Nuptial Flights and Mating

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The vast majority of virgin queens die within hours after leaving the mother nest. Most are destroyed by predators (Figure 3-4) and hostile workers of alien nests, with the others being variously drowned, overheated, and desiccated. In species with large nest populations, such as the leafcutter ants (Atta) and fire ants (Solenopsis), it is not uncommon for one colony to release hundreds or thousands of the young winged queens in less than an hour. If the surrounding area is dominated by stable, mature colonies, only one or two of the queens might become the progenetrices of new colonies. Most of the rest will die before they can construct a first shelter--or even before they can find a mate. In an unusual study of its kind, Whitcomb et al. (1973) have produced a catalog of the many kinds of predators that decimate young queens of the red imported fire ant Solenopsis invicta. The few individuals that navigate all the dangers must also avoid breeding with males of other species, thus producing inviable or sterile offspring.

Figure 3-4. The mortality of colony-founding queens is extremely high. In this case a recently dealated queen of Pogonomyrmex maricopa has been captured by a crab spider. (From Holldobler, 1976b.).

It follows that the brief interval between leaving the home nest and settling into a new, incipient nest is a period of intense natural selection among queens, a dangerous odyssey that must be precisely timed and executed in order to succeed. We should expect to find an array of physiological and behavioral mechanisms that enable the young queens simultaneously to avoid enemies, get to the right habitat on time in order to build a secure nest, and mate with males of the same species. Field studies have shown that such specialized traits exist in abundance.

As also expected from the evolutionary argument, mating patterns vary greatly from one species to the next. However, most of the patterns thus far studied fall into one or the other of two broad classes, or "syndromes" (Hölldobler and Bartz, 1985). In the first, the female-calling syndrome, the females, which are often wingless and sometimes just fertile workers, do not travel far from the nest. Standing on the ground or low vegetation, they release sex pheromones to "call" the winged males to them (Figure 3-5). This pattern is displayed by Amblyopone and Rhytidoponera, which are members of the phylogenetically primitive subfamily Ponerinae (Haskins, 1978); presumably also by the very primitive Nothomyrmecia macrops (Hölldobler and Taylor, 1983); at least one pseudomyrmecine, the Neotropical acacia ant Pseudomyrmex ferrugineus (Janzen, 1967); and the socially parasitic species of the myrmicine genera Doronomyrmex, Formicoxenus, Harpagoxenus, and Leptothorax (Buschinger, 1968a,b, 1971a,b, 1975b; see Figure 3-6).

In general, the colonies of female-calling species are typically small at maturity, with 20 to 1,000 workers, and produce relatively few reproductives. So far as known the females mate only once. An unusual variation on this pattern is followed by the Florida harvester ant Pogonomyrmex badius. Females gather on the surface of their home nest and are inseminated by males; afterward they fly off to start new colonies. Van Pelt (1953) thought that the males came from the same nest as the females with whom they copulate, but S. D. Porter (personal communication) observed that they usually fly for about a quarter-hour first before settling on a nest different from their own. Porter observed one case in which a male mated with two females after alighting.

Figure 3-5. An example of one of the two major categories of mating behavior in ants, the female-calling syndrome, in the Australian ponerine Rhytidoponera metallica. (a) An ergatoid or worker-like female (black) assumes the calling posture, during which she releases a sex pheromone from the pygidial gland located between the VIth and VIIth abdominal tergites. (b) A male approaches her and touches her with his antennae. (c) The male mounts the female, grasps her by the prothorax, and extrudes his copulatory organ in search of the female's genitals. (d) Copulation occurs. (From Holldobler and Haskins, 1977; drawing by T. Holldobler-Forysth.)

The second combination of traits during mating is the male-aggregation syndrome. Males from many colonies gather at specific mating sites, usually prominent features of the landscape such as sunflecked clearings, forest borders, hilltops, the crowns of trees, and even the tops of tall buildings. Sometimes, as in some species of Lasius and Solenopsis, the males cruise in large numbers at characteristic heights above the ground. The females fly into the swarms, often from long distances, in order to mate (see Figures 3-7 through 3-9 and Plate 2), and afterward they typically disperse widely before shedding their wings and excavating a nest. The winged queens and males of the fire ant Solenopsis invicta, for example, fly up to heights of 250 meters or more; 99 percent then descend to the ground within a 2-kilometer radius of their origin, while a very few travel as far as 10 kilometers. The ability of a single mature colony to disseminate fertile queens in many directions over long distances is one of the reasons the fire ant is so difficult to eradicate (Markin et al., 1971). Male-aggregation species typically differ from those utilizing female calling in two other key respects: the mature colonies are large, containing from several thousand to over a million workers and producing hundreds to thousands of reproductive adults yearly, and multiple insemination is common. An unusual reversal of the usual swarming procedure was recently discovered in some Pheidole species of the southwestern United States: the winged queens gather in aerial swarms, where they maintain a more or less uniform distance from each other while attracting males with pheromones. The males fly into the female swarms and mate with individual females (Hölldobler, unpublished). Swarms of variable composition, some predominantly male and others predominantly female (occasionally exclusively female), have been reported by Eberhard (1978) in the coccid-tending formicine Acropyga paramaribensis of northern South America.

Figure 3-6. Sexual behavior of the socially parasitic ant Leptothorax pacis. a pair in copulation. (From Buschinger, 1971b).

Ant species can be classified another way into two broad types. When the males alight on the surface of the mating site, either in response to female calling or in swarms to compete directly with one another, they are often typically large and robust in form and possess well-developed mandibles. In contrast, males that gather in aerial swarms are usually (but not invariably) smaller relative to the queen than are males of the first type. Also, their mandibles are reduced in size and dentition, sometimes consisting of nothing more than vestigial lobate or strap-shaped organs. An example of this type is the small myrmicine Pheidole sitarches of the southwestern United States. Up to 50 males form circular swarms that hover from a few centimeters to two meters above the surface of woodland clearings. The virgin queens fly in slow, even circles through the aggregations until mounted in midair by a male, whereupon the pair cease flying and spiral to the ground together to complete the copulation (Wilson, 1957b).

The swarms of some ant species are among the more dramatic spectacles of the insect world. W. W. Froggatt (in Wheeler, 1916c) describes the flight of the giant Australian bulldog ant Myrmecia sanguinea as follows:

"On January 30th, after some very hot, stormy weather, while I was at Chevy Chase, near Armidale, N.S.W., I crossed the paddock and climbed to the top of Mt. Roul, an isolated, flat-topped, basaltic hill, which rises about 300 feet above the surrounding open, cleared country. The summit, about half an acre in extent, is covered with low "black-thorn" bushes (Bursaria spinifera). I saw no signs of bull-dog ant nests till I reached the summit. Then I was enveloped in a regular cloud of the great winged ants. They were out in thousands and thousands, resting on the rocks and grass. The air was full of them, but they were chiefly flying in great numbers about the bushes where the males were copulating with the females. As soon as a male (and there were hundreds of males to every female) captured a female on a bush, other males surrounded the couple till there was a struggling mass of ants forming a ball as large as one's fist. Then something seemed to give way, the ball would fall to the ground and the ants would scatter. As many as half a dozen of these balls would keep forming on every little bush and this went on throughout the morning. I was a bit frightened at first but the ants took no notice of me, as the males were all so eager in their endeavors to seize the females."

Donisthorpe (1915) tells of the mass flights of the abundant Myrmica rubra from the distinctively British viewpoint of an earlier observer:

"Farren-White in 1876 observed a swarm of ants near Stonehouse rising and falling over a small beech tree. The effect of those in the air--gyrating and meeting each other in their course, as seen against the deep blue sky--reminded him of the little dodder, with its tiny clustered blossoms and its network of ramifying scarlet threads, over the gorse or heather at Bournemouth. He noticed the swarm about thirty paces off, and it began to assume the appearance of curling smoke; at forty paces he could quite imagine the tree to be on fire. At fifty paces the smoke had nearly vanished into thin air."

A still different mating pattern was described in the Australian formicine species Notoncus ectatommoides by W. L. Brown (1955a):

"In a cropped lawn at Montville, numerous small holes appeared, each opened by workers and accompanied by a minute pile of dark earthen particles. From these holes, males began to issue almost immediately in numbers, until within a few minutes there had accumulated on the surface a surprisingly large number of this sex and also a few workers. The males traveled aimlessly over the sward in low, flitting flight from one blade of grass to another, never rising more than a foot or so from the ground. Movement seemed to take place at random in all directions. Suddenly, however, the males of one area all rushed simultaneously to a single focal point, which proved to be a winged female emerging from a small hole. In a few seconds, the female was surrounded by a dense swarm of males in the form of a ball, which at times must have exceeded 2 cm in diameter. This ball moved in a half-tumbling, half-dragging motion over and among the densely packed grass blades, and held together for perhaps 20 seconds, after which the female escaped, flying straight upward. She appeared not to be encumbered by a male, and no males were seen to follow her for more than a foot above the ground; she flew steadily, and soon passed out of sight.

Meanwhile, the lawn had become dotted with similar balls of frenzied males, each surrounding a female in a fashion similar to the first. Obviously, many more males than females were involved in this particular flight. On each occasion, the female left the ball after 20-30 seconds and flew straight upward."

In a similar fashion males and females of Formica obscuripes conduct nuptial swarms on the ground. Talbot (1972) observed them flying to "swarming grounds" near their nests which were maintained throughout the nuptial flight season and perhaps even from year to year. The males fly back and forth above the ground searching for females which "stand on grasses, forbs or bushes," and apparently signal their presence to the males by pheromones.

No encompassing theory exists to explain the extreme variation in the patterns of mating behavior so far observed. However, a close examination of individual species reveals details that clearly contribute to the greater success of the sexual castes. For example, flying queens of the formicine Lasius neoniger stay strictly within open fields, the exclusive habitat of the earthbound colonies. Fewer than one percent make the mistake of venturing into adjacent woodland, a habitat dominated by the otherwise closely similar Lasius alienus. In one experimental study (Wilson and Hunt, 1966), newly inseminated and flightless queens were labeled with radioactive material for easy tracking and displaced to woodland sites. They attempted to crawl out but were unable to do so. In other words the Lasius queens depend on controlled flight patterns to survive.

Like orientation, the timing of the flights is important for successful mating and colony foundation. Flights conducted as part of the female-calling syndrome do not appear to be well synchronized at the level of either the colony or the population of colonies. The search by airborne males for solitary calling females in fact resembles that of many solitary wasps (Buschinger, 1975; Haskins, 1978). In contrast, flights leading to male aggregation are tightly synchronized within the colony as well as among colonies of the same species.

The manner in which this coordination is achieved is typified by Pogonomyrmex harvester ants of the southwestern United States. The process has been described by Hölldobler (1976b). Just prior to take-off, males and females move restlessly in and out of the sandy crater nests or gather in clusters around the entrance, as shown in Figure 3-7. This preflight activity is especially pronounced in Pogonomyrmex maricopa, a morning flyer, the queens and males of which evidently need more time to warm up before taking wing. As the time of departure approaches, the reproductives run back and forth in mounting intensity. Now, in a frenzy, they climb up and down on grass leaves or small bushes around the nest. At this point many more workers pour out of the nest, running excitedly around the nest and attacking any moving object encountered (including the careless myrmecologist). When the first reproductives try to take flight, the workers at first delay many of them by pulling or carrying them back to the nest. However, once the flight is in full progress, workers cease to interfere. Although the timing of the take-off overlaps considerably between the two sexes, the males generally fly from the nest first. Once aloft both sexes appear at first to drift with the wind, but after a few seconds they take a course upwind or across the wind. Soon afterward they arrive at the swarm sites, centered on conspicuous landmarks such as tree crowns and the tops of hills or (in the case of Pogonomyrmex rugosus) merely flat local areas in the desert.

A similar marching order is observed by the carpenter ant Camponotus herculeanus, which nests in the trunks of both living and dead trees in the boreal forests of Eurasia and North America. Males leave before the queens, although the periods broadly overlap. The early departure of the winged forms is inhibited by the workers, who drag or carry many back to the nest entrance (Figure 3-10). However, when the males do succeed in taking flight, they discharge a pheromone from their mandibular gland. The concentration of this substance is highest at the peak of male activity--the gland emission can now be smelled readily by humans--enough to trigger the mass take-off of the females (Figure 3-11). Blum (1981b) reports methyl 6-methylsalicylate and mellein as two of the three components of the secretion. This pleasantly aromatic combination is shared by most other species of Camponotus, but considerable differentiation nevertheless is achieved by the addition of other substances, such as octanoic acid and methyl anthranilate, according to species (see also Lloyd et al., 1984). A similar function may be accomplished by vibrational signals rather than pheromones in Pogonomyrmex harvester ants. Both males and virgin queens stridulate just before and during take-off, running the sharp posterior rim of their postpetiole over the actively moving, striated file on the first gastric tergite (Markl et al., 1977).

