31 fossil genera
209 fossil species
|See Phylogeny of Formicidae for details.|
Ward et al. (2016) - The ant subfamily Formicinae is a large and successful group, comprising about 3030 described species, distributed globally across a wide range of terrestrial environments (Brown 2000; Bolton 2003; AntCat 2015). The subfamily includes such well-known taxa as wood ants and their relatives (Formica), carpenter ants (Camponotus), weaver ants (Oecophylla), and honeypot ants (Myrmecocystus), and a diverse array of about fifty other genera. The females (workers and gynes) of this subfamily are readily distinguished from all other ants by the presence of an acidopore, a nozzle-shaped structure at the apex of the seventh abdominal sternum used to spray formic acid (Bolton 1994). Formicine workers have a flexible promesonotal suture (secondarily immobile in a few taxa), closed metacoxal cavities, single petiolar node, complete tergosternal fusion of the petiole (second abdominal segment), and no functional sting; abdominal segments 4–6 are very large relative to the sternites, which they overlap laterally and usually also ventrally (Bolton 2003).
For relationships among these genera see Phylogeny of Formicinae.
The mesosoma is attached to the gaster by a single distinct segment, the petiole. The gaster is smooth, without constrictions between the segments. The sting is absent and the tip of the gaster has a small circular opening (an acidopore) which is often surrounded by a ring of short hairs. Species of Formicinae are most often confused with species of the subfamily Dolichoderinae because both have a single segmented petiole, lack a sting and are often similar in overall body size and shape. This is especially true for the smaller species such as those in Plagiolepis. However, formicines can always be separated from dolichoderines because the tip of the gaster has a small circular opening while all dolichoderines have a slit-like opening.
Males: Boudinot (2015) - The Formicinae are uniquely identified by the following combination of characters: mandibles never serrate; antennal toruli usually situated posterad posterior clypeal margin; antenna 8–13-merous; oblique mesopleural sulcus present; at most six closed cells present on forewing; jugal lobe absent; petiolar peduncle short to absent; petiole narrowly attached to abdominal segment III; abdominal segment III unpetiolated; abdominal segment IV without cinctus between pre- and postsclerites; abdominal sternum IX unpronged and edentate.
|See images of genera within this subfamily|
Keys including this Subfamily
- Key to Australian Ant Subfamilies
- Key to Iberian Peninsula Subfamilies
- Key to Subfamilies, Males
- Key to Subfamilies of North America
- Key to subfamilies of the Neotropical region
Keys to Genus in this Subfamily
- Key to Prenolepis Group Genera (Formicinae)
- Key to Australian Genera of Formicinae
- Key to North American Genera of Formicinae
- Key to Formicinae genera of the southwestern Australian Botanical Province
- Formicinae of the southwestern Australian Botanical Province
- Myrmicinae of the southwestern Australian Botanical Province
- Key to Philippine Formicinae
Distribution and Species Richness based on AntMaps
|Tribes||Valid Genera||% World Genera||Invalid Genera||Valid Species/Subsp.||% World Species||Invalid Species/Subsp.|
|Fossil Genera||% World Fossil Genera||Valid Fossil Species/Subsp.||% World Fossil Species/Subsp.|
Fossils known from: Aix-en-Provence, France (Late Oligocene), Arkansas amber, Malvern, Arkansas, United States (Lutetian, Middle Eocene), Auxillac, Auvergne, France (Early Oligocene), Baltic amber (Bartonian, Middle to Late Eocene), Barstow Formation, California, United States (Burdigalian to Langhian, Early to Middle Miocene), Bembridge Marls, Isle of Wight, UK (Priabonian, Late Eocene), Berezovsky massif, Zakarpatskaya, Ukraine (Serravallian, Miocene), Bitterfeld amber (Bartonian, Middle to Late Eocene), Bol’shaya Svetlovodnaya, Sikhote-Alin, Russia (Priabonian, Late Eocene), Bournemouth, Dorset, U.K. (Bartonian, Middle Eocene), Brunn-Vösendorf, Austria (Late Miocene), Brunstatt, Haut-Rhin, France (Early Oligocene), Decín, Czech Republic (Early Miocene), Dominican amber, Dominican Republic (Burdigalian, Early Miocene), Eckfeld, Germany (Lutetian, Middle Eocene), Elko, Nevada, United States (Miocene), Florissant, Colorado, United States (Late Eocene), Fonseca Formation, Fonseca, Minas Gerais State, Brazil (Oligocene), Foulden Maar diatomite, New Zealand (Aquitanian, Early Miocene), Fushun amber, Liaoning, China (Ypresian, Early Eocene), Green River Formation, Colorado, United States (Lutetian, Middle Eocene), Joursac, Auvergne, France (Late Miocene), Kishenehn Formation shale, Montana, United States (Lutetian, Middle Eocene), Kleinkems, Germany (Early Oligocene), Klondike Formation, Republic, Washington, United States (Lutetian, Middle Eocene), Lac Chambon, Auvergne, France (Pliocene), Mfwangano Island, Lake Victoria, Kenya (Early Miocene), Malyi Kamyshlak, Kerch, Crimea, Russian Federation (Middle Miocene), Messel, Germany (Lutetian, Middle Eocene), Mokrina (Krottensee), Czech Republic (Late Burdigalian, Early Miocene), Montagne d'Andance, Saint-Bauzile, Ardèche, France (Early Turolian, Late Miocene), Oeningen, Switzerland (Messinian, Late Miocene), Oise amber, France (Ypresian, Early Eocene), Ormety, Georgia (Serravallian, Miocene), Parschlug, Austria (Serravallian, Miocene), Quesnel, British Columbia, Canada (Early Miocene?), Radoboj, Croatia (Burdigalian, Early Miocene), Raritan (New Jersey) amber, New Jersey, United States (Turonian, Late Cretaceous), Rott, Westphalia, Germany (Late Oligocene), Rovno amber (Priabonian, Late Eocene), Sakhalin amber, Ukraine (Thanetian, Paleocene), Schossnitz (= Sosnica?), Silesia, Poland (Late Miocene), Shanwang, China (Early Miocene), Sicilian amber, Italy (Late/Upper Miocene), Varvara Formation, Apomarma, Messara Basin, Crete, Greece (Messinian, Late Miocene), Vishnevaya Balka Creek, Stavropol, Russian Federation (Middle Miocene), Mossy Creek, near Wellborn, Texas, United States (Late Eocene), Willershausen, Lower Saxony, Germany (Late Pliocene).
