What is the name of this small black insect?

Location: South India

Size: 1-2 cm

Precise identification of insects requires use of microscopic lens and an immobile specimen.
I have neither of those here; also, my reference material is focused on North American species. Nevertheless, with what I have available to me, it is most likely a member of order Hemiptera, family Alydidae:

Alydidae are mostly active during the day, so your specimen was probably resting and saw a nearby artificial source of light. They use their piercing mouth-parts to suck the juices out of herbaceous seeds. Unless yours was discovered in a heavily urbanized area, I would venture that this one feeds off nearby grasses and trees.

What is the name of this small black insect? - Biology

Mites are small arachnids (eight-legged arthropods).

Mites are not a defined taxon, but the name is used for members of several groups in the subclass acari of the class Arachnida. Mites span two different groups of arachnids:

Most mites are tiny, less than 1 mm (0.04 in) in length, and have a simple, unsegmented body plan. Their small size makes them easily overlooked some species live in water, many live in soil as decomposers, others live on plants, sometimes creating galls, while others again are predators or parasites. This last type includes the commercially important Varroa parasite of honey bees, as well as the scabies mite of humans. Most species are harmless to humans but a few are associated with allergies or may transmit diseases.

The scientific discipline devoted to the study of ticks and mites is called acarology.


The name is directly from the onomatopoeic Latin cicada. [2] [3] [b]

The superfamily Cicadoidea is a sister of the Cercopoidea (the froghoppers). Cicadas are arranged into two families, the Tettigarctidae and Cicadidae. The two extant species of the Tettigarctidae include one in southern Australia and the other in Tasmania. The family Cicadidae is subdivided into the subfamilies Cicadinae, Tibicininae (or Tettigadinae), Tettigomyiinae, and Cicadettinae [5] [6] they are found on all continents except Antarctica. Some previous works also included a family-level taxon called the Tibiceninae. The largest species is the Malaysian emperor cicada Megapomponia imperatoria its wingspan is up to about 20 cm (8 in). [7] Cicadas are also notable for the great length of time some species take to mature. [8]

At least 3000 cicada species are distributed worldwide, with the majority being in the tropics. Most genera are restricted to a single biogeographical region, and many species have a very limited range. This high degree of endemism has been used to study the biogeography of complex island groups such as in Indonesia and Asia. [9] There are several hundred described species in Australia and New Zealand, [c] around 150 in South Africa, over 170 in America north of Mexico, [10] at least 800 in Latin America, [11] and over 200 in Southeast Asia and the Western Pacific. [12]

About 100 species occur in the Palaearctic. A few species are found in southern Europe, [8] and a single species was known from England, the New Forest cicada, Cicadetta montana, which also occurs in continental Europe. [13] Many species await formal description and many well-known species are yet to be studied carefully using modern acoustic analysis tools that allow their songs to be characterized.

Many of the North American species are the annual or jarfly or dog-day cicadas, members of the Neotibicen, Megatibicen, or Hadoa genera, so named because they emerge in late July and August. [14] The best-known North American genus, however, may be Magicicada. These periodical cicadas have an extremely long life cycle of 13 or 17 years, with adults suddenly and briefly emerging in large numbers. [14] [15]

Australian cicadas are found on tropical islands and cold coastal beaches around Tasmania, in tropical wetlands, high and low deserts, alpine areas of New South Wales and Victoria, large cities including Sydney, Melbourne, and Brisbane, and Tasmanian highlands and snowfields. Many of them have common names such as cherry nose, brown baker, red eye, greengrocer, yellow Monday, whisky drinker, double drummer, and black prince. The Australian greengrocer, Cyclochila australasiae, is among the loudest insects in the world. [16]

More than 40 species from five genera populate New Zealand, ranging from sea level to mountain tops, and all are endemic to New Zealand and its surrounding islands (Kermadec Islands, Chatham Islands). One species is found on Norfolk Island, which technically is part of Australia. [17] The closest relatives of the NZ cicadas live in New Caledonia and Australia.