Many entomologists, including especially Kannowski (1959a, 1963) and Weber (1972), have observed that each ant species, at least those displaying the male-aggregation syndrome, swarms at a precise time in the 24-hour diel cycle; and the time differs among species. Under controlled laboratory conditions, McCluskey (1958, 1965, 1967, 1974) and McCluskey and Soong (1979) demonstrated in fact that the rhythms of males are generally if not universally circadian and endogenous. Once set in a laboratory regime of 12 hours light alternating with 12 hours dark, the rhythms persist for up to a week in total darkness. They are also quite precise. McCluskey found that males of the harvester ant Messor (= Veromessor) andrei increase in movement during the last hour of darkness, then peak during the first hour of light. Throughout the remainder of the 24-hour cycle they are quiescent, usually stirring themselves only to groom, solicit food from the workers, or walk sluggishly about the nest. Males of the Argentine ant Iridomyrmex humilis, in contrast, are most active at the very end of the light period. Similarly distinctive rhythms, each spanning only one or two hours, have been documented by McCluskey and his co-workers across a wide diversity of species from four subfamilies (Ponerinae, Myrmicinae, Dolichoderinae, and Formicinae), including some that are wholly nocturnal.

Queens of at least two species, Pogonomyrmex californicus and Mesor (= Veromessor) pergandei, also display circadian rhythms, and these are more or less synchronous with those of the males (McCluskey, 1967; McCluskey and Carter, 1969). In the case of P. californicus at least, the rhythm persists even after the female has flown and lost her wings. But it ceases when she is mated.

In summary, the time of day in which flights occur is programmed by a species-specific diel rhythm. But what determines the particular day on which the flights occur? Several studies, including that by Boomsma and Leusink (1981), have shown that weather conditions play a major role in the timing of nuptial flights. One of the commonest triggering stimuli is rain, especially in species that occur in dry habitats such as deserts, grassland, and forest clearings. A typical species in this respect is Lasius neoniger, one of the most abundant ants in abandoned fields and other open environments in eastern North America. This small formicine emerges in immense swarms in the late afternoon in the second half of August or early September. The flights almost always occur within 24 hours after moderate or heavy rainfall on warm, humid days with little wind. For an hour or so the air seems filled with the winged ants. They rise from the ground like snowfall in reverse. After mating, the queens find themselves on moistened soil that is easier to excavate. They are also protected from desiccation due to overheating (Wilson, 1955a). A very similar pattern is followed by the North American leafcutter ant Atta texana, except that the flights occur well before dawn, between 0300 and 0415 hours (Moser, 1967a).

Because there are relatively few "best days" in which the young queens can be successfully launched, species belonging to the same genus are likely to swarm at the same time and location. In one respect this is a favorable result, since an apparent function of mass emergence and swarming in cicadas, termites, and other insects is the reduction of mortality by overloading predators (Wilson, 1975b). But in another respect it can be detrimental. In the tumult of the swarms, with males struggling to copulate with each female encountered, there is a strong likelihood of interspecific hybridization resulting in either sterility or the production of less viable hybrids. Applying the standard argument from natural selection theory, this circumstance favors the evolution of premating isolating mechanisms. The conventional explanation does seem compatible with a great deal of evidence. Species belonging to genera as phylogenetically diverse as Myrmecia, Pheidole, Solenopsis, and Lasius have been observed to conduct their nuptial flights within the major habitats occupied by the colonies, thus automatically avoiding sexual contact with closely related species limited to other major habitats. How widespread and efficient this isolating mechanism is among ants in general has not been determined. But it cannot be the sole device in deserts, savannas, and tropical moist forests, where large numbers of congeneric species nest closely together. To take an extreme case, in many forest localities in the Amazon Basin, thirty or more species of Pheidole can be found within a single plot of a few square kilometers. Another intrinsic isolating mechanism is differentiation in the preferred mating site within the major habitat. Among the sympatric species of Pogonomyrmex of Arizona, Pogonomyrmex desertorum and Pogonomyrmex maricopa congregate on bushes and trees, while Pogonomyrmex barbatus and Pogonomyrmex rugosus gather at different sites on the ground. In addition, males mark the sites with secretions from their mandibular glands, and apparently the females and other males are attracted by volatile pheromones contained in the material (Hölldobler, 1976b). It is possible (but not yet experimentally verified) that the pheromones are species-specific and serve as an additional isolating device.

Many congeneric species are further separated by the timing of their mating flight, either the season of the year or the hour of the day. In Figures 3-12 and 3-13 we have presented two sets of data from army ants that suggest just such a mutually repulsing spread of flight times across the seasons and the daily cycle respectively. The males of army ants, on which the data were based, fly for an unknown distance before entering the columns or bivouacs of alien colonies belonging to the same species. If the receptiveness of the workers is synchronized by the same circadian rhythm, even the hours of flight can serve as an effective barrier to "mistakes" and interspecific hybridization. Such staggering in the diel flight schedule appears to be common among ants. In Michigan, for example, Myrmica emeryana flies between 0600 and 0800 hours, Myrmica americana between 1230 and 1630 hours, and Myrmica fracticornis between 1800 and 1930 hours (Kannowski, 1959a). Similarly, in Arizona Pogonomyrmex maricopa flies between 1000 and 1130 hours, Pogonomyrmex barbatus between 1530 and 1700 hours, and Pogonomyrmex rugosus between 1630 and 1800 hours. As morning flyers, the Pogonomyrmex maricopa queens appear to be at some disadvantage. The heat of midday prevents them from beginning nest excavation for three or four hours, during which time they are subject to higher predation than the other species (Hölldobler, 1976b). Some of the most closely related European species of Leptothorax swarm at different times of the day; others come into contact, and occasionally hybridize (Plateaux, 1978, 1987).

Another potential advantage of synchronous nuptial swarming is the increase in the numbers of colonies participating and hence the degree of outbreeding. The sparse data on allozyme variation in ants collected so far indicates that outbreeding is indeed nearly total (Craig and Crozier, 1979; Pamilo and Varvio-Aho, 1979; Pearson, 1983; Ward, 1983a). Hence mating is either effectively at random, as demonstrated in experimental choice tests with Pogonomyrmex californicus by Mintzer (1982a), or disassortative, that is, directed away from nestmates.

The glandular sources of sex pheromones produced by female ants have been identified only for a few species. The reproductive females of Rhytidoponera metallica call males with a sex attractant from the pygidial gland, an intersegmental structure between the VIth and VIIth abdominal tergites (Hölldobler and Haskins, 1977). Although some of the contents of this gland have been chemically identified (Meinwald et al., 1983), the specific behavior-releasing components have not yet been established experimentally. In several myrmicine species glands associated with the sting apparatus have been pinpointed as the sources of female sex pheromones. Virgin queens release a male attracting pheromone from the poison gland in the myrmicines Xenomyrmex floridanus (Hölldobler, 1971); Harpagoxenus sublaevis (Buschinger, 1972a); Doronomyrmex kutteri and Doronomyrmex pacis (Buschinger, 1975b; see Figure 3-6); and Formicoxenus nitidulus (Buschinger, 1976a,b).

Buschinger (1972b) was also able to demonstrate that males of Doronomyrmex kutteri and D. pacis react to the other species' female sex pheromones, and that hybridization is possible in laboratory experiments. In the field, however, both species, which occur sympatrically, appear to be sexually isolated by different diel rhythms in mating activity. In general, specificity in sexual communication is consistent with phylogenetic relationships among the leptothoracines. The Canadian slavemaker Harpagoxenus canadensis shows the same mating behavior as the European H. sublaevis, and males of both species respond to the other species' female sex pheromones. Very similar sexual behavior and responses to sex pheromones have been described in several other social parasites of the "subgenus Mychothorax" of Leptothorax, whose hosts, like those of H. canadensis and H. sublaevis, also belong to the "subgenus Mychothorax." The same is true of at least some non-parasitic members of the subgenus. In fact, there appears to be no pheromone specificity among the Leptothorax species. In contrast, Protomognathus americanus males do not respond to H. canadensis or H. sublaevis pheromones. This anomaly suggests that P. americanus may be more closely related to its host of the "subgenus Leptothorax" than to the other genus of Harpagoxenus or the "subgenus Mychothorax" (Buschinger, 1975b, 1981; Buschinger and Alloway, 1979).

Poison gland secretions of Pogonomyrmex females also elicit attraction in males (Hölldobler, 1976b). In Monomorium pharaonis, on the other hand, the female sex pheromone is derived from the Dufour's gland and the bursa pouches (Hölldobler and Wüst, 1973).

Male ants are richly endowed with exocrine glands (Hölldobler and Engel-Siegel, 1982), but little is known about their function. One important fact, noted earlier, is that Camponotus herculeanus males discharge mandibular gland contents when departing from the nest that stimulate the virgin reproductive females to launch as well the nuptial flight. A variety of compounds of the mandibular gland secretions of several Camponotus species have been identified (for review see Blum, 1981b), but it is not yet clear which substance or combination of compounds elicits the behavior. Similarly, the males of Lasius neoniger discharge their mandibular gland contents sometime during the nuptial flight (Law et al., 1965), but the precise timing and function remain unknown.

Males of Pogonomyrmex discharge mandibular gland secretions when arriving at the mating sites. The collectively discharged pheromone appears to attract the virgin females to the lek (Hölldobler, 1976b). It is possible that in other species where males have well-developed mandibular glands and distinct blends of compounds, the secretions also function in promoting aggregation and competition. Examples include Lasius' and Acanthomyops (Law et al., 1965), Camponotus (Brand et al., 1973b,c; review by Blum, 1981), Calomyrmex (Brown and Moore, 1979), Myrmecocystus (review by Blum, 1981b), Tetramorium caespitum (Pasteels et al., 1980), and Polyrhachis doddi (Bellas and Hölldobler, 1985).

A hypothesis concerning a possible novel role of male pheromones in sexual selection in army ants has recently been proposed by Franks and Hölldobler (1987). A detailed morphological examination of the reproductives has shown a close resemblance of conspecific males and females. Males are remarkably queen-like. They are large and robust, and their long, cylindrical abdomens are partially filled with an impressive battery of exocrine glands similar in form and location to those of females. Because queens are flightless and never leave their colony, males must fly between colonies and run the gauntlet of the workers before they approach the queen. For this reason, the workers can choose which males will be admitted and which virgin queens will be inseminated by the males. Army ant workers might therefore be involved in a unique form of sexual selection in which they choose both the matriarch and patriarch of new colonies. If this interpretation is correct, males resemble queens not because they are deceitful mimics; instead, under the influence of sexual selection they have come to use the same channels of communication to demonstrate their potential fitness to the workers as those used by queens.

Worker involvement in sexual selection might not be restricted to the army ants. Wheeler (1910a) noted that males of Leptogenys elongata are also accepted into alien colonies to mate with the wingless ergatoid females, and Maschwitz and Mühlenberg (1975) observed that males run along permanent foraging trails of Leptogenys ocellifera, apparently in an attempt to find access to ergatoid females. It may therefore be significant that Hölldobler and Engel-Siegel (1982) discovered very large exocrine sternal glands in Leptogenys males. Some other ponerines have ergatoid queens and therefore are not likely to engage in ordinary nuptial swarms, including species of Diacamma, Dinoponera, Megaponera, and Ophthalmopone. Longhurst and Howse (1979a) observed that males of Megaponera foetens enter the nests of alien colonies, after utilizing recruitment pheromone trails laid by workers to guide them to the nest. No information is available, however, on the exocrine glandular system of Megaponera or for that matter most other ponerine genera. Males of Ophthalmopone berthoudi also enter strange nests after dispersal flights, but so far as known do not follow odor trails--O. berthoudi workers in fact forage in an exclusively solitary manner and hence are less likely to lay recruitment trails of any kind (Peeters and Crewe, 1986a, 1987).