List of Tribes and Genera
Boudinot (2015) - In terms of both number of described species (~3,000) and genera (51), the Formicinae is one of the most diverse lineages of ants. Genera of the Formicinae are relatively easily delimitable based on males (B. Boudinot, in prep.), but little work has been done to render males identifiable. Males are unknown or at least undescribed for seven genera (Agraulomyrmex, Alloformica, Bregmatomyrma, Forelophilus, Pseudonotoncus, Santschiella, Teratomyrmex), and the identity of Echinopla and Phasmomyrmex is uncertain.
These are some of the most common ants in Australia and can be found everywhere, often in large numbers. Many seem to feed principally upon nectar and other plant exudates, directly or indirectly via Homoptera, while others are general scavengers, foraging on the ground or on vegetation. They can be found at all times of the day and night. Nests are usually fairly large, with hundreds or thousands of workers, and range from small and cryptic to large and obvious. They are generally in soil but some species are associated with rotten wood while a few are arboreal (nesting in hollows in tree trunks or branches). Workers are generally active and fast moving and many will defend their nests vigorously, attacking intruders with their large mandibles and formic acid sprays.
Species of formicines are found world wide and are second only to the Myrmicinae in numbers of species, with over 3700 described species and subspecies and 49 genera. In Australia there are just over 400 described species and subspecies in 21 genera, with many species yet to be described. Of the 19 genera, six are found only in Australia.
LeBrun et al. (2015) - In studies with Solenopsis species and eight Formicinae species it was found that formic acid serves as a useful antioxidant of fire ant venom. Their results suggest this ability is widespread within the subfamily, at least for those ants that express the specific observed detoxification behavior of acidopore grooming (or other formic acid, self-application behavior) after alkaloid venom exposure. An increased survivorship of ants exposed to venom was found for those species that exhibited acidopore grooming. This behavior was first found in and described for Nylanderia fulva: Solenopsis invicta is one of twenty exclusively New World fire ants, a subgroup of species within the large genus Solenopsis characterized by large, aggressive colonies of polymorphic workers with piperidine alkaloid-based venom (Blum 1992). Nylanderia fulva detoxifies S. invicta venom by applying its own venom, formic acid, to body parts exposed to S. invicta venom, using a prescribed series of stereotyped actions, or acidopore grooming. Standing on its hind and middle legs, the worker curls its gaster underneath its body. It then touches its acidopore (specialized exocrine-gland duct located at the gaster tip, uniquely shared among formicines) to its mandibles, runs its front legs through its mandibles, and grooms itself vigorously by rubbing its legs over its body, periodically reapplying its acidopore to its mandibles. Acidopore grooming by N. fulva results in greatly increased survivorship following conflict with fire ants or artificial exposure to fire ant venom (LeBrun et al. 2014). Although it is not yet demonstrated how formic acid alters the bioactivity of fire ant venom, formic acid protonates the nitrogen in fire ant venom alkaloids forming a protic ionic liquid with distinct physical properties (Chen et al. 2014). Among changes to the venom including the denaturation of associated venom enzymes and an increase in viscosity, the formate salt of the alkaloid is more polar and less lipophilic, which may reduce the ability of the protonated alkaloid to penetrate the waxy cuticle or cell membranes (Meinwald 2014). In addition to fire ant venom detoxification, the toxicity of formic acid itself is reduced as a result of salt formation. Acidopore grooming provides an effective mechanism for detoxification because, in conflicts with other ants, fire ants primarily apply venom topically by gaster flagging (a venom dispersal behavior) and smearing or flicking venom droplets exuded on the tips of their stingers onto competitors (Obin and Vandermeer 1985).