Palaeontology Edit

Fossil Cicadomorpha first appeared in the Late Triassic. The superfamily Palaeontinoidea contains three families. The Upper Permian Dunstaniidae are found in Australia and South Africa, and also in younger rocks from China. The Upper Triassic Mesogereonidae are found in Australia and South Africa. [18] This group, though, is currently thought to be more distantly related to Cicadomorpha than previously thought. [19]

The Palaeontinidae or "giant cicadas" come from the Jurassic and Lower Cretaceous of Eurasia and South America. [18] The first of these was a fore wing discovered in the Taynton Limestone Formation of Oxfordshire, England it was initially described as a butterfly in 1873, before being recognised as a cicada-like form and renamed Palaeontina oolitica. [20]

Most fossil Cicadidae are known from the Cenozoic, [21] and the oldest unambiguously identified specimen is Davispia bearcreekensis (subfamily Tibicininae) from 59–56 million years ago (Mya). One fossil genus and species (Burmacicada protera) based on a first-instar nymph has recently been reported from 98–99 Mya in the Late Cretaceous, [22] although questions remain about its assignment to the Cicadidae. [21]

Description Edit

Cicadas are large insects made conspicuous by the courtship calls of the males. They are characterized by having three joints in their tarsi, and having small antennae with conical bases and three to six segments, including a seta at the tip. [23] The Auchenorrhyncha differ from other hemipterans by having a rostrum that arises from the posteroventral part of the head, complex sound-producing membranes, and a mechanism for linking the wings that involves a down-rolled edging on the rear of the fore wing and an upwardly protruding flap on the hind wing. [9]

Cicadas are feeble jumpers, and nymphs lack the ability to jump altogether. Another defining characteristic is the adaptations of the fore limbs of nymphs for underground life. The relict family Tettigarctidae differs from the Cicadidae in having the prothorax extending as far as the scutellum, and by lacking the tympanal apparatus. [9]

The adult insect, known as an imago, is 2 to 5 cm (1 to 2 in) in total length in most species. The largest, the empress cicada (Megapomponia imperatoria), has a head-body length around 7 cm (2.8 in), and its wingspan is 18–20 cm (7–8 in). [8] [24] Cicadas have prominent compound eyes set wide apart on the sides of the head. The short antennae protrude between the eyes or in front of them. They also have three small ocelli located on the top of the head in a triangle between the two large eyes this distinguishes cicadas from other members of the Hemiptera. The mouthparts form a long, sharp rostrum that they insert into the plant to feed. [25] The postclypeus is a large, nose-like structure that lies between the eyes and makes up most of the front of the head it contains the pumping musculature. [26]

The thorax has three segments and houses the powerful wing muscles. They have two pairs of membranous wings that may be hyaline, cloudy, or pigmented. The wing venation varies between species and may help in identification. The middle thoracic segment has an operculum on the underside, which may extend posteriorly and obscure parts of the abdomen. The abdomen is segmented, with the hindermost segments housing the reproductive organs, and terminates in females with a large, saw-edged ovipositor. In males, the abdomen is largely hollow and used as a resonating chamber. [25]

The surface of the fore wing is superhydrophobic it is covered with minute, waxy cones, blunt spikes that create a water-repellent film. Rain rolls across the surface, removing dirt in the process. In the absence of rain, dew condenses on the wings. When the droplets coalesce, they leap several millimetres into the air, which also serves to clean the wings. [27] Bacteria landing on the wing surface are not repelled rather, their membranes are torn apart by the nanoscale-sized spikes, making the wing surface the first-known biomaterial that can kill bacteria. [28]

Temperature regulation Edit

Desert cicadas such as Diceroprocta apache are unusual among insects in controlling their temperature by evaporative cooling, analogous to sweating in mammals. When their temperature rises above about 39°C, they suck excess sap from the food plants and extrude the excess water through pores in the tergum at a modest cost in energy. Such a rapid loss of water can be sustained only by feeding on water-rich xylem sap. At lower temperatures, feeding cicadas would normally need to excrete the excess water. By evaporative cooling, desert cicadas can reduce their bodily temperature by some 5°C. [29] [30] Some non-desert cicada species such as Magicicada tredecem also cool themselves evaporatively, but less dramatically. [31] Conversely, many other cicadas can voluntarily raise their body temperatures as much as 22 °C (40 °F) above ambient temperature. [32]