Male ants compete for females in a rigorous fashion, whether they are orienting to calling females in the primitive manner, flying in aerial swarms, or massing on the surface of the ground and vegetation. The competitive nature of mating is vividly illustrated by Pogonomyrmex rugosus (see Plate 2). The males gather in what can properly be called leks of the vertebrate kind. That is, the males occupy the same site year after year, use pheromones to attract other reproductives of the same species, and then compete with one another for access to the females. In the desert near Portal, Arizona, Hölldobler (1976b) was able to locate only one such site in an area of approximately 120,000 m2. The mating arena covered 4800 m2 of completely flat land unmarked by any distinctive physical features. The winged reproductives approached the arena upwind, which may suggest the presence of an olfactory cue. The first individuals to arrive (at around 1630 hours) were males, which alighted and began to race about in a frenzied manner. Soon afterward the first females alighted. They were immediately surrounded by three to ten males, as shown in Figures 3-8 and 3-9. At the height of the activity thousands of such mating clusters carpeted the ground, in densities as high as 50 per square meter. The queens actively terminated mating after several copulations, and stridulated when prevented from leaving by other suitors. This stridulatory vibration evidently served as a "female liberation signal" that communicated the female's non-receptivity to approaching males and induced them to cease pursuit (Markl et al., 1977). The females then climbed onto grass leaves to launch their flights or else flew directly from the ground. Some landed a short distance away, but others traveled at least 100 meters and possibly much farther. Each then shed her wings and began to excavate a nest chamber in the soil (Figure 3-14).

The general activity at the Pogonomyrmex rugosus mating site lasted about two hours, ending completely by 1900 hours as darkness approached. The males then withdrew into shelters around the mating site, such as crevices beneath grass clumps or little cavities in the soil. There they remained clustered overnight and through the following day until 1500-1600 hours, when they resumed activity. As on the previous day, new males flew in to the site to swell the population, and shortly afterwards females began to arrive. This cycle was repeated on three more consecutive days.

The ant leks differ from those of the sage grouse, hammerheaded bats, and Hawaiian Drosophila (see for example, Bradbury, 1985) in one important respect. Ant males are constrained in a way that vertebrates and fruit flies are not: each male ecloses from the pupa into full maturity with all of the sperm that he will ever possess. Dissections of males from phylogenetically divergent genera such as Nothomyrmecia, Camponotus, Lasius, Myrmica, and Pogonomyrmex reveal that the males' testes have degenerated and all of the sperm have migrated to the expanded vas deferens (Hölldobler, 1966, and unpublished data). When the male mates, it discharges most or all of the sperm together with the secretions of the mucus gland; he is thus incapable of additional inseminations (see Figure 3-15). As a result, reproductive success in male ants does not increase with repeated copulations, as it does with other kinds of insects whose males continuously replenish their sperm supply. Furthermore, it does not appear, from the few cases known, that males have enough sperm to inseminate more than one female. In cases where the queen is destined to produce very large numbers of offspring, one male is not even able to supply all of her needs. In Atta sexdens, for example, each newly eclosed male has between 40 and 80 million spermatozoans, while each newly mated female contains between 200 and 310 million spermatozoans in her spermathecae (Kerr, 1962). Fire ant queens (Solenopsis invicta) receive a supply of about 7 million sperm initially, which they gradually parcel out over a period of almost 7 years until the supply is exhausted (Tschinkel and Porter, 1988). Male ants are thus under strong pressure in natural selection to husband their sperm carefully.

One obvious question concerning the ultimate reproductive success of males is whether it is better for a male to invest all of his sperm in a single female or else to copulate with several females. As Hölldobler and Bartz (1985) pointed out, it is important to note that in ants, unlike other nonsocial species, a male's sperm does not all go towards effective reproduction. This is because in order for an ant colony to begin to produce any reproductive forms, it first must produce many workers. In most advanced ant societies workers are rarely reproductive, and because workers are females derived from fertilized eggs, a substantial portion of a male's sperm is used in colony growth and maintenance rather than in direct production of new queens. The consequent trade-off for a male is obvious. If he inseminates only a single female, and if she mates with no other male, then the male is certain to father any reproductives that she eventually produces. However, mortality of colony-founding females is extremely high. Hence a male that inseminates only a single female puts all of his sperm in one fragile basket. If he were to inseminate several females, on the other hand, he increases the chance that his sperm will end up in a successful foundress of a colony. In this case, however, he might decrease the chance that his sperm is used by the queen to make alates. On the other hand, if males do inseminate several females, there may be selection favoring males whose sperm mixes with other males' sperm in the females' spermathecae. Mixing sperm increases the chance that each male will have at least some offspring among the new crop of alate queens. From allozyme variation studies in multiple mating ant species, it does in fact appear that workers in colonies are fathered by several males (Pamilo, 1982b,c; Pearson, 1983; Ward, 1983a).

In species with large mature colonies, whose females must mate with several males in order to acquire sufficient sperm, males seldom attempt to monopolize females (Cole, 1983b). In many other kinds of insects, and other organisms as well, sperm competition is an important selective force, and males are often favored to ensure that no other male copulates with his mate (Parker, 1970). In multiple mating ant species, however, a male that prevents his mate from mating again may well prevent her from acquiring enough sperm to generate a mature colony, that is, one large enough to produce reproductives. Males in these species therefore should be selected to mate with females that are already mated. The active vying for position in waiting lines behind copulating pairs in Pogonomyrmex species indicates at least that males do not discriminate against previously mated females, but it does not prove the optimum multiple mating hypothesis.

To summarize, male ants are faced with two limiting resources: a restricted number of females available for mating, and a finite supply of sperm that suffices for only one or at most several matings. An expected consequence in evolution is the fierce competition of the kind observed in the Pogonomyrmex leks. Davidson (1982) observed that Pogonomyrmex barbatus and Pogonomyrmex desertorum males indiscriminately seize females and attempt to mate, while the females actively resist copulation. As a result, large males are disproportionately successful at gaining access to mates. In addition, large females mate even more disproportionately with large males. And still further, the average size of males produced by an individual colony depends on the total number of reproductives reared in a given season, which in turn is a function of the size and vigor of the colony. In short, the bigger the colony, the more likely its individual males are to succeed in the mating arenas. Why hasn't this selection pressure created ever larger males in evolution? Davidson offers two reasons: larger males mean fewer males per colony, an obvious trade-off in colony fitness, and very large males (as opposed to merely large ones) have been observed to lose some of their advantage to those slightly smaller. The result is the existence of an optimum male size in Pogonomyrmex.

A confounding bit of data reported by Davidson (1982) is that not only do larger females tend to mate with larger males, but smaller females tend to mate with smaller males. If sexual selection is operating such that females choose larger males, why do small females not also choose to mate with larger males? The answer to this question may be that males are selected to be choosy as well. As we have pointed out male ants have only so much sperm at their disposal, and they cannot afford to be profligate. Selection may favor males who compete for larger females because there is a better chance that large females will survive to produce a mature colony. A result of the competition would be that the smaller, less competitive males must settle for the smaller, less desirable females.

The evolution of male biology has been subjected to few rigorous studies, and most questions concerning trends and optimality in its evolution remain unanswered. We are in a somewhat better position with reference to both data and theory on the number of female matings. As documented in Table 3-1, which includes most or all of the information available, some fraction of the queens of fully three-quarters of all species copulate with more than one male. It is also true, as revealed by allozyme marker studies (Pamilo, 1982b,c; Pearson, 1983; Ward, 1983a), that the sperm from different fathers contribute randomly to fertilization. Cole (1983b) established that multiple matings (polyandry) occurs more frequently in species with large colony size. He concluded, as West-Eberhard (1975) and a few other previous writers had suggested earlier on more intuitive grounds, that polyandry was therefore likely to be a response to the need on the part of queens in large colonies for more spermatozoans than one male can provide. In a study of 25 species in 5 subfamilies, Tschinkel (1987a) added stronger evidence from the number of sperm acquired by queens. In comparisons across species, the number of sperm increases very rapidly with the number of ovarioles. It ranged in Tschinkel's sample from a few tens of thousands in Ponera and Hypoponera, which form small, slow-growing colonies, to 400 million in the leafcutter Atta texana, which attains populations of over a million workers at a time. At another level, the number of sperm stored per ovariole (as opposed to per queen) increased from 2,000 for queens with only six ovarioles to about 30,000 for queens with about 200 ovarioles.

Not satisfied with the intuitively simplest explanation, however, Crozier and Page (1985) went on to employ the method of multiple competing hypotheses to test the adaptiveness of polyandry. They constructed no less than eight such explanations (some admittedly very improbable) to account for the trend documented by Cole. The explanation of limited male contribution favored by Cole and earlier authors was downgraded, because "males of species with big females are generally larger than those with small females, so there is no absolute bar to male size (within reason!)." This does not seem to be a very strong counterargument. Aerial swarmers can potentially benefit from smaller size, which confers greater agility during the approach to incoming queens. Also, as we have noted with reference to mating in Pogonomyrmex, there is a trade-off between male size and male numbers, still poorly analyzed, that might contribute to the preferred production by colonies of smaller males.

Hence it is prudent to keep alive the limited-sperm hypothesis of polyandry. Crozier and Page, after discarding most of the other competing explanations, hold on to three (not counting the limited-sperm hypothesis, which we favor) as both inherently plausible and compatible with the correlation between colony size and polyandry. The first is that caste determination might be genetic, and if so polyandry would allow fuller expression of the caste system in each colony. It follows that species with more complex caste differentiation (a trait associated with large colonies) should be more polyandrous than species with simpler caste systems. As Crozier and Page note, there is no evidence for genetic caste determination in ants to the present time, although recent evidence suggests some kind of genetic predisposition toward various forms of labor specialization in honeybees (Calderone and Page, 1988; Frumhoff and Baker, 1988; Robinson and Page, 1988). All of the many substantial studies to date have implicated a single genotype with multiple developmental pathways controlled by nutritive and other environmental factors (see Chapter 8). The second surviving explanation in the Crozier-Page analysis is that polyandry maximizes the production of divergent worker genotypes, quite apart from caste phenotypes, and hence the range of environmental conditions that a colony can tolerate. Broad-niche species, most often those possessing large colonies, should be more polyandrous than species with narrow niches. This broad relationship has not yet been tested empirically. The third favored hypothesis is that multiple matings reduce the chances of disaster due to the production of diploid males. Males of Hymenoptera, it will be recalled, ordinarily come from unfertilized eggs and are determined as males simply by being haploid, that is, having only one set of sex-determining genes. When one or a very few loci are involved in the process, and recessive male-determining alleles exist, it is also possible to get males from fertilized eggs, the so-called diploid males. The queen of an ant colony can ordinarily control male egg production precisely by opening or closing her spermathecal valve "at will," thus determining whether an egg in the vaginal passage is fertilized. But she has no control whatever over the production of diploid male eggs, because the effort to produce females will still result in a fixed percentage of males by Mendelian chance alone. This circumstance does not matter much if the strategy of the colony is to produce males during early stages of colony growth (beyond the very earliest, fragile stage of colony founding), a not uncommon event in species with a small mature colony size. But it can add a substantial energetic burden on species whose strategy is to hold off production of drones until the colony is large. By mixing sperm from multiple males, the variance of such a load is reduced. In other words, more colonies are likely to have some diploid males, but on the average they are less likely to produce large numbers of diploid males.

Reasoning in another mode, Woyciechowski and Lomnicki (1987) proposed that multiple matings prevent workers from producing male offspring. According to their model of kin selection, workers are at an advantage if they produce sons and care for nephews in the presence of a mother queen who mated only once, but they should avoid personal reproduction and care for brothers in the presence of a mother queen who mated several times. The existing data on queen mating patterns and worker reproduction are not adequate to test the hypothesis.

More recently, Sherman et al. (1988) have argued that the role of polyandry is to increase genetic variation within colonies, thereby reducing the likelihood that parasites or pathogens can decimate the worker force by overcoming all of its physiological and behavioral defenses at once. A balanced portfolio of investments in genetic variation, in other words, is more likely to produce the highest long-term probability of survival and successful growth. This argument is logical, but in our opinion does not accord with the remarkable correlations that exist between polyandry, sperm count, and colony size. This latter relation favors the limited-sperm hypothesis but no other.

In any case, the reproductive behavior of ants is a still poorly explored domain with rich possibilities for general evolutionary biology. More studies are needed along all fronts, including the comparative natural history of nuptial flights, the detailed analysis of individual males and females during mating, genetic studies of sex determination, and more sophisticated models of reproductive competition at the individual and colony levels.