The specific chemistry of the reaction of formic acid with venom alkaloids and its use when challenged with specific venom types indicates that alkaloid venoms are targets of detoxification grooming. Solenopsis thief ants, and Monomorium species stand out as brood-predators of formicine ants that produce piperidine, pyrrolidine, and pyrroline venom, providing an important ecological context for the use of detoxification behavior. Detoxification behavior also represents a mechanism that can influence the order of assemblage dominance hierarchies surrounding food competition. Thus, this behavior likely influences ant-assemblages through a variety of ecological pathways.
Known Haploid Counts: 6, 8, 9, 10, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 25, 26, 27, 28.
Haploid Count Details: 10 (Taxon: Stigmacros), 10 (Taxon: Camponotus), 10 (Taxon: Camponotus crassus), 10 • 20 (Taxon: Camponotus compressus), 12 (Taxon: Echinopla), 12 (Taxon: Oecophylla longinoda), 13 (Taxon: Camponotus), 13 • 14 (Taxon: Camponotus japonicus), 14 (Taxon: Camponotus ligniperda), 14 (Taxon: Lasius fuliginosus), 14 (Taxon: Polyrhachis illaudata), 14 (Taxon: Camponotus), 14 (Taxon: Pseudolasius), 15 (Taxon: Nylanderia), 15 (Taxon: Acropyga), 15 (Taxon: Lasius sakagamii), 15 (Taxon: Lasius alienus), 15 (Taxon: Lasius brunneus), 15 (Taxon: Lasius flavus), 15 (Taxon: Lasius nearcticus), 15 (Taxon: Lasius niger), 15 (Taxon: Lasius umbratus), 15 (Taxon: Nylanderia parvula), 15 (Taxon: Pseudolasius), 16 (Taxon: Prenolepis jerdoni), 16 (Taxon: Paratrechina longicornis), 17 (Taxon: Anoplolepis gracilipes), 17 (Taxon: Pseudolasius), 17 (Taxon: Camponotus foreli), 18 (Taxon: Camponotus rufoglaucus), 18 (Taxon: Polyrhachis illaudata), 18 (Taxon: Camponotus), 18 • 20 (Taxon: Camponotus cruentatus), 19 (Taxon: Camponotus festinus), 19 (Taxon: Camponotus), 19 (Taxon: Camponotus), 19 (Taxon: Pseudolasius), 20 (Taxon: Camponotus atriceps), 20 (Taxon: Camponotus sylvaticus), 20 (Taxon: Prenolepis jerdoni), 20 (Taxon: Camponotus balzani), 20 (Taxon: Polyrhachis hippomanes), 20 (Taxon: Camponotus), 20 (Taxon: Camponotus), 20 (Taxon: Camponotus), 20 (Taxon: Camponotus rufipes), 21 (Taxon: Polyrhachis ammon), 21 (Taxon: Polyrhachis), 21 (Taxon: Polyrhachis dives), 21 (Taxon: Polyrhachis rastellata), 21 (Taxon: Polyrhachis hector), 22 (Taxon: Notoncus ectatommoides), 23 (Taxon: Camponotus consobrinus), 23 (Taxon: Camponotus), 23 (Taxon: Camponotus), 25 (Taxon: Prenolepis jerdoni), 26 (Taxon: Formica yessensis), 26 (Taxon: Formica aquilonia), 26 (Taxon: Formica exsecta), 26 (Taxon: Formica lugubris), 26 (Taxon: Formica dakotensis), 26 (Taxon: Formica obscuripes), 26 (Taxon: Formica polyctena), 26 (Taxon: Formica pratensis), 26 (Taxon: Formica pressilabris), 26 (Taxon: Formica rufa), 26 (Taxon: Formica sanguinea), 26 (Taxon: Formica transkaucasica), 26 (Taxon: Formica truncorum), 26 (Taxon: Formica ulkei), 26 (Taxon: Formica uralensis), 26 (Taxon: Cataglyphis iberica), 26 (Taxon: Formica pergandei), 26 (Taxon: Formica subintegra), 26 (Taxon: Cataglyphis bicolor), 26 (Taxon: Camponotus), 26 (Taxon: Iberoformica subrufa), 26 (Taxon: Formica), 26 (Taxon: Formica reflexa), 26 (Taxon: Formica picea), 27 (Taxon: Formica gerardi), 27 (Taxon: Formica cinerea), 27 (Taxon: Formica cunicularia), 27 (Taxon: Formica fusca), 27 (Taxon: Formica gagates), 27 (Taxon: Formica rufibarbis), 27 (Taxon: Formica montana), 27 (Taxon: Prenolepis jerdoni), 27 (Taxon: Formica japonica), 27 (Taxon: Formica), 27 (Taxon: Polyergus samurai), 28 (Taxon: Formica truncorum), 6 (Taxon: Formica candida), 8 (Taxon: Nylanderia), 8 (Taxon: Oecophylla smaragdina), 8 (Taxon: Paratrechina longicornis), 8 (Taxon: Pseudolasius), 9 (Taxon: Prolasius), 9 (Taxon: Camponotus vitiosus), 9 (Taxon: Plagiolepis schmitzii), 9 (Taxon: Plagiolepis schmitzii), 9 (Taxon: Plagiolepis pygmaea), 9 (Taxon: Plagiolepis pygmaea), 9 (Taxon: Camponotus), 9 (Taxon: Camponotus).
