terminata (W. L. E. Schmidt, 1842); 45 – M. caeruleicolor (Pic, 1906) comb. nov; 46 – M. heydeni (Marseul, 1879) comb. nov. Scale bars: 20 μm (Figs 41, 44–46); 50 μm (Figs 42, 43), 200 μm (Figs 39, 40).

Figs 88–92. 88 – Microhoria cervi sp. nov., aedeagus in ventral view; 89 – M. sulaimanica sp. nov., aedeagus in ventral view. 90–92 – male sternum VIII: 90 – M. sawda sp. nov.; 91 – M. bacillisternum sp. nov.; 92 – M. cervi sp. nov. Scale bars: 0.2 mm – A (Fig. 92), B (Figs 89–91), C (Fig. 88).

Figs 96–99. 96–98 – Aedeagus: 96 – M. depressa (LaFerté-Sénectère, 1849), Azerbaijan, Zarat (ZKDC), lateral view; 97 – M. edmondi (Pic, 1893), syntype, lateral view; 98 – M. fergana sp. nov., lateral view. 99 – M. fergana sp. nov., male abdominal segment VIII in ventral view. Scale bars:A – (Fig. 99), B (Figs 96, 97), C (Fig. 98).

Figs 1–6. 1 – Liparoderus venator (Dufour, 1849), Spain, Zaragoza (D. Molina det., photo by B. Campo, www.biodiversidadvirtual.org.). 2 – Microhoria raveli (Pic, 1899), female internal reproductive structures, after DE MARZO (1996). 3–6 – Aedeagus in ventral view: 3 – Sapintus oceanicus (LaFerté- Sénectère, 1849), India, Tamil Nadu, Poovar (ZKDC); 4 – Anthelephila cyanea (Hope, 1833), South Africa, Western Cape prov., Greyton (ZKDC); 5 – Microhoria garavuti sp. nov., holotype; 6 – M. cervi sp. nov., paratype, Al Mughsayl (a = parameral plate, b = phallobase, c = baculi of endophallus, d = primary gonopore). Scale not specified.