Flight Month Data

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Taxon Subfamily Month Notes
Acromyrmex versicolor Myrmicinae Jul Aug
Acropyga sauteri Formicinae Mar Apr May Jun
Aenictus lifuiae Dorylinae Aug Taiwan
Amblyopone australis Amblyoponinae Feb Mar
Aneuretus simoni Aneuretinae Jul Aug
Anoplolepis gracilipes Formicinae Apr
Aphaenogaster fulva Myrmicinae Aug Sep Oct
Aphaenogaster lamellidens Myrmicinae May Jun
Aphaenogaster occidentalis Myrmicinae Jun Jul Aug Sep
Aphaenogaster picea Myrmicinae May
Aphaenogaster rudis Myrmicinae Jul Aug
Aphaenogaster subterranea Myrmicinae Jul Aug Sep
Aphaenogaster tennesseensis Myrmicinae Jun
Aphaenogaster texana Myrmicinae Jun
Atta mexicana Myrmicinae Jun Jul
Atta texana Myrmicinae Apr May Jun
Azteca instabilis Dolichoderinae Jan Feb Mar Apr May Jun Jul Aug Sep Oct Nov Dec
Brachymyrmex depilis Formicinae Mar Apr
Brachymyrmex obscurior Formicinae Jun
Brachymyrmex patagonicus Formicinae Apr May Jun Jul Aug Sep Oct
Brachyponera chinensis Ponerinae May Jun Jul
Camponotus aethiops Formicinae Jun Jul Aug
Camponotus americanus Formicinae Apr May Jun
Camponotus anthrax Formicinae May
Camponotus atriceps Formicinae May Jun Jul
Camponotus auriventris Formicinae Apr
Camponotus caryae Formicinae Apr May
Camponotus castaneus Formicinae Mar Apr May Jun Jul
Camponotus chromaiodes Formicinae Mar Apr May Jun Jul
Camponotus cingulatus Formicinae May Jun Nov
Camponotus claviscapus Formicinae May Jun Jul Aug Sep
Camponotus consobrinus Formicinae Dec
Camponotus cruentatus Formicinae May Jun
Camponotus curviscapus Formicinae May Jun Jul
Camponotus dalmaticus Formicinae Apr May
Camponotus decipiens Formicinae May
Camponotus essigi Formicinae May
Camponotus fallax Formicinae May Jun
Camponotus floridanus Formicinae Apr May Jun Jul Aug Sep
Camponotus fragilis Formicinae Jul Aug
Camponotus herculeanus Formicinae May Jun Jul
Camponotus hyatti Formicinae Mar Apr May
Camponotus inaequalis Formicinae Mar as ''Camponotus tortuganus'' (March)
Camponotus japonicus Formicinae May Jun Japan
Camponotus laevigatus Formicinae Mar
Camponotus laevissimus Formicinae Apr May
Camponotus lateralis Formicinae Apr
Camponotus ligniperda Formicinae May Jun
Camponotus modoc Formicinae Apr May Jun Jul Washington (May)
Camponotus mucronatus Formicinae Jun Jul
Camponotus nawai Formicinae Aug Japan
Camponotus nearcticus Formicinae Apr May Jun
Camponotus nicobarensis Formicinae Apr
Camponotus novaeboracensis Formicinae Apr May Jun Jul Washington (May)
Camponotus novogranadensis Formicinae Jun Jul Aug Sep Oct Nov Dec
Camponotus obscuripes Formicinae May Jun Jul Aug in Japan, May to June in lowlands, and until August in the mountains
Camponotus ocreatus Formicinae Mar Apr
Camponotus pennsylvanicus Formicinae Apr May Jun Jul
Camponotus piceus Formicinae May Jun Jul
Camponotus planatus Formicinae May Jun
Camponotus sanctaefidei Formicinae May Jun Jul Aug Sep Oct Nov
Camponotus sansabeanus Formicinae Mar Apr May
Camponotus saxatilis Formicinae Jul Aug
Camponotus semitestaceus Formicinae Mar
Camponotus sexguttatus Formicinae Apr
Camponotus socius Formicinae May
Camponotus subbarbatus Formicinae Apr May Jun
Camponotus substitutus Formicinae Oct Nov
Camponotus suffusus Formicinae Mar
Camponotus texanus Formicinae Mar
Camponotus turkestanus Formicinae Apr May Jun
Camponotus vagus Formicinae Apr May Jun
Camponotus vicinus Formicinae Mar Apr May Jun Jul
Camponotus yamaokai Formicinae May Japan
Camponotus yogi Formicinae Sep
Cardiocondyla mauritanica Myrmicinae Jun
Carebara diversa Myrmicinae Apr
Cataglyphis cursor Formicinae Jul Aug Sep
Cataglyphis nodus Formicinae Jun Jul
Colobopsis impressa Formicinae Apr May Jun Jul
Colobopsis mississippiensis Formicinae Jun Jul
Colobopsis nipponica Formicinae Jun Jul Japan
Colobopsis obliqua Formicinae May Jun
Colobopsis truncata Formicinae Jun Jul Aug
Crematogaster ashmeadi Myrmicinae Aug Sep
Crematogaster cerasi Myrmicinae Aug Sep Oct
Crematogaster depilis Myrmicinae Jun
Crematogaster lineolata Myrmicinae Oct
Crematogaster matsumurai Myrmicinae Jul Aug Sep Japan
Crematogaster mutans Myrmicinae Apr
Crematogaster osakensis Myrmicinae Sep Japan
Crematogaster scutellaris Myrmicinae Aug Sep Oct
Crematogaster stollii Myrmicinae Jun Jul Aug Sep Oct
Cryptopone ochracea Ponerinae Aug Sep
Cryptopone testacea Ponerinae Jan Apr May Jun Aug Dec
Cyphomyrmex costatus Myrmicinae Feb Mar Apr Sep Dec
Cyphomyrmex minutus Myrmicinae May
Cyphomyrmex rimosus Myrmicinae Apr May Jun Jul
Diacamma rugosum Ponerinae Jan Feb Mar Apr
Dinomyrmex gigas Formicinae Jan Feb Mar Apr May Jun Jul Aug Sep Oct Nov Dec
Dolichoderus bispinosus Dolichoderinae Feb Mar Apr May Jun Jul Aug Sep Oct Nov
Dolichoderus debilis Dolichoderinae Jan Feb Mar Apr May Jun Jul Aug Sep Oct Nov Dec
Dolichoderus lutosus Dolichoderinae Jan Feb Mar Apr May Jun Jul Aug Sep Oct Nov Dec
Dolichoderus plagiatus Dolichoderinae May
Dolichoderus quadripunctatus Dolichoderinae Jul Aug Sep
Dolichoderus sibiricus Dolichoderinae Aug Japan
Dolichoderus taschenbergi Dolichoderinae Jun
Dolopomyrmex pilatus Myrmicinae Mar
Dorymyrmex bicolor Dolichoderinae Feb Mar Apr
Dorymyrmex bureni Dolichoderinae Apr May Jun Jul Aug
Dorymyrmex insanus Dolichoderinae Jan Feb Mar Apr May Jul Aug Sep Oct Nov Dec
Ectatomma ruidum Ectatomminae Jan Mar Apr May Jun Jul Aug Sep Oct Nov Dec
Ectatomma tuberculatum Ectatomminae May Jun Jul Aug Sep Oct Nov Dec
Forelius pruinosus Dolichoderinae May Jun Jul Aug
Formica altipetens Formicinae Jul Aug
Formica aquilonia Formicinae Jun Jul
Formica argentea Formicinae Jul Aug
Formica aserva Formicinae Jun Jul
Formica biophilica Formicinae Jun Jul
Formica bruni Formicinae Jul Aug
Formica brunneonitida Formicinae Jun Jul Aug
Formica caucasicola Formicinae Jul Aug
Formica cinerea Formicinae Jun Jul Aug
Formica clara Formicinae Jun Jul
Formica cunicularia Formicinae Jun Jul
Formica exsecta Formicinae Jun Jul Aug
Formica exsectoides Formicinae Jul
Formica foreli Formicinae Jul Aug
Formica forsslundi Formicinae Jul
Formica francoeuri Formicinae Apr May Jun
Formica fusca Formicinae Jun Jul Aug
Formica fuscocinerea Formicinae Jun Jul Aug Sep
Formica gagates Formicinae Jul Aug
Formica gagatoides Formicinae Jul Aug
Formica glacialis Formicinae Jul Aug
Formica glauca Formicinae Jun Jul
Formica incerta Formicinae Jul Aug
Formica integra Formicinae May
Formica integroides Formicinae Apr
Formica japonica Formicinae Jun Jul Aug Sep early June to mid July in Tokyo, Japan (Kawai, 1942) and from August to September in Kochi, Japan (Okamoto, 1957)
Formica lemani Formicinae Jun Jul Aug Sep
Formica longiceps Formicinae Jul Aug
Formica lugubris Formicinae May Jun Jul
Formica manchu Formicinae Jun Jul Aug
Formica mesasiatica Formicinae Jun Jul Aug
Formica moki Formicinae Jun Jul
Formica neogagates Formicinae Aug
Formica obscuripes Formicinae Apr May Washington (May)
Formica obscuriventris Formicinae Jun
Formica pacifica Formicinae Apr
Formica paralugubris Formicinae May Jun Jul
Formica pisarskii Formicinae Aug
Formica podzolica Formicinae Jun Jul Aug
Formica polyctena Formicinae Apr May Jun
Formica pratensis Formicinae Apr May Jun Jul Aug Sep
Formica pressilabris Formicinae Jun Jul Aug
Formica ravida Formicinae May Jun Jul
Formica rubicunda Formicinae Jul
Formica rufa Formicinae May Jun
Formica rufibarbis Formicinae Jun Jul Aug
Formica sanguinea Formicinae May Jun Jul Aug sep Japan
Formica selysi Formicinae Jun Jul Aug Sep
Formica subaenescens Formicinae Aug
Formica subpolita Formicinae Jun
Formica subsericea Formicinae Apr May Jun Jul Aug
Formica transkaucasica Formicinae Jul Aug
Formica transmontanis Formicinae Oct Nov
Formica truncorum Formicinae Jun Jul Aug
Formica ulkei Formicinae Jul Aug
Formica uralensis Formicinae Jun Jul Aug
Formica yessensis Formicinae Aug Japan
Formicoxenus nitidulus Myrmicinae Jul Aug
Gnamptogenys continua Ectatomminae Jan Feb Mar Apr May Jun Jul Aug Sep Oct Nov Dec
Gnamptogenys hartmani Ectatomminae Jan Mar Apr May Jun Jul Aug Sep Oct
Harpagoxenus sublaevis Myrmicinae Jul Aug
Hypoponera nippona Ponerinae Aug Japan
Hypoponera opacior Ponerinae Jul
Hypoponera punctatissima Ponerinae Jun Jul Aug Sep
Iridomyrmex bicknelli Dolichoderinae Apr Yass, New South Wales
Iridomyrmex purpureus Dolichoderinae Oct
Lasius alienus Formicinae Jul Aug Sep
Lasius austriacus Formicinae Jul Aug Sep
Lasius bicornis Formicinae Apr May Jun Jul Aug Sep
Lasius brevicornis Formicinae Aug Sep
Lasius brunneus Formicinae May Jun Jul Aug
Lasius capitatus Formicinae Sep Japan
Lasius carniolicus Formicinae May Jun Jul Aug Sep Oct
Lasius citrinus Formicinae Apr May Jun Jul Aug
Lasius claviger Formicinae Aug Sep Oct Nov Dec
Lasius distinguendus Formicinae Jul Aug Sep
Lasius emarginatus Formicinae Jun Jul Aug
Lasius flavus Formicinae Jun Jul Aug Sep
Lasius fuliginosus Formicinae Jun Jul Aug Sep
Lasius hayashi Formicinae Jul Aug Japan
Lasius interjectus Formicinae Apr May Jun
Lasius japonicus Formicinae Jul Aug Japan
Lasius jensi Formicinae Jun Jul Aug Sep
Lasius lasioides Formicinae May Jun Jul
Lasius latipes Formicinae Jun Jul Aug Sep
Lasius meridionalis Formicinae Jun Jul Aug Sep
Lasius mixtus Formicinae Jul Aug Sep
Lasius morisitai Formicinae Jul Japan
Lasius murphyi Formicinae Mar
Lasius myops Formicinae May Jun Jul Aug Sep
Lasius nearcticus Formicinae Jul Aug Sep
Lasius neoniger Formicinae Jul Aug Sep Oct Nov Dec
Lasius niger Formicinae Jun Jul Aug Sep
Lasius nipponensis Formicinae Jun Jul Japan
Lasius orientalis Formicinae Jul Japan
Lasius paralienus Formicinae Aug Sep Oct
Lasius platythorax Formicinae Jun Jul Aug
Lasius productus Formicinae Aug Sep Japan
Lasius psammophilus Formicinae Jul Aug
Lasius reginae Formicinae Aug Sep
Lasius sabularum Formicinae Aug Sep Oct
Lasius sakagamii Formicinae Jun Jul Aug Sep Oct Japan
Lasius sonobei Formicinae Aug Sep Japan
Lasius spathepus Formicinae Jun Jul Aug Japan
Lasius subumbratus Formicinae Jul Aug