Known Diploid Counts: 16, 18, 20, 24, 26, 28, 29, 30, 31, 32, 34, 35, 36, 38, 39, 40, 42, 44, 46, 48, 50, 52, 54, 78.
Diploid Count Details: 16 (Taxon: Nylanderia), 16 (Taxon: Nylanderia), 16 (Taxon: Nylanderia), 16 (Taxon: Prenolepis imparis), 16 (Taxon: Oecophylla smaragdina), 16 (Taxon: Paratrechina longicornis), 16 (Taxon: Paratrechina longicornis), 18 (Taxon: Prolasius), 18 (Taxon: Prolasius), 18 (Taxon: Plagiolepis), 18 (Taxon: Plagiolepis), 18 (Taxon: Camponotus vitiosus), 18 (Taxon: Plagiolepis schmitzii), 18 (Taxon: Plagiolepis schmitzii), 18 (Taxon: Plagiolepis pygmaea), 18 (Taxon: Plagiolepis pygmaea), 18 (Taxon: Lepisiota capensis), 18 (Taxon: Lepisiota), 18 (Taxon: Lepisiota), 18 (Taxon: Camponotus), 18 (Taxon: Brachymyrmex), 18 (Taxon: Camponotus), 18 (Taxon: Camponotus), 20 (Taxon: Stigmacros), 20 (Taxon: Camponotus dolendus), 20 (Taxon: Camponotus mitis), 20 (Taxon: Polyrhachis), 20 (Taxon: Camponotus), 20 (Taxon: Camponotus), 20 (Taxon: Camponotus), 20 (Taxon: Camponotus), 20 (Taxon: Camponotus crassus), 24 (Taxon: Camponotus barbatus taylori), 24 (Taxon: Echinopla), 26 (Taxon: Nylanderia), 26 (Taxon: Camponotus japonicus), 26 (Taxon: Camponotus mus), 26 (Taxon: Camponotus variegatus), 28 (Taxon: Nylanderia), 28 (Taxon: Camponotus lateralis), 28 (Taxon: Camponotus ligniperda), 28 (Taxon: Camponotus vagus), 28 (Taxon: Lasius alienus), 28 (Taxon: Calomyrmex), 28 (Taxon: Lasius fuliginosus), 28 (Taxon: Lasius pallitarsis), 28 (Taxon: Polyrhachis illaudata), 28 (Taxon: Camponotus kiusiuensis), 28 (Taxon: Camponotus obscuripes), 28 • 29 (Taxon: Acropyga acutiventris), 30 (Taxon: Nylanderia), 30 (Taxon: Nylanderia), 30 (Taxon: Nylanderia), 30 (Taxon: Lasius sakagamii), 30 (Taxon: Lasius alienus), 30 (Taxon: Lasius emarginatus), 30 (Taxon: Lasius flavus), 30 (Taxon: Lasius nearcticus), 30 (Taxon: Prenolepis jerdoni), 30 (Taxon: Lasius niger), 30 (Taxon: Lasius umbratus), 30 (Taxon: Lasius talpa), 30 (Taxon: Nylanderia indica), 30 (Taxon: Pseudolasius), 30 (Taxon: Pseudolasius), 30 (Taxon: Pseudolasius), 30 (Taxon: Pseudolasius), 30 (Taxon: Pseudolasius), 30 (Taxon: Nylanderia), 30 • 32 (Taxon: Acropyga), 31 (Taxon: Prenolepis jerdoni), 32 (Taxon: Prenolepis jerdoni), 32 (Taxon: Prenolepis jerdoni), 32 (Taxon: Camponotus), 32 (Taxon: Camponotus), 34 (Taxon: Anoplolepis gracilipes), 34 (Taxon: Prenolepis jerdoni), 34 (Taxon: Prenolepis jerdoni), 34 (Taxon: Camponotus), 34 (Taxon: Camponotus), 34 (Taxon: Camponotus foreli), 35 (Taxon: Camponotus), 36 (Taxon: Prenolepis jerdoni), 36 (Taxon: Camponotus), 38 (Taxon: Stigmacros), 38 (Taxon: Camponotus festinus), 38 (Taxon: Camponotus), 38 (Taxon: Camponotus), 38 (Taxon: Camponotus), 38 (Taxon: Camponotus), 38 (Taxon: Camponotus), 39 (Taxon: Camponotus), 39 (Taxon: Camponotus), 39 • 40 (Taxon: Camponotus crassisquamis), 39 • 40 (Taxon: Camponotus rufipes), 40 (Taxon: Camponotus atriceps), 40 (Taxon: Camponotus sylvaticus), 40 (Taxon: Camponotus compressus), 40 (Taxon: Camponotus parius), 40 (Taxon: Camponotus balzani), 40 (Taxon: Camponotus bonariensis), 40 (Taxon: Camponotus cingulatus), 40 (Taxon: Camponotus sericeiventris), 40 (Taxon: Polyrhachis hippomanes), 40 (Taxon: Camponotus), 40 (Taxon: Camponotus), 40 (Taxon: Camponotus), 40 (Taxon: Camponotus punctulatus), 40 (Taxon: Camponotus renggeri), 40 (Taxon: Camponotus), 40 (Taxon: Camponotus thraso), 40 (Taxon: Camponotus), 42 (Taxon: Polyrhachis lamellidens), 42 (Taxon: Camponotus aethiops), 42 (Taxon: Polyrhachis