Song Edit

The "singing" of male cicadas is produced principally and in the majority of species using a special structure called a tymbal, a pair of which lies below each side of the anterior abdominal region. The structure is buckled by muscular action and being made of resilin unbuckled rapidly on muscle relaxation and the rapid action of muscles produces their characteristic sounds. Some cicadas however, have mechanisms for stridulation, sometimes in addition to the tymbals. Here, the wings are rubbed over a series of midthoracic ridges. In the Chinese species Subpsaltria yangi, both males and females can stridulate. [33] The sounds may further be modulated by membranous coverings and by resonant cavities. [23]

The male abdomen in some species is largely hollow, and acts as a sound box. By rapidly vibrating these membranes, a cicada combines the clicks into apparently continuous notes, and enlarged chambers derived from the tracheae serve as resonance chambers with which it amplifies the sound. The cicada also modulates the song by positioning its abdomen toward or away from the substrate. Partly by the pattern in which it combines the clicks, each species produces its own distinctive mating songs and acoustic signals, ensuring that the song attracts only appropriate mates. [14] The tettigarctid (or hairy) cicadas Tettigarcta crinita of Australia and T. tomentosa have rudimentary tymbals in both sexes and do not produce airborne sounds. Both males and females produce vibrations that are transmitted through the tree substrate. They are considered as representing the original state from which other cicada communication has evolved. [34]

Average temperature of the natural habitat for the South American species Fidicina rana is about 29 °C (84 °F). During sound production, the temperature of the tymbal muscles was found to be significantly higher. [35] Many cicadas sing most actively during the hottest hours of a summer day roughly a 24-hour cycle. [36] Most cicadas are diurnal in their calling and depend on external heat to warm them up, while a few are capable of raising their temperatures using muscle action and some species are known to call at dusk. [32] Kanakia gigas and Froggattoides typicus are among the few that are known to be truly nocturnal and there may be other nocturnal species living in tropical forests. [37] [38]

Cicadas call from varying heights on trees. Where multiple species occur, the species may use different heights and timing of calling. [39] [40] While the vast majority of cicadas call from above the ground, two Californian species, Okanagana pallidula and O. vanduzeei are known to call from hollows made at the base of the tree below the ground level. The adaptive significance is unclear, as the calls are not amplified or modified by the burrow structure, but this may avoid predation. [41]

Although only males produce the cicadas' distinctive sounds, both sexes have membranous structures called tympana (singular – tympanum) by which they detect sounds, the equivalent of having ears. Males disable their own tympana while calling, thereby preventing damage to their hearing [42] a necessity partly because some cicadas produce sounds up to 120 dB (SPL) [42] which is among the loudest of all insect-produced sounds. [43] The song is loud enough to cause permanent hearing loss in humans should the cicada be at "close range". In contrast, some small species have songs so high in pitch that they are inaudible to humans. [44]

For the human ear, telling precisely where a cicada song originates is often difficult. The pitch is nearly constant, the sound is continuous to the human ear, and cicadas sing in scattered groups. In addition to the mating song, many species have a distinct distress call, usually a broken and erratic sound emitted by the insect when seized or panicked. Some species also have courtship songs, generally quieter, and produced after a female has been drawn to the calling song. Males also produce encounter calls, whether in courtship or to maintain personal space within choruses. [45]

The song of cicadas is considered by entomologists to be unique to a given species, and a number of resources exist to collect and analyse cicada sounds. [46]

Life cycle Edit

In some species of cicadas, the males remain in one location and call to attract females. Sometimes, several males aggregate and call in chorus. In other species, the males move from place to place, usually with quieter calls, while searching for females. The Tettigarctidae differ from other cicadas in producing vibrations in the substrate rather than audible sounds. [9] After mating, the female cuts slits into the bark of a twig where she deposits her eggs. [9] Both male and female cicadas die within a few weeks after emerging from the soil. Although they have mouthparts and are able to consume some plant liquids for nutrition, the amount eaten is very small and the insects have a natural adult lifespan of less than two months.