Lasius talpa Formicinae Aug Sep Japan
Lasius umbratus Formicinae Jun Jul Aug Sep
Leptogenys elongata Ponerinae May Jun
Leptogenys punctaticeps Ponerinae Jan Feb Mar Apr May Jun Jul Aug Sep Oct Nov Dec
Leptothorax acervorum Myrmicinae Jun Jul Aug Sep
Leptothorax gredleri Myrmicinae Jul Aug
Leptothorax kutteri Myrmicinae Jul Aug
Leptothorax muscorum Myrmicinae Jun Jul Aug Sep
Leptothorax pacis Myrmicinae Jul
Linepithema humile Dolichoderinae May Jun Jul
Liometopum luctuosum Dolichoderinae Mar
Liometopum microcephalum Dolichoderinae Jun Jul
Liometopum occidentale Dolichoderinae Mar Apr May Jun
Manica rubida Myrmicinae May Jun Jul Aug Sep
Manica yessensis Myrmicinae Aug Japan
Mayaponera arhuaca Ponerinae Jan Feb Mar Apr May Jun Jul Aug Sep
Mayaponera constricta Ponerinae Jan Feb Mar Apr May Jun Jul Aug Sep Oct Dec
Megalomyrmex symmetochus Myrmicinae May Jun Jul Aug
Meranoplus bicolor Myrmicinae Jan Feb Apr May Jun Aug Sep Oct Nov Dec
Messor aciculatus Myrmicinae Apr May Japan
Messor barbarus Myrmicinae Sep Oct Nov
Messor capitatus Myrmicinae Jan Nov Dec
Messor ebeninus Myrmicinae Nov Dec
Messor structor Myrmicinae Mar Apr May Jun Jul Aug Sep
Monomorium ergatogyna Myrmicinae Jun
Monomorium floricola Myrmicinae Jun
Monomorium minimum Myrmicinae Mar
Myrmecia brevinoda Myrmeciinae Mar
Myrmecia tarsata Myrmeciinae Nov
Myrmecina americana Myrmicinae Aug Sep Oct
Myrmecina graminicola Myrmicinae Aug Sep
Myrmecocystus creightoni Formicinae Feb
Myrmecocystus kennedyi Formicinae Mar Apr
Myrmecocystus mendax Formicinae Jul
Myrmecocystus mexicanus Formicinae Jul Aug
Myrmecocystus mimicus Formicinae Jun Jul Aug
Myrmecocystus navajo Formicinae Jul Aug
Myrmecocystus romainei Formicinae Jul
Myrmecocystus semirufus Formicinae Mar
Myrmecocystus testaceus Formicinae Mar Apr May
Myrmecocystus yuma Formicinae Jul Aug
Myrmica aloba Myrmicinae Jul Aug Sep
Myrmica bibikoffi Myrmicinae Aug
Myrmica constricta Myrmicinae Aug Sep
Myrmica deplanata Myrmicinae Jul Aug Sep
Myrmica forcipata Myrmicinae Jul Aug
Myrmica gallienii Myrmicinae Aug Sep Oct
Myrmica hellenica Myrmicinae Aug Sep Oct
Myrmica hirsuta Myrmicinae Aug Sep
Myrmica jessensis Myrmicinae Sep Japan
Myrmica karavajevi Myrmicinae Jul Aug Sep
Myrmica kotokui Myrmicinae Aug Sep Oct Japan
Myrmica kurokii Myrmicinae Aug
Myrmica lobicornis Myrmicinae Jul Aug Sep
Myrmica lobulicornis Myrmicinae Jul Aug Sep
Myrmica lonae Myrmicinae Jul Aug Sep
Myrmica pinetorum Myrmicinae May
Myrmica pisarskii Myrmicinae Jul Aug Sep
Myrmica punctiventris Myrmicinae Sep Oct Nov
Myrmica rubra Myrmicinae Aug
Myrmica ruginodis Myrmicinae Jul Aug
Myrmica rugulosa Myrmicinae Aug Sep Oct
Myrmica sabuleti Myrmicinae Jul Aug Sep
Myrmica salina Myrmicinae Aug
Myrmica scabrinodis Myrmicinae Jul Aug Sep
Myrmica schencki Myrmicinae Jul Aug
Myrmica specioides Myrmicinae Jul Aug Sep
Myrmica sulcinodis Myrmicinae Jul Aug Sep
Myrmica vandeli Myrmicinae Jul Aug Sep
Myrmica wesmaeli Myrmicinae Aug Sep
Myrmicaria natalensis Myrmicinae Jan
Myrmicocrypta dilacerata Myrmicinae Jan Feb Mar Apr May Jun Dec
Neivamyrmex klugii distans Dorylinae Mar Apr May Jun
Neivamyrmex melshaemeri Dorylinae Apr May Jun Jul Aug
Neivamyrmex nigrescens Dorylinae Jul
Neivamyrmex pilosus Dorylinae Mar Apr May late dry season to early wet season
Neivamyrmex pseudops Dorylinae Jan Feb Mar Apr May Jun Jul Aug Sep Oct Dec
Neivamyrmex swainsonii Dorylinae Mar Apr May
Novomessor cockerelli Myrmicinae Jul
Nylanderia faisonensis Formicinae Apr May
Nylanderia flavipes Formicinae May Jun Japan
Nylanderia parvula Formicinae May Jun
Nylanderia terricola Formicinae Feb
Nylanderia vaga Formicinae Oct
Nylanderia vividula Formicinae Mar Apr
Odontomachus bauri Ponerinae Jan Feb Mar Apr May Jun Jul Aug Sep Oct Nov Dec
Odontomachus clarus Ponerinae Jun Jul
Odontomachus simillimus Ponerinae Feb
Pachycondyla harpax Ponerinae Jan Feb Mar Apr May Jun Jul Nov Dec
Paraparatrechina sakurae Formicinae Oct Nov Japan
Paraponera clavata Paraponerinae Jan Feb Mar Apr May Jun Jul Aug Sep Oct Nov Dec
Paratrechina longicornis Formicinae Jun Jul
Pheidole bicarinata Myrmicinae Jun Jul Aug
Pheidole californica Myrmicinae Apr May
Pheidole ceres Myrmicinae Jul
Pheidole christopherseni Myrmicinae Jan Feb Mar Nov Dec
Pheidole dentata Myrmicinae Apr May Jun
Pheidole dentigula Myrmicinae May
Pheidole desertorum Myrmicinae Jun Jul Aug
Pheidole diversipilosa Myrmicinae Jun
Pheidole flavens Myrmicinae May
Pheidole floridana Myrmicinae Apr May
Pheidole gilvescens Myrmicinae Aug
Pheidole hyatti Myrmicinae May Jun Jul
Pheidole megacephala Myrmicinae Aug
Pheidole moerens Myrmicinae May Jun Jul
Pheidole morrisii Myrmicinae May Jun
Pheidole navigans Myrmicinae Jul Aug
Pheidole obscurithorax Myrmicinae May Jun Jul
Pheidole obtusospinosa Myrmicinae Jul Aug Sep
Pheidole pallidula Myrmicinae May Jun Jul Aug
Pheidole pilifera Myrmicinae Jun Jul Aug
Pheidole tysoni Myrmicinae Aug
Pheidole vistana Myrmicinae Jun Jul
Pheidole xerophila Myrmicinae Jul Aug
Plagiolepis pallescens Formicinae Jun Jul Aug
Plagiolepis xene Formicinae Jul Aug
Pogonomyrmex badius Myrmicinae May Jun Jul
Pogonomyrmex barbatus Myrmicinae Apr May Jun Jul Aug Sep
Pogonomyrmex californicus Myrmicinae Apr May Jun Jul Aug
Pogonomyrmex magnacanthus Myrmicinae Apr May
Pogonomyrmex maricopa Myrmicinae Jul Aug
Pogonomyrmex montanus Myrmicinae Jul
Pogonomyrmex occidentalis Myrmicinae Aug Sep
Pogonomyrmex rugosus Myrmicinae Jul Aug Sep
Pogonomyrmex subdentatus Myrmicinae Mar Apr May Jun
Pogonomyrmex subnitidus Myrmicinae Jun
Pogonomyrmex tenuispinus Myrmicinae Jul
Polyergus breviceps Formicinae Aug Sep
Polyergus lucidus Formicinae Jul Aug Sep
Polyergus mexicanus Formicinae Aug
Polyergus rufescens Formicinae Jul Aug Sep
Polyergus topoffi Formicinae Jul
Polyergus vinosus Formicinae May
Polyrhachis lamellidens Formicinae Sep Oct Nov
Ponera coarctata Ponerinae Aug Sep
Ponera kohmoku Ponerinae Aug Japan
Ponera pennsylvanica Ponerinae Mar Apr May
Poneracantha triangularis Ectatomminae Jul
Prenolepis imparis Formicinae Jan Feb Mar Apr May Jun Dec
Prenolepis nitens Formicinae Apr May
Proatta butteli Myrmicinae Apr
Proceratium chickasaw Proceratiinae Aug
Proceratium silaceum Proceratiinae Jan
Procryptocerus belti Myrmicinae May Jun Jul Aug Sep Nov Dec
Pseudomyrmex apache Pseudomyrmecinae Apr
Pseudomyrmex caeciliae Pseudomyrmecinae May
Pseudomyrmex ejectus Pseudomyrmecinae May Jun Jul
Pseudomyrmex gracilis Pseudomyrmecinae Mar Apr May Jun Jul Aug Sep Oct Nov
Solenopsis amblychila Myrmicinae Apr
Solenopsis carolinensis Myrmicinae May Jun
Solenopsis fugax Myrmicinae Aug Sep
Solenopsis geminata Myrmicinae Apr May
Solenopsis invicta Myrmicinae Apr May Jun Jul
Solenopsis krockowi Myrmicinae Jul Aug
Solenopsis molesta Myrmicinae Jun Jul Aug
Solenopsis pergandei Myrmicinae May Jun
Solenopsis tennesseensis Myrmicinae Jul
Solenopsis texana Myrmicinae Jul
Solenopsis xyloni Myrmicinae Apr May Jun Jul Aug
Stenamma debile Myrmicinae Aug Sep Oct
Stenamma nipponense Myrmicinae Sep Japan
Stigmatomma pallipes Amblyoponinae Aug Sep
Strongylognathus alpinus Myrmicinae Jul Aug Sep Oct
Strongylognathus bulgaricus Myrmicinae Jul Aug Sep
Strongylognathus huberi Myrmicinae Jul Aug Sep
Strongylognathus italicus Myrmicinae Aug Sep
Strongylognathus kratochvili Myrmicinae Jul Aug Sep
Strongylognathus testaceus Myrmicinae Jun Jul Aug
Strumigenys dolichognatha Myrmicinae Mar Apr May Jun Jul Aug Sep Oct Nov
Strumigenys elongata Myrmicinae Feb Mar Apr May Jun Jul Aug Sep
Strumigenys gundlachi Myrmicinae Apr May Jun Jul Aug Sep
Strumigenys lewisi Myrmicinae Aug Japan
Strumigenys margaritae Myrmicinae Jul Aug
Strumigenys membranifera Myrmicinae Feb Mar Apr May Jun Jul Aug Sep Oct Nov
Strumigenys minutula Myrmicinae Jun Jul Hong Kong
Strumigenys zeteki Myrmicinae Jan Feb Mar Apr May Jun Jul Aug
Syscia augustae Dorylinae Jun Jul Aug
Tapinoma erraticum Dolichoderinae May Jun Jul
Tapinoma madeirense Dolichoderinae Jun Jul
Tapinoma magnum Dolichoderinae Apr May Jun Jul Aug Sep April-June in Italy, Germany, and the Netherlands; August-September in Algeria
Tapinoma minutum Dolichoderinae Mar
Tapinoma sessile Dolichoderinae Apr May Jun Jul
Technomyrmex albipes Dolichoderinae Jul
Technomyrmex gibbosus Dolichoderinae Sep Japan
Temnothorax affinis Myrmicinae Jul Aug
Temnothorax albipennis Myrmicinae Jul
Temnothorax andrei Myrmicinae Jun Jul
Temnothorax caguatan Myrmicinae Jun Jul
Temnothorax congruus Myrmicinae Jul Japan
Temnothorax corticalis Myrmicinae Jul Aug Sep
Temnothorax crassispinus Myrmicinae Jul Aug Sep
Temnothorax curvispinosus Myrmicinae Jun Jul Aug
Temnothorax interruptus Myrmicinae Jun Jul Aug Sep
Temnothorax nigriceps Myrmicinae Jun Jul Aug Sep
Temnothorax nylanderi Myrmicinae Jul Aug Sep
Temnothorax parvulus Myrmicinae Aug Sep
Temnothorax ravouxi Myrmicinae Jun Jul Aug
Temnothorax saxonicus Myrmicinae Jul
Temnothorax sordidulus Myrmicinae Jul
Temnothorax spinosior Myrmicinae Jul Japan
Temnothorax tuberum Myrmicinae Jun Jul Aug
Temnothorax unifasciatus Myrmicinae Jun Jul Aug
Temnothorax vivianoi Myrmicinae Aug Sicily, Italy
Tetramorium atratulum Myrmicinae May Jun Jul Aug Sep
Tetramorium bicarinatum Myrmicinae May Jun
Tetramorium caespitum Myrmicinae Jun Jul Aug
Tetramorium hungaricum Myrmicinae Jun
Tetramorium immigrans Myrmicinae Jul
Tetramorium impurum Myrmicinae Jul Aug Sep Oct
Tetramorium moravicum Myrmicinae May Jun
Tetramorium semilaeve Myrmicinae Jul Aug
Tetramorium tsushimae Myrmicinae May Jun Jul
Trachymyrmex septentrionalis Myrmicinae May Jun Jul Aug
Typhlomyrmex rogenhoferi Ectatomminae Mar Apr May Jun Jul Aug
Veromessor andrei Myrmicinae Jun Jul
Veromessor julianus Myrmicinae Feb Mar
Veromessor pergandei Myrmicinae Feb Mar Apr
Wadeura guianensis Ponerinae Jul Aug Sep Oct Nov
Wasmannia auropunctata Myrmicinae Apr May Jun Jul Aug Sep Oct Nov
Xenomyrmex stollii Myrmicinae Jan Feb Mar Apr May Jun Jul Aug Sep Oct Nov Dec