ammon), 42 (Taxon: Camponotus alii), 42 (Taxon: Polyrhachis), 42 (Taxon: Polyrhachis rastellata), 42 (Taxon: Polyrhachis hector), 42 (Taxon: Polyrhachis lacteipennis), 44 (Taxon: Notoncus ectatommoides), 44 (Taxon: Camponotus sericeus), 44 (Taxon: Camponotus femoratus), 44 (Taxon: Camponotus), 46 (Taxon: Camponotus consobrinus), 46 (Taxon: Camponotus), 46 (Taxon: Camponotus), 48 (Taxon: Polyrhachis gribodoi), 48 (Taxon: Camponotus), 50 (Taxon: Camponotus pilicornis), 50 (Taxon: Opisthopsis rufithorax), 52 (Taxon: Formica yessensis), 52 (Taxon: Formica candida), 52 (Taxon: Formica exsecta), 52 (Taxon: Formica lugubris), 52 (Taxon: Formica polyctena), 52 (Taxon: Formica pratensis), 52 (Taxon: Formica rufa), 52 (Taxon: Formica sanguinea), 52 (Taxon: Formica truncorum), 52 (Taxon: Cataglyphis setipes), 52 (Taxon: Camponotus), 52 (Taxon: Formica frontalis), 52 (Taxon: Camponotus), 52 (Taxon: Formica picea), 54 (Taxon: Formica cinerea), 54 (Taxon: Formica cunicularia), 54 (Taxon: Formica fusca), 54 (Taxon: Formica gagates), 54 (Taxon: Formica lemani), 54 (Taxon: Formica rufibarbis), 54 (Taxon: Formica montana), 54 (Taxon: Formica japonica), 54 (Taxon: Polyergus samurai), 78 (Taxon: Gigantiops destructor).
The following information is derived from Barry Bolton's Online Catalogue of the Ants of the World.
- FORMICINAE [subfamily of Formicidae]
- Formicariae Latreille, 1809: 124. Type-genus: Formica Linnaeus, 1758: 579.
Lepeletier de Saint-Fargeau, 1835: 197 [Formicites]; Mayr, 1862: 651; Emery, 1877a: 70 [Formicidae]; Bondroit, 1918: 17 [Formicitae]; Wheeler, W.M. 1920: 53, subsequent authors; Bolton, 2003: 20, 93; Ward, et al. 2016: 343.
Mayr, 1862: 651 (genera key); Mayr, 1865: 6 (diagnosis); Handlirsch, 1907: 859 (†fossil taxa catalogue); Dalla Torre, 1893: 171 (catalogue); Emery, 1895j: 772 (synoptic classification); Emery, 1896e: 187 (genera key); Wheeler, W.M. 1910g: 143 (diagnosis); Forel, 1912i: 88 (tribes key); Forel, 1917: 248 (synoptic classification); Arnold, 1920a: 551 (diagnosis); Forel, 1921c: 139 (diagnosis); Wheeler, W.M. 1922a: 210, 691 (diagnosis, tribes key); Emery, 1925b: 2 (diagnosis, tribe key, catalogue); Brown & Nutting, 1950: 127 (venation, phylogeny); Eisner, 1957: 465 (proventriculus morphology); Hung & Brown, 1966: 198 (gastric apex, structure); Bernard, 1967: 267 (diagnosis); Gotwald, 1969: 120 (mouthparts morphology); Wheeler, G.C. & Wheeler, J. 1972a: 41 (diagnosis); Brown, 1973b: 169 (genera, distribution); Wheeler, G.C. & Wheeler, J. 1976b: 62 (larvae, review and synthesis); Snelling, R.R. 1981: 402 (synoptic classification); Wheeler, G.C. & Wheeler, J. 1985: 258 (synoptic classification); Billen, 1986: 173 (Dufour's gland); Dlussky & Fedoseeva, 1988: 77 (synoptic classification); Hölldobler & Wilson, 1990: 9 (synoptic classification, genera keys); Agosti, 1991: 295 (genus group diagnoses); Shattuck, 1992b: 201 (phylogeny); Baroni Urbani, et al. 1992: 317 (phylogeny); Bolton, 1994: 42 (diagnosis, synoptic classification, genera keys); Bolton, 1995a: 1039 (census); Bolton, 1995b: 11 (catalogue); Wenseleers, et al. 1998: 121 (cloacal gland); Dlussky & Rasnitsyn, 2002: 417 (diagnosis for impression fossils); Bolton, 2003: 20, 93 (diagnosis, synopsis); Brady, et al. 2006: 18173 (phylogeny); Moreau, et al. 2006: 102 (phylogeny); Keller, 2011: 1 (morphology, phylogeny); Boudinot, 2015: 51 (diagnosis); Ward, et al. 2016: 345 (reclassification, synopsis); Fisher & Bolton, 2016: 47 (diagnosis).