When the eggs hatch, the newly hatched nymphs drop to the ground and burrow. Cicadas live underground as nymphs for most of their lives at depths down to about 2.5 m (8 ft). Nymphs have strong front legs for digging and excavating chambers in close proximity to roots, where they feed on xylem sap. In the process, their bodies and interior of the burrow become coated in anal fluids. In wet habitats, larger species construct mud towers above ground to aerate their burrows. In the final nymphal instar, they construct an exit tunnel to the surface and emerge. [9] They then moult (shed their skins) on a nearby plant for the last time, and emerge as adults. The exuviae or abandoned exoskeletons remain, still clinging to the bark of the tree. [47]

Most cicadas go through a life cycle that lasts 2–5 years. Some species have much longer life cycles, such as the North American genus, Magicicada, which has a number of distinct "broods" that go through either a 17-year, or in some parts of the region, a 13-year life cycle. The long life cycles may have developed as a response to predators, such as the cicada killer wasp and praying mantis. [48] [49] [50] A specialist predator with a shorter life cycle of at least two years could not reliably prey upon the cicadas. [51] An alternate hypothesis is that these long life cycles evolved during the ice ages so as to overcome cold spells and that as species co-emerged and hybridized they left distinct species that did not hybridize having periods matching prime numbers. [52]

A teneral cicada that has just emerged and is waiting to dry before flying away

Homeowners should consider working with a licensed pest professional to employ a preventative pest management plan. There are also a few things that can be done around the property to prevent a little black ant infestation.

Homeowners should seal cracks and crevices in exterior walls with a silicone-based caulk, ensure firewood is stored at least 20 feet away from the home, and keep shrubbery well trimmed. Location of the nest is also important. While it can be difficult to see these ants due to their small size, their nests can be found by following the trial of workers back to the colony.


Overwintering adult boxelder bugs emerge from hibernation in late March to early April when the boxelder buds open. During this time, the adults leave their overwintering sites to return to their host trees for the warmer months. They first feed on fallen boxelder seeds and later move to the female boxelder trees or maple trees where they eat newly developing leaves. Occasionally, boxelder bugs will feed on the fruits of plum and apple trees.

The females lay clusters of straw-yellow eggs on stones, eaves, grass, shrubs and trees – especially in the bark crevices of boxelder trees. The eggs turn red as the embryos develop and hatch about two weeks later. The nymphs feed on fallen boxelder seeds and later on new leaves. There are two generations per year in the warmer regions of the United States.

In the fall, boxelder bugs become gregarious and congregate on the south side of rocks, trees and buildings where the sun hits. After large masses gather, they migrate to nearby buildings or homes to overwinter. These pests tend to hide in small cracks and crevices in walls to insulate themselves from the cold winter temperatures.

What is the name of this small black insect? - Biology

Biting midges can be a nuisance to campers, fishermen, hunters, hikers, gardeners, and others who spend time outdoors during early morning and evenings, and even during the daytime on cloudy days when winds are calm. They will readily bite humans the bites are irritating, painful, and can cause long-lasting painful lesions for some people.

A common observation upon experiencing a bite from this insect is that something is biting, but the person suffering cannot see what it is. Biting midges are sometimes incorrectly referred to as sand flies. Sand flies are insects that belong to a different biological group and should not be confused with the biting midges.

Figure 1. Culicoides furens shown next to a U.S. dime and pencil point to demonstrate the relative size of this adult biting midge species. Photograph by Roxanne Connelly, Florida Medical Entomology Laboratory, University of Florida.

Distribution (Back to Top)

There are over 4,000 species of biting midges in the Ceratopogonidae family, and over 1,000 in just one genus, Culicoides. The distribution of midges in the genus Culicoides is world-wide 47 species are known to occur in Florida. Species belonging to the genus Leptoconops occur in the tropics, sub-tropics, the Caribbean, and some coastal areas of southeast Florida.

The natural habitats of biting midges vary by species. Areas with substantial salt marsh habitat are major producers of many biting midge species. Additional sources for some species, like the bluetongue virus vector Culicoides sonorensis Wirth and Jones, include highly organic soil that is wet but not underwater such as those found with high manure loads in swine-, sheep- and cattle-farming operations. These insects do not establish inside homes, apartments, or inside humans or other animals.

Description (Back to Top)

Immature Stages: The eggs can be cigar-, banana-, or sausage-shaped and approximately 0.25 mm long. They are white when first laid but later turn brown or black. The eggs are laid on moist soil and cannot withstand drying out. Some species can lay up to 450 eggs per batch and as many as seven batches in a lifespan. Eggs typically hatch within two to 10 days of being laid time to hatch is dependent on the species and temperatures.