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Month Taxa
January Azteca instabilis, Cryptopone testacea, Diacamma rugosum, Dinomyrmex gigas, Dolichoderus debilis, Dolichoderus lutosus, Dorymyrmex insanus, Ectatomma ruidum, Gnamptogenys continua, Gnamptogenys hartmani, Leptogenys punctaticeps, Mayaponera arhuaca, Mayaponera constricta, Meranoplus bicolor, Messor capitatus, Myrmicaria natalensis, Myrmicocrypta dilacerata, Neivamyrmex pseudops, Odontomachus bauri, Pachycondyla harpax, Paraponera clavata, Pheidole christopherseni, Prenolepis imparis, Proceratium silaceum, Strumigenys zeteki, Xenomyrmex stollii
February Amblyopone australis, Azteca instabilis, Cyphomyrmex costatus, Diacamma rugosum, Dinomyrmex gigas, Dolichoderus bispinosus, Dolichoderus debilis, Dolichoderus lutosus, Dorymyrmex bicolor, Dorymyrmex insanus, Gnamptogenys continua, Leptogenys punctaticeps, Mayaponera arhuaca, Mayaponera constricta, Meranoplus bicolor, Myrmecocystus creightoni, Myrmicocrypta dilacerata, Neivamyrmex pseudops, Nylanderia terricola, Odontomachus bauri, Odontomachus simillimus, Pachycondyla harpax, Paraponera clavata, Pheidole christopherseni, Prenolepis imparis, Strumigenys elongata, Strumigenys membranifera, Strumigenys zeteki, Veromessor julianus, Veromessor pergandei, Xenomyrmex stollii
March Acropyga sauteri, Amblyopone australis, Azteca instabilis, Brachymyrmex depilis, Camponotus castaneus, Camponotus chromaiodes, Camponotus hyatti, Camponotus inaequalis, Camponotus laevigatus, Camponotus ocreatus, Camponotus sansabeanus, Camponotus semitestaceus, Camponotus suffusus, Camponotus texanus, Camponotus vicinus, Cyphomyrmex costatus, Diacamma rugosum, Dinomyrmex gigas, Dolichoderus bispinosus, Dolichoderus debilis, Dolichoderus lutosus, Dolopomyrmex pilatus, Dorymyrmex bicolor, Dorymyrmex insanus, Ectatomma ruidum, Gnamptogenys continua, Gnamptogenys hartmani, Lasius murphyi, Leptogenys punctaticeps, Liometopum luctuosum, Liometopum occidentale, Mayaponera arhuaca, Mayaponera constricta, Messor structor, Monomorium minimum, Myrmecia brevinoda, Myrmecocystus kennedyi, Myrmecocystus semirufus, Myrmecocystus testaceus, Myrmicocrypta dilacerata, Neivamyrmex klugii distans, Neivamyrmex pilosus, Neivamyrmex pseudops, Neivamyrmex swainsonii, Nylanderia vividula, Odontomachus bauri, Pachycondyla harpax, Paraponera clavata, Pheidole christopherseni, Pogonomyrmex subdentatus, Ponera pennsylvanica, Prenolepis imparis, Pseudomyrmex gracilis, Strumigenys dolichognatha, Strumigenys elongata, Strumigenys membranifera, Strumigenys zeteki, Tapinoma minutum, Typhlomyrmex rogenhoferi, Veromessor julianus, Veromessor pergandei, Xenomyrmex stollii
April Acropyga sauteri, Anoplolepis gracilipes, Atta texana, Azteca instabilis, Brachymyrmex depilis, Brachymyrmex patagonicus, Camponotus americanus, Camponotus auriventris, Camponotus caryae, Camponotus castaneus, Camponotus chromaiodes, Camponotus dalmaticus, Camponotus floridanus, Camponotus hyatti, Camponotus laevissimus, Camponotus lateralis, Camponotus modoc, Camponotus nearcticus, Camponotus nicobarensis, Camponotus novaeboracensis, Camponotus ocreatus, Camponotus pennsylvanicus, Camponotus sansabeanus, Camponotus sexguttatus, Camponotus subbarbatus, Camponotus turkestanus, Camponotus vagus, Camponotus vicinus, Carebara diversa, Colobopsis impressa, Crematogaster mutans, Cryptopone testacea, Cyphomyrmex costatus, Cyphomyrmex rimosus, Diacamma rugosum, Dinomyrmex gigas, Dolichoderus bispinosus, Dolichoderus debilis, Dolichoderus lutosus, Dorymyrmex bicolor, Dorymyrmex bureni, Dorymyrmex insanus, Ectatomma ruidum, Formica francoeuri, Formica integroides, Formica obscuripes, Formica pacifica, Formica polyctena, Formica pratensis, Formica subsericea, Gnamptogenys continua, Gnamptogenys hartmani, Iridomyrmex bicknelli, Lasius bicornis, Lasius citrinus, Lasius interjectus, Leptogenys punctaticeps, Liometopum occidentale, Mayaponera arhuaca, Mayaponera constricta, Meranoplus bicolor, Messor aciculatus, Messor structor, Myrmecocystus kennedyi, Myrmecocystus testaceus, Myrmicocrypta dilacerata, Neivamyrmex klugii distans, Neivamyrmex melshaemeri, Neivamyrmex pilosus, Neivamyrmex pseudops, Neivamyrmex swainsonii, Nylanderia faisonensis, Nylanderia vividula, Odontomachus bauri, Pachycondyla harpax, Paraponera clavata, Pheidole californica, Pheidole dentata, Pheidole floridana, Pogonomyrmex barbatus, Pogonomyrmex californicus, Pogonomyrmex magnacanthus, Pogonomyrmex subdentatus, Ponera pennsylvanica, Prenolepis imparis, Prenolepis nitens, Proatta butteli, Pseudomyrmex apache, Pseudomyrmex gracilis, Solenopsis amblychila, Solenopsis geminata, Solenopsis invicta, Solenopsis xyloni, Strumigenys dolichognatha, Strumigenys elongata, Strumigenys gundlachi, Strumigenys membranifera, Strumigenys zeteki, Tapinoma magnum, Tapinoma sessile, Typhlomyrmex rogenhoferi, Veromessor pergandei, Wasmannia auropunctata, Xenomyrmex stollii
May Acropyga sauteri, Aphaenogaster lamellidens, Aphaenogaster picea, Atta texana, Azteca instabilis, Brachymyrmex patagonicus, Brachyponera chinensis, Camponotus americanus, Camponotus anthrax, Camponotus atriceps, Camponotus caryae, Camponotus castaneus, Camponotus chromaiodes, Camponotus cingulatus, Camponotus claviscapus, Camponotus cruentatus, Camponotus curviscapus, Camponotus dalmaticus, Camponotus decipiens, Camponotus essigi, Camponotus fallax, Camponotus floridanus, Camponotus herculeanus, Camponotus hyatti, Camponotus japonicus, Camponotus laevissimus, Camponotus ligniperda, Camponotus modoc, Camponotus nearcticus, Camponotus novaeboracensis, Camponotus obscuripes, Camponotus pennsylvanicus, Camponotus piceus, Camponotus planatus, Camponotus sanctaefidei, Camponotus sansabeanus, Camponotus socius, Camponotus subbarbatus, Camponotus turkestanus, Camponotus vagus, Camponotus vicinus, Camponotus yamaokai, Colobopsis impressa, Colobopsis obliqua, Cryptopone testacea, Cyphomyrmex minutus, Cyphomyrmex rimosus, Dinomyrmex gigas, Dolichoderus bispinosus, Dolichoderus debilis, Dolichoderus lutosus, Dolichoderus plagiatus, Dorymyrmex bureni, Dorymyrmex insanus, Ectatomma ruidum, Ectatomma tuberculatum, Forelius pruinosus, Formica francoeuri, Formica integra, Formica lugubris, Formica obscuripes, Formica paralugubris, Formica polyctena, Formica pratensis, Formica ravida, Formica rufa, Formica sanguinea, Formica subsericea, Gnamptogenys continua, Gnamptogenys hartmani, Lasius bicornis, Lasius brunneus, Lasius carniolicus, Lasius citrinus, Lasius interjectus, Lasius lasioides, Lasius myops, Leptogenys elongata, Leptogenys punctaticeps, Linepithema humile, Liometopum occidentale, Manica rubida, Mayaponera arhuaca, Mayaponera constricta, Megalomyrmex symmetochus, Meranoplus bicolor, Messor aciculatus, Messor structor, Myrmecocystus testaceus, Myrmica pinetorum, Myrmicocrypta dilacerata, Neivamyrmex klugii distans, Neivamyrmex melshaemeri, Neivamyrmex pilosus, Neivamyrmex pseudops, Neivamyrmex swainsonii, Nylanderia faisonensis, Nylanderia flavipes, Nylanderia parvula, Odontomachus bauri, Pachycondyla harpax, Paraponera clavata, Pheidole californica, Pheidole dentata, Pheidole dentigula, Pheidole flavens, Pheidole floridana, Pheidole hyatti, Pheidole moerens, Pheidole morrisii, Pheidole obscurithorax, Pheidole pallidula, Pogonomyrmex badius, Pogonomyrmex barbatus, Pogonomyrmex californicus, Pogonomyrmex magnacanthus, Pogonomyrmex subdentatus, Polyergus vinosus, Ponera pennsylvanica, Prenolepis imparis, Prenolepis nitens, Procryptocerus belti, Pseudomyrmex caeciliae, Pseudomyrmex ejectus, Pseudomyrmex gracilis, Solenopsis carolinensis, Solenopsis geminata, Solenopsis invicta, Solenopsis pergandei, Solenopsis xyloni, Strumigenys dolichognatha, Strumigenys elongata, Strumigenys gundlachi, Strumigenys membranifera, Strumigenys zeteki, Tapinoma erraticum, Tapinoma magnum, Tapinoma sessile, Tetramorium atratulum, Tetramorium bicarinatum, Tetramorium moravicum, Tetramorium tsushimae, Trachymyrmex septentrionalis, Typhlomyrmex rogenhoferi, Wasmannia auropunctata, Xenomyrmex stollii
June Acropyga sauteri, Aphaenogaster lamellidens, Aphaenogaster occidentalis, Aphaenogaster tennesseensis, Aphaenogaster texana, Atta mexicana, Atta texana, Azteca instabilis, Brachymyrmex obscurior, Brachymyrmex patagonicus, Brachyponera chinensis, Camponotus aethiops, Camponotus americanus, Camponotus atriceps, Camponotus castaneus, Camponotus chromaiodes, Camponotus cingulatus, Camponotus claviscapus, Camponotus cruentatus, Camponotus curviscapus, Camponotus fallax, Camponotus floridanus, Camponotus herculeanus, Camponotus japonicus, Camponotus ligniperda, Camponotus modoc, Camponotus mucronatus, Camponotus nearcticus, Camponotus novaeboracensis, Camponotus novogranadensis, Camponotus obscuripes, Camponotus pennsylvanicus, Camponotus piceus, Camponotus planatus, Camponotus sanctaefidei, Camponotus subbarbatus, Camponotus turkestanus, Camponotus vagus, Camponotus vicinus, Cardiocondyla mauritanica, Cataglyphis nodus, Colobopsis impressa, Colobopsis mississippiensis, Colobopsis nipponica, Colobopsis obliqua, Colobopsis truncata, Crematogaster depilis, Crematogaster stollii, Cryptopone testacea, Cyphomyrmex rimosus, Dinomyrmex gigas, Dolichoderus bispinosus, Dolichoderus debilis, Dolichoderus lutosus, Dolichoderus taschenbergi, Dorymyrmex bureni, Ectatomma ruidum, Ectatomma tuberculatum, Forelius pruinosus, Formica aquilonia, Formica aserva, Formica biophilica, Formica brunneonitida, Formica cinerea, Formica clara, Formica cunicularia, Formica exsecta, Formica francoeuri, Formica fusca, Formica fuscocinerea, Formica glauca, Formica japonica, Formica lemani, Formica lugubris, Formica manchu, Formica mesasiatica, Formica moki, Formica obscuriventris, Formica paralugubris, Formica podzolica, Formica polyctena, Formica pratensis, Formica pressilabris, Formica ravida, Formica rufa, Formica rufibarbis, Formica sanguinea, Formica selysi, Formica subpolita, Formica subsericea, Formica truncorum, Formica uralensis, Gnamptogenys continua, Gnamptogenys hartmani, Hypoponera punctatissima, Lasius bicornis, Lasius brunneus, Lasius carniolicus, Lasius citrinus, Lasius emarginatus, Lasius flavus, Lasius fuliginosus, Lasius interjectus, Lasius jensi, Lasius lasioides, Lasius latipes, Lasius meridionalis, Lasius myops, Lasius niger, Lasius nipponensis, Lasius platythorax, Lasius sakagamii, Lasius spathepus, Lasius umbratus, Leptogenys elongata, Leptogenys punctaticeps, Leptothorax acervorum, Leptothorax muscorum, Linepithema humile, Liometopum microcephalum, Liometopum occidentale, Manica rubida, Mayaponera arhuaca, Mayaponera constricta, Megalomyrmex symmetochus, Meranoplus bicolor, Messor structor, Monomorium ergatogyna, Monomorium floricola, Myrmecocystus mimicus, Myrmicocrypta dilacerata, Neivamyrmex klugii distans, Neivamyrmex melshaemeri, Neivamyrmex pseudops, Nylanderia flavipes, Nylanderia parvula, Odontomachus bauri, Odontomachus clarus, Pachycondyla harpax, Paraponera clavata, Paratrechina longicornis, Pheidole bicarinata, Pheidole dentata, Pheidole desertorum, Pheidole diversipilosa, Pheidole hyatti, Pheidole moerens, Pheidole morrisii, Pheidole obscurithorax, Pheidole pallidula, Pheidole pilifera, Pheidole vistana, Plagiolepis pallescens, Pogonomyrmex badius, Pogonomyrmex barbatus, Pogonomyrmex californicus, Pogonomyrmex subdentatus, Pogonomyrmex subnitidus, Prenolepis imparis, Procryptocerus belti, Pseudomyrmex ejectus, Pseudomyrmex gracilis, Solenopsis carolinensis, Solenopsis invicta, Solenopsis molesta, Solenopsis pergandei, Solenopsis xyloni, Strongylognathus testaceus, Strumigenys dolichognatha, Strumigenys elongata, Strumigenys gundlachi, Strumigenys membranifera, Strumigenys minutula, Strumigenys zeteki, Syscia augustae, Tapinoma erraticum, Tapinoma madeirense, Tapinoma magnum, Tapinoma sessile, Temnothorax andrei, Temnothorax caguatan, Temnothorax curvispinosus, Temnothorax interruptus, Temnothorax nigriceps, Temnothorax ravouxi, Temnothorax tuberum, Temnothorax unifasciatus, Tetramorium atratulum, Tetramorium bicarinatum, Tetramorium caespitum, Tetramorium hungaricum, Tetramorium moravicum, Tetramorium tsushimae, Trachymyrmex septentrionalis, Typhlomyrmex rogenhoferi, Veromessor andrei, Wasmannia auropunctata, Xenomyrmex stollii
July Acromyrmex versicolor, Aneuretus simoni, Aphaenogaster occidentalis, Aphaenogaster rudis, Aphaenogaster subterranea, Atta mexicana, Azteca instabilis, Brachymyrmex patagonicus, Brachyponera chinensis, Camponotus aethiops, Camponotus atriceps, Camponotus castaneus, Camponotus chromaiodes, Camponotus claviscapus, Camponotus curviscapus, Camponotus floridanus, Camponotus fragilis, Camponotus herculeanus, Camponotus modoc, Camponotus mucronatus, Camponotus novaeboracensis, Camponotus novogranadensis, Camponotus obscuripes, Camponotus pennsylvanicus, Camponotus piceus, Camponotus sanctaefidei, Camponotus saxatilis, Camponotus vicinus, Cataglyphis cursor, Cataglyphis nodus, Colobopsis impressa, Colobopsis mississippiensis, Colobopsis nipponica, Colobopsis truncata, Crematogaster matsumurai, Crematogaster stollii, Cyphomyrmex rimosus, Dinomyrmex gigas, Dolichoderus bispinosus, Dolichoderus debilis, Dolichoderus lutosus, Dolichoderus quadripunctatus, Dorymyrmex bureni, Dorymyrmex insanus, Ectatomma ruidum, Ectatomma tuberculatum, Forelius pruinosus, Formica altipetens, Formica aquilonia, Formica argentea, Formica aserva, Formica biophilica, Formica bruni, Formica brunneonitida, Formica caucasicola, Formica cinerea, Formica clara, Formica cunicularia, Formica exsecta, Formica exsectoides, Formica foreli, Formica forsslundi, Formica fusca, Formica fuscocinerea, Formica gagates, Formica gagatoides, Formica glacialis, Formica glauca, Formica incerta, Formica japonica, Formica lemani, Formica longiceps, Formica lugubris, Formica manchu, Formica mesasiatica, Formica moki, Formica paralugubris, Formica podzolica, Formica pratensis, Formica pressilabris, Formica ravida, Formica rubicunda, Formica rufibarbis, Formica sanguinea, Formica selysi, Formica subsericea, Formica transkaucasica, Formica truncorum, Formica ulkei, Formica uralensis, Formicoxenus nitidulus, Gnamptogenys continua, Gnamptogenys hartmani, Harpagoxenus sublaevis, Hypoponera opacior, Hypoponera punctatissima, Lasius alienus, Lasius austriacus, Lasius bicornis, Lasius brunneus, Lasius carniolicus, Lasius citrinus, Lasius distinguendus, Lasius emarginatus, Lasius flavus, Lasius fuliginosus, Lasius hayashi, Lasius japonicus, Lasius jensi, Lasius lasioides, Lasius latipes, Lasius meridionalis, Lasius mixtus, Lasius morisitai, Lasius myops, Lasius nearcticus, Lasius neoniger, Lasius niger, Lasius nipponensis, Lasius orientalis, Lasius platythorax, Lasius psammophilus, Lasius sakagamii, Lasius spathepus, Lasius subumbratus, Lasius umbratus, Leptogenys punctaticeps, Leptothorax acervorum, Leptothorax gredleri, Leptothorax kutteri, Leptothorax muscorum, Leptothorax pacis, Linepithema humile, Liometopum microcephalum, Manica rubida, Mayaponera arhuaca, Mayaponera constricta, Megalomyrmex symmetochus, Messor structor, Myrmecocystus mendax, Myrmecocystus mexicanus, Myrmecocystus mimicus, Myrmecocystus navajo, Myrmecocystus romainei, Myrmecocystus yuma, Myrmica aloba, Myrmica deplanata, Myrmica forcipata, Myrmica karavajevi, Myrmica lobicornis, Myrmica lobulicornis, Myrmica lonae, Myrmica pisarskii, Myrmica ruginodis, Myrmica sabuleti, Myrmica scabrinodis, Myrmica schencki, Myrmica specioides, Myrmica sulcinodis, Myrmica vandeli, Neivamyrmex melshaemeri, Neivamyrmex nigrescens, Neivamyrmex pseudops, Novomessor cockerelli, Odontomachus bauri, Odontomachus clarus, Pachycondyla harpax, Paraponera clavata, Paratrechina longicornis, Pheidole bicarinata, Pheidole ceres, Pheidole desertorum, Pheidole hyatti, Pheidole moerens, Pheidole navigans, Pheidole obscurithorax, Pheidole obtusospinosa, Pheidole pallidula, Pheidole pilifera, Pheidole vistana, Pheidole xerophila, Plagiolepis pallescens, Plagiolepis xene, Pogonomyrmex badius, Pogonomyrmex barbatus, Pogonomyrmex californicus, Pogonomyrmex maricopa, Pogonomyrmex montanus, Pogonomyrmex rugosus, Pogonomyrmex tenuispinus, Polyergus lucidus, Polyergus rufescens, Polyergus topoffi, Poneracantha triangularis, Procryptocerus belti, Pseudomyrmex ejectus, Pseudomyrmex gracilis, Solenopsis invicta, Solenopsis krockowi, Solenopsis molesta, Solenopsis tennesseensis, Solenopsis texana, Solenopsis xyloni, Strongylognathus alpinus, Strongylognathus bulgaricus, Strongylognathus huberi, Strongylognathus kratochvili, Strongylognathus testaceus, Strumigenys dolichognatha, Strumigenys elongata, Strumigenys gundlachi, Strumigenys margaritae, Strumigenys membranifera, Strumigenys minutula, Strumigenys zeteki, Syscia augustae, Tapinoma erraticum, Tapinoma madeirense, Tapinoma magnum, Tapinoma sessile, Technomyrmex albipes, Temnothorax affinis, Temnothorax albipennis, Temnothorax andrei, Temnothorax caguatan, Temnothorax congruus, Temnothorax corticalis, Temnothorax crassispinus, Temnothorax curvispinosus, Temnothorax interruptus, Temnothorax nigriceps, Temnothorax nylanderi, Temnothorax ravouxi, Temnothorax saxonicus, Temnothorax sordidulus, Temnothorax spinosior, Temnothorax tuberum, Temnothorax unifasciatus, Tetramorium atratulum, Tetramorium caespitum, Tetramorium immigrans, Tetramorium impurum, Tetramorium semilaeve, Tetramorium tsushimae, Trachymyrmex septentrionalis, Typhlomyrmex rogenhoferi, Veromessor andrei, Wadeura guianensis, Wasmannia auropunctata, Xenomyrmex stollii