Regional and National Faunas with Keys
Mayr, 1855: 299 (Austria); Mayr, 1861: 25 (Europe); Mayr, 1868b: 25 (†Baltic Amber); André, 1874: 167 (Europe); Forel, 1874: 22 (Switzerland); Saunders, E. 1880: 203 (Britain); André, 1882a: 126 (Europe and Algeria); Provancher, 1887: 225 (Canada); Cresson, 1887: 94 (U.S.A. genera); Nasonov, 1889: 50 (Russia); Forel, 1891b: 8 (Madagascar genera); Lameere, 1892: 62 (Belgium); Forel, 1892j: 220 (India and Sri Lanka); Bingham, 1903: 308 (India, Sri Lanka and Burma); Ruzsky, 1905b: 100 (Russian Empire); Wasmann, 1906: 7 (Luxemburg); Bondroit, 1910: 481 (Belgium); Wheeler, W.M. 1910g: 560 (North America genera); Stitz, 1914: 80 (Central Europe); Gallardo, 1915: 35 (Argentina genera); Forel, 1915d: 45 (Switzerland); Donisthorpe, 1915d: 184 (Britain); Emery, 1916b: 216 (Italy); Wheeler, W.M. 1916m: 590 (U.S.A., Connecticut); Bondroit, 1918: 17 (France and Belgium); Arnold, 1920a: 552 (South Africa); Kutter, 1920b: 134 (Switzerland); Soudek, 1922: 61 (Czechoslovakia); Lomnicki, 1925a: 160 (Poland); Stärcke, 1926: 118, 146 (Netherlands); Karavaiev, 1927c: 273 (Ukraine); Donisthorpe, 1927b: 205 (Britain); Menozzi & Russo, 1930: 172 (Dominican Republic); Arnol'di, 1933b: 601 (Russia); Menozzi, 1933b: 90 (Israel genera); Karavaiev, 1936: 173 (Ukraine); Smith, M.R. 1937: 865 (Puerto Rico); Stitz, 1939: 230 (Germany); Kratochvíl, 1941: 97 (Central Europe); Novák & Sadil, 1941: 97 (Central Europe); Cole, 1942: 373 (U.S.A., Utah); Smith, M.R. 1943f: 309 (U.S.A., males); Buren, 1944a: 292 (U.S.A., Iowa); Holgersen, 1943b: 173 (Norway); Holgersen, 1944: 199 (Norway); Smith, M.R. 1947f: 599 (U.S.A. genera); van Boven, 1947: 181 (Belgium); Creighton, 1950a: 355 (Nearctic); Kusnezov, 1956: 31 (Argentina); Brown, 1958h: 42 (New Zealand); van Boven, 1959: 11 (Netherlands); Gregg, 1963: 447 (U.S.A., Colorado); Wheeler, G.C. & Wheeler, J. 1963: 160 (U.S.A., North Dakota); Collingwood, 1964: 104 (Britain); Bernard, 1967: 268 (Western Europe); Wilson & Taylor, 1967: 17 (Polynesia); van Boven, 1970b: 26 (Netherlands); Kempf, 1972a: 266 (Neotropical, synoptic classification); Bolton, 1973a: 329 (West Africa genera); Bolton & Collingwood, 1975: 3 (Britain); Snelling & Hunt, 1976: 104 (Chile); Tarbinsky, 1976: 126 (Kyrghyzstan); van Boven, 1977: 126 (Belgium); Kutter, 1977c: 183 (Switzerland); Arnol'di & Dlussky, 1978: 548 (former European U.S.S.R.); Collingwood, 1978: 88 (Iberian Peninsula); Collingwood, 1979: 85 (Fennoscandia and Denmark); Greenslade, 1979: 32 (South Australia genera); Schembri & Collingwood, 1981: 436 (Malta); Prins, 1983: 8 (Southern Africa genera); Allred, 1982: 444 (U.S.A., Utah); Verhaeghe, Deligne, et al., 1984: 106 (Belgium genera); Baroni Urbani, 1984: 81 (Neotropical genera); Gösswald, 1985: 263 (Germany); Collingwood, 1985: 273 (Saudi Arabia); Wheeler, G.C. & Wheeler, J. 1986g: 58 (U.S.A., Nevada); Nilsson & Douwes, 1987: 68 (Norway); Agosti & Collingwood, 1987b: 279 (Balkans); Dlussky, et al. 1990: 124 (Turkmenistan); Kupyanskaya, 1990: 162 (Far Eastern Russia); Morisita, et al., 1991: 10 (Japan); Atanasov & Dlussky, 1992: 49 (Bulgaria); Shattuck, 1992b: 199 (higher classification, phylogeny); Lattke, in Jaffe, 1993: 150 (Neotropical genera); Arakelian, 1994: 76 (Armenia); Wu, J. & Wang, 1995: 125 (China genera); Kupyanskaya, 1995: 332 (Far Eastern Russia); Collingwood & Agosti, 1996: 361 (Saudi Arabia); Seifert, 1996b: 166 (Central Europe); Skinner & Allen, 1996: 41 (Britain); Collingwood & Prince, 1998: 21 (Portugal); Shattuck, 1999: 25, 86 (Australia genera, synopsis); Andersen, 2000: 68 (northern Australia genera); Zhou, 2001b: 165 (China, Guangxi); Czechowski, et al. 2002: 147 (Poland); Aktaç & Radchenko, 2002: 54 (Turkey genera); Yoshimura & Onoyama, 2002: 425 (Japan genera, males); Mackay & Mackay, 2002: 236 (U.S.A., New Mexico); Palacio & Fernández, in Fernández, 2003d: 242 (Neotropical genera); Radchenko, 2005b: 187 (North Korea); Coovert, 2005: 113 (U.S.A., Ohio); Clouse, 2007b: 190 (Micronesia); Seifert, 2007: 150 (North and Central Europe); Terayama, 2009: 202 (Taiwan); Heterick, 2009: 30 (south-western Australia genera); Boer, 2010: 17 (Benelux); Czechowski, et al. 2012: 351 (Poland); General & Alpert, 2012: 71 (Philippines genera key) ; Dlussky & Perfilieva, 2014: 433 (British Eocene species key); Baccaro, et al. 2015: 77, 176 (Brazil genera key, text); Radchenko, 2016: 266 (Ukraine).