The larvae are worm-like, creamy white, and approximately 2 to 5 mm long. Larvae develop through four instars the first instar larvae possess a functional spine-bearing proleg. Pupal color can be pale yellow to light brown to dark brown. They are 2 to 5 mm in length with an unsegmented cephalothorax that has a pair of respiratory horns that may bear spines or wrinkles. During this stage, the insects possess a spiny integument which can be used to identify the fly to species level.

Adults: The adult no-see-ums are gray and less than 1/8-inch long. The two wings possess dense hairs and give rise to pigmentation patterns. These wing patterns are used by biologists to identify species. The large compound eyes are more or less contiguous above the bases of the 15-segmented antennae. The pedicel of the males' antennae houses the Johnston's organ. The mouthparts are well-developed with cutting teeth on elongated mandibles in the proboscis, adapted for blood-sucking in females, but not in males. The thorax extends slightly over the head, and the abdomen is nine-segmented and tapered at the end.

Figure 2. Adult biting midge, Culicoides sonorensis Wirth and Jones, showing blood-filled abdomen and the characteristic wings patterns used for species identification. Photograph by Ed T. Schmidtmann, USDA/ARS.

Life Cycle (Back to Top)

Adults: Biting midges are holometabolous, progressing from egg to larva to pupa, and finally to the adult stage. The complete cycle can occur in two to six weeks, but is dependent on the species and environmental conditions. The adults are most abundant near productive breeding sites, but will disperse to mate and to feed. The mean distance for female flight is 2 km, less than half of that distance for males.

Male Culicoides typically emerge before the females and are ready to mate when the female emerges from the pupal stage. Mating typically occurs in flight when females fly into swarms of males and the insects are oriented end to end with the ventral parts of the genitalia in contact. Some species mate without swarming instead, the males go to hosts where the female is likely to feed on blood mating occurs when she finishes feeding.

Eggs: Males and females feed on nectar, but the females require blood for their eggs to mature. The females will blood-feed primarily around dawn and dusk however, there are some species that prefer to feed during the day. Some species are autogenous and therefore may produce the first batch of viable eggs without a blood meal using reserves stored from the larval period blood meals are required for subsequent batches of eggs.

The number of eggs produced varies among species and size of bloodmeal. For example, Culicoides furens (Poey) can lay 50 to 110 eggs per bloodmeal, and C. mississippiensis Hoffman, 25 to 50 eggs per bloodmeal. The adults can live two to seven weeks in a laboratory setting, but only a few weeks under natural conditions.

Larvae: Larvae require water, air and food and are not strictly aquatic or terrestrial. They cannot develop without moisture. The larvae are present in and around salt-marsh and mangrove swamps, on shores of streams and ponds, and in muddy substrates. They feed on small organisms. Most species cannot exist more than a few inches below the air-water interface.

In the tropics, the larval habitat of many species is in rotting fruit, bromeliads, and other water-holding plants. Other larval habitats include mud, sand, and debris at edges of ponds, lakes and springs, tree holes, and slime-covered bark. The larval stage can last from two weeks to a year, depending on the species, temperatures, and geographic area.

While some larvae can develop in wet manure-contaminated areas (Mullen 2002), they do not develop inside the animal. The larvae also do not develop inside humans or other animals.

Pupae: The pupal stage typically lasts

Medical Significance (Back to Top)

In the U.S., the biting midges are primarily a nuisance and the major medical issue associated with Culicoides is allergic reactions to the bites. However, like other blood feeding Diptera, Culicoides species are vectors of pathogens that can cause disease in humans and animals. In Central and South America, western and central Africa, and some Caribbean islands, biting midges are the vectors of filarial worms in the genus Mansonella. These parasites cause infection in humans that produces dermatitis and skin lesions because the adult worms are located in the skin.

Biting midges, primarily the species Culicoides sonorensis, are responsible for transmission of bluetongue virus to sheep and cattle in the U.S. Bluetongue is a serious disease of ruminants. Bluetongue viruses are found world-wide and are transmitted by different Culicoides species in different regions. Many countries that are bluetongue free prohibit the movement of livestock from bluetongue endemic regions. The annual economic damage in lost trade is in the millions of dollars.

Other animal disease causing pathogens transmitted by the