August Acromyrmex versicolor, Aenictus lifuiae, Aneuretus simoni, Aphaenogaster fulva, Aphaenogaster occidentalis, Aphaenogaster rudis, Aphaenogaster subterranea, Azteca instabilis, Brachymyrmex patagonicus, Camponotus aethiops, Camponotus claviscapus, Camponotus floridanus, Camponotus fragilis, Camponotus nawai, Camponotus novogranadensis, Camponotus obscuripes, Camponotus sanctaefidei, Camponotus saxatilis, Cataglyphis cursor, Colobopsis truncata, Crematogaster ashmeadi, Crematogaster cerasi, Crematogaster matsumurai, Crematogaster scutellaris, Crematogaster stollii, Cryptopone ochracea, Cryptopone testacea, Dinomyrmex gigas, Dolichoderus bispinosus, Dolichoderus debilis, Dolichoderus lutosus, Dolichoderus quadripunctatus, Dolichoderus sibiricus, Dorymyrmex bureni, Dorymyrmex insanus, Ectatomma ruidum, Ectatomma tuberculatum, Forelius pruinosus, Formica altipetens, Formica argentea, Formica bruni, Formica brunneonitida, Formica caucasicola, Formica cinerea, Formica exsecta, Formica foreli, Formica fusca, Formica fuscocinerea, Formica gagates, Formica gagatoides, Formica glacialis, Formica incerta, Formica japonica, Formica lemani, Formica longiceps, Formica manchu, Formica mesasiatica, Formica neogagates, Formica pisarskii, Formica podzolica, Formica pratensis, Formica pressilabris, Formica rufibarbis, Formica sanguinea, Formica selysi, Formica subaenescens, Formica subsericea, Formica transkaucasica, Formica truncorum, Formica ulkei, Formica uralensis, Formica yessensis, Formicoxenus nitidulus, Gnamptogenys continua, Gnamptogenys hartmani, Harpagoxenus sublaevis, Hypoponera nippona, Hypoponera punctatissima, Lasius alienus, Lasius austriacus, Lasius bicornis, Lasius brevicornis, Lasius brunneus, Lasius carniolicus, Lasius citrinus, Lasius claviger, Lasius distinguendus, Lasius emarginatus, Lasius flavus, Lasius fuliginosus, Lasius hayashi, Lasius japonicus, Lasius jensi, Lasius latipes, Lasius meridionalis, Lasius mixtus, Lasius myops, Lasius nearcticus, Lasius neoniger, Lasius niger, Lasius paralienus, Lasius platythorax, Lasius productus, Lasius psammophilus, Lasius reginae, Lasius sabularum, Lasius sakagamii, Lasius sonobei, Lasius spathepus, Lasius subumbratus, Lasius talpa, Lasius umbratus, Leptogenys punctaticeps, Leptothorax acervorum, Leptothorax gredleri, Leptothorax kutteri, Leptothorax muscorum, Manica rubida, Manica yessensis, Mayaponera arhuaca, Mayaponera constricta, Megalomyrmex symmetochus, Meranoplus bicolor, Messor structor, Myrmecina americana, Myrmecina graminicola, Myrmecocystus mexicanus, Myrmecocystus mimicus, Myrmecocystus navajo, Myrmecocystus yuma, Myrmica aloba, Myrmica bibikoffi, Myrmica constricta, Myrmica deplanata, Myrmica forcipata, Myrmica gallienii, Myrmica hellenica, Myrmica hirsuta, Myrmica karavajevi, Myrmica kotokui, Myrmica kurokii, Myrmica lobicornis, Myrmica lobulicornis, Myrmica lonae, Myrmica pisarskii, Myrmica rubra, Myrmica ruginodis, Myrmica rugulosa, Myrmica sabuleti, Myrmica salina, Myrmica scabrinodis, Myrmica schencki, Myrmica specioides, Myrmica sulcinodis, Myrmica vandeli, Myrmica wesmaeli, Neivamyrmex melshaemeri, Neivamyrmex pseudops, Odontomachus bauri, Paraponera clavata, Pheidole bicarinata, Pheidole desertorum, Pheidole gilvescens, Pheidole megacephala, Pheidole navigans, Pheidole obtusospinosa, Pheidole pallidula, Pheidole pilifera, Pheidole tysoni, Pheidole xerophila, Plagiolepis pallescens, Plagiolepis xene, Pogonomyrmex barbatus, Pogonomyrmex californicus, Pogonomyrmex maricopa, Pogonomyrmex occidentalis, Pogonomyrmex rugosus, Polyergus breviceps, Polyergus lucidus, Polyergus mexicanus, Polyergus rufescens, Ponera coarctata, Ponera kohmoku, Proceratium chickasaw, Procryptocerus belti, Pseudomyrmex gracilis, Solenopsis fugax, Solenopsis krockowi, Solenopsis molesta, Solenopsis xyloni, Stenamma debile, Stigmatomma pallipes, Strongylognathus alpinus, Strongylognathus bulgaricus, Strongylognathus huberi, Strongylognathus italicus, Strongylognathus kratochvili, Strongylognathus testaceus, Strumigenys dolichognatha, Strumigenys elongata, Strumigenys gundlachi, Strumigenys lewisi, Strumigenys margaritae, Strumigenys membranifera, Strumigenys zeteki, Syscia augustae, Tapinoma magnum, Temnothorax affinis, Temnothorax corticalis, Temnothorax crassispinus, Temnothorax curvispinosus, Temnothorax interruptus, Temnothorax nigriceps, Temnothorax nylanderi, Temnothorax parvulus, Temnothorax ravouxi, Temnothorax tuberum, Temnothorax unifasciatus, Temnothorax vivianoi, Tetramorium atratulum, Tetramorium caespitum, Tetramorium impurum, Tetramorium semilaeve, Trachymyrmex septentrionalis, Typhlomyrmex rogenhoferi, Wadeura guianensis, Wasmannia auropunctata, Xenomyrmex stollii
September Aphaenogaster fulva, Aphaenogaster occidentalis, Aphaenogaster subterranea, Azteca instabilis, Brachymyrmex patagonicus, Camponotus claviscapus, Camponotus floridanus, Camponotus novogranadensis, Camponotus sanctaefidei, Camponotus yogi, Cataglyphis cursor, Crematogaster ashmeadi, Crematogaster cerasi, Crematogaster matsumurai, Crematogaster osakensis, Crematogaster scutellaris, Crematogaster stollii, Cryptopone ochracea, Cyphomyrmex costatus, Dinomyrmex gigas, Dolichoderus bispinosus, Dolichoderus debilis, Dolichoderus lutosus, Dolichoderus quadripunctatus, Dorymyrmex insanus, Ectatomma ruidum, Ectatomma tuberculatum, Formica fuscocinerea, Formica japonica, Formica lemani, Formica pratensis, Formica sanguinea, Formica selysi, Gnamptogenys continua, Gnamptogenys hartmani, Hypoponera punctatissima, Lasius alienus, Lasius austriacus, Lasius bicornis, Lasius brevicornis, Lasius capitatus, Lasius carniolicus, Lasius claviger, Lasius distinguendus, Lasius flavus, Lasius fuliginosus, Lasius jensi, Lasius latipes, Lasius meridionalis, Lasius mixtus, Lasius myops, Lasius nearcticus, Lasius neoniger, Lasius niger, Lasius paralienus, Lasius productus, Lasius reginae, Lasius sabularum, Lasius sakagamii, Lasius sonobei, Lasius talpa, Lasius umbratus, Leptogenys punctaticeps, Leptothorax acervorum, Leptothorax muscorum, Manica rubida, Mayaponera arhuaca, Mayaponera constricta, Meranoplus bicolor, Messor barbarus, Messor structor, Myrmecina americana, Myrmecina graminicola, Myrmica aloba, Myrmica constricta, Myrmica deplanata, Myrmica gallienii, Myrmica hellenica, Myrmica hirsuta, Myrmica jessensis, Myrmica karavajevi, Myrmica kotokui, Myrmica lobicornis, Myrmica lobulicornis, Myrmica lonae, Myrmica pisarskii, Myrmica punctiventris, Myrmica rugulosa, Myrmica sabuleti, Myrmica scabrinodis, Myrmica specioides, Myrmica sulcinodis, Myrmica vandeli, Myrmica wesmaeli, Neivamyrmex pseudops, Odontomachus bauri, Paraponera clavata, Pheidole obtusospinosa, Pogonomyrmex barbatus, Pogonomyrmex occidentalis, Pogonomyrmex rugosus, Polyergus breviceps, Polyergus lucidus, Polyergus rufescens, Polyrhachis lamellidens, Ponera coarctata, Procryptocerus belti, Pseudomyrmex gracilis, Solenopsis fugax, Stenamma debile, Stenamma nipponense, Stigmatomma pallipes, Strongylognathus alpinus, Strongylognathus bulgaricus, Strongylognathus huberi, Strongylognathus italicus, Strongylognathus kratochvili, Strumigenys dolichognatha, Strumigenys elongata, Strumigenys gundlachi, Strumigenys membranifera, Tapinoma magnum, Technomyrmex gibbosus, Temnothorax corticalis, Temnothorax crassispinus, Temnothorax interruptus, Temnothorax nigriceps, Temnothorax nylanderi, Temnothorax parvulus, Tetramorium atratulum, Tetramorium impurum, Wadeura guianensis, Wasmannia auropunctata, Xenomyrmex stollii
October Aphaenogaster fulva, Azteca instabilis, Brachymyrmex patagonicus, Camponotus novogranadensis, Camponotus sanctaefidei, Camponotus substitutus, Crematogaster cerasi, Crematogaster lineolata, Crematogaster scutellaris, Crematogaster stollii, Dinomyrmex gigas, Dolichoderus bispinosus, Dolichoderus debilis, Dolichoderus lutosus, Dorymyrmex insanus, Ectatomma ruidum, Ectatomma tuberculatum, Formica transmontanis, Gnamptogenys continua, Gnamptogenys hartmani, Iridomyrmex purpureus, Lasius carniolicus, Lasius claviger, Lasius neoniger, Lasius paralienus, Lasius sabularum, Lasius sakagamii, Leptogenys punctaticeps, Mayaponera constricta, Meranoplus bicolor, Messor barbarus, Myrmecina americana, Myrmica gallienii, Myrmica hellenica, Myrmica kotokui, Myrmica punctiventris, Myrmica rugulosa, Neivamyrmex pseudops, Nylanderia vaga, Odontomachus bauri, Paraparatrechina sakurae, Paraponera clavata, Polyrhachis lamellidens, Pseudomyrmex gracilis, Stenamma debile, Strongylognathus alpinus, Strumigenys dolichognatha, Strumigenys membranifera, Tetramorium impurum, Wadeura guianensis, Wasmannia auropunctata, Xenomyrmex stollii
November Azteca instabilis, Camponotus cingulatus, Camponotus novogranadensis, Camponotus sanctaefidei, Camponotus substitutus, Dinomyrmex gigas, Dolichoderus bispinosus, Dolichoderus debilis, Dolichoderus lutosus, Dorymyrmex insanus, Ectatomma ruidum, Ectatomma tuberculatum, Formica transmontanis, Gnamptogenys continua, Lasius claviger, Lasius neoniger, Leptogenys punctaticeps, Meranoplus bicolor, Messor barbarus, Messor capitatus, Messor ebeninus, Myrmecia tarsata, Myrmica punctiventris, Odontomachus bauri, Pachycondyla harpax, Paraparatrechina sakurae, Paraponera clavata, Pheidole christopherseni, Polyrhachis lamellidens, Procryptocerus belti, Pseudomyrmex gracilis, Strumigenys dolichognatha, Strumigenys membranifera, Wadeura guianensis, Wasmannia auropunctata, Xenomyrmex stollii
December Azteca instabilis, Camponotus consobrinus, Camponotus novogranadensis, Cryptopone testacea, Cyphomyrmex costatus, Dinomyrmex gigas, Dolichoderus debilis, Dolichoderus lutosus, Dorymyrmex insanus, Ectatomma ruidum, Ectatomma tuberculatum, Gnamptogenys continua, Lasius claviger, Lasius neoniger, Leptogenys punctaticeps, Mayaponera constricta, Meranoplus bicolor, Messor capitatus, Messor ebeninus, Myrmicocrypta dilacerata, Neivamyrmex pseudops, Odontomachus bauri, Pachycondyla harpax, Paraponera clavata, Pheidole christopherseni, Prenolepis imparis, Procryptocerus belti, Xenomyrmex stollii

References