The formicomorph subfamilies
Diagnosis Clypeus broad from front to back. Antennal sockets inclined upward toward midline of head (note 1) and situated well behind anterior margin of head. Promesonotal suture usually present and flexible, the pronotum and mesonotum capable of movement relative to each other (note 2). Dorsal cuticular flap of metapleural gland reduced anteriorly and extended posteromedially (note 3). Propodeal lobes absent (note 4). Waist of one segment, the petiole, with complete tergosternalfusion (note 5) (also in male). Helcium sternite small and retracted, overlapped by the tergite (note 6) (also in male). Abdominal segment III (first gastral) without or with partial tergostemal fusion (note 7) (also in male), segment IV without tergosternal fusion. Abdominal segments IV - VII without differentiated presclerites (also in male). Stridulitrum absent from abdominal tergite IV. Abdominal tergites IV - VI hypertrophied with respect to their stemites (note 8). Postpygidial glands absent (note 9). Pretarsal claws without a preapical tooth on the inner margin (also in male). Sting apparatus with furcula reduced and fused to sting base, or lost (note 10). Jugal lobe absent from hind wing of alates. [Synopsis, p. 79.]
Notes (1) Antennal sockets that are horizontal and in the plane of the transverse axis of the head, such as are present in the dorylomorphs and leptanillomorphs, are perhaps best regarded as the plesiomorphic condition. The proceratiines exhibit this feature but there it appears secondary. Inclined to vertical antennal sockets are universal in formicomorphs, myrmeciomorphs, myrmicomorphs and many poneromorphs. (2) In a few Formicinae the promesonotal suture may be present but fused and immobile; very rarely the suture may be vestigial or even absent (Echinopla, a few Polyrhachis species). For distribution of the character through the family see notes under myrmicomorph subfamilies. (3) Metapleural gland is secondarily absent in some Formicinae; for distribution of the character in the subfamily see under Camponotini. (4) Propodeal lobes are present only in Oecophylla and are regarded as independently evolved in that genus. (5) Tergosternal fusion of the petiole is universal in the formicomorph subfamilies, the myrmicomorphs, and the subfamilies Leptanillinae and Leptanilloidinae. In the poneromorphs it occurs in Paraponerini, Proceratiini, Typhlomyrmecini and some Ectatommini; it is partial (anterior) in Amblyoponini. The fusion is regarded as independently acquired in each of these. (6) This is the usual, and probably plesiomorphic, condition in most formicid groups. For different helcium structures, in which the sternite bulges ventrally, see notes under dorylomorph subfamilies and under Myrmicinae. (7) In many dolichoderine genera and in the formicine tribe Plagiolepidini the basal part of abdominal segment III, close to the helcium, has tergosternal fusion with complete obliteration of the suture, but posterior to this the sclerites are not fused and the tergite regularly overlaps the sternite. For distribution of this character through the family see notes under dorylomorph subfamilies. (8) The sternites of abdominal segments IV - VI are small relative to the tergites, and often are entirely ventral; the tergites comprise most of the sides and usually also lap extensively onto the ventral surface of each segment. A similar hypertrophy of abdominal tergite IV occurs in some myrmicine tribes. In Tatuidris (ppAgroecomyrmecini]]), Proceratiini, Loboponera (Ponerini) and some species of Gnamptogenys (Ectatommini), abdominal sternite IV is reduced, but in all these the reduction is associated with a corresponding expansion and strong vaulting of the fourth tergite and reduction of succeeding segments. (9) Postpygidial glands are also absent in Myrmicinae. (10) Fusion or loss of the furcula from the sting apparatus is also found in the dorylomorph subfamilies, some leptanillomorphs and in those Myrmicinae that have reduced stings. It is probably apomorphic in each case.
Comments (i) The three formicomorph subfamilies form a monophyletic group but there is still ambiguity concerning the phylogenetic relationships among them, and concerning the identity of their sister-group. It is suspected, but still remains to be conclusively proved, that the myrmeciomorphs constitute the sister-group, though the extinct †Formiciinae may also be a candidate for this status. (ii) Dolichoderinae and Formicinae are certainly monophyletic but some slight uncertainty must remain about Aneuretinae because there is only one extant monotypic genus that can be intensively studied and all other genus rank taxa are fossils; see comments under Aneuretinae. (iii) With some exceptions (Dolichoderus, many camponotines and particularly Echinopla) the cuticle of formicomorphs is thin and flexible, much more so than in any other group. (iv) The males of many individual dolichoderine and formicine genera can be recognised, but as yet no characters have been found to distinguish male Formicinae as a group from male Do1ichoderinae as a group.
Diagnosis With characters of formicomorph subfamilies. Metacoxal cavities fully closed by a slender bar of cuticle, without a suture in the annulus (note 1) (also in male). He1cium usually attached low on anterior face of abdominal segment III (first gastral) (note 2) (also in male). Helcium tergite dorsally with an extensive U-shaped emargination in its anterior margin (note 3) (also in male). Pygidium simple. Acidopore present at apex ofhypopygium. Formic acid producing glands present. Sting vestigial to absent, not functional; lancets disarticulated from sting. Proventriculus sclerotised (note 4). Pavan I s gland absent. Pygidial gland absent. Cloacal gland usually present (note 5). Pupal cocoons absent or present (note 6). [Synopsis, p. 93.]
Notes (1) Closure of the metacoxal cavities in this way is duplicated only in Dolichoderinae; see note (1) there. (2) Helcium is attached relatively high on abdominal segment III only in Oecophylla. The change from the low position otherwise universal in Formicinae is secondary and is associated with the ability to reflex the gaster over the alitrunk. This is unique to Oecophylla in this subfamily; elsewhere the ability is found only in the myrmicines Crematogaster and, to a lesser extent, Recurvidris. (3) Some species or species groups in genera Polyrhachis and Echinopla (Camponotini) have the emargination of the anterior margin of the helcium tergite reduced, vestigial, or even absent; this is certainly a secondary adaptation. A similar and probably synapomorphic emargination of the helcium is present throughout the Dolichoderinae but does not occur anywhere else in the family Formicidae. (4) Proventriculus is also sclerotised in many Dolichoderinae. (5) Cloacal gland is absent only in Oecophylla, perhaps secondarily so. (6) Formicinae is polymorphic in terms of pupal cocoons. They are regularly absent in nest-weaving forms and sporadically also absent elsewhere in the subfamily.
Comments (i) This is one of the most easily diagnosed and longest-established of the formicid subfamilies. The taxonomic history of Formicinae has recently been extended back to the Cretaceous (Grimaldi & Agosti, 2000). (ii) The earlier higher classifications of Formicinae (Forel, 1912f; Emery, 1925b), which were more or less adhered to until relatively recently, were largely founded on the morphology of the proventriculus. There are now serious doubts about the assumption that the sepalous proventriculus has arisen only once in the subfamily, and about the evolution and distribution of other aspects of the organ's morphology, so that a proventriculus-based classification has become unstable and cannot be maintained (see also comments under Melophorini). The outline of an alternative system has been proposed by Agosti (1991) and is adapted and expanded here.
- Boudinot, B.E. 2015. Contributions to the knowledge of Formicidae (Hymenoptera, Aculeata): a new diagnosis of the family, the first global male-based key to subfamilies, and a treatment of early branching lineages. European Journal of Taxonomy 120, 1-62 (http://dx.doi.org/10.5852/ejt.2015.120).
- LeBrun, E. G., P. J. Diebold, M. R. Orr, and L. E. Gilbert. 2015. Widespread Chemical Detoxification of Alkaloid Venom by Formicine Ants. Journal of Chemical Ecology. 41:884-895. doi:10.1007/s10886-015-0625-3
- Ward, P.S., Blaimer, B.B., Fisher, B.L. 2016. A revised phylogenetic classification of the ant subfamily Formicinae (Hymenoptera: Formicidae), with resurrection of the genera Colobopsis and Dinomyrmex. Zootaxa. 4072(3):343–357. (doi 10.11646/zootaxa.4072.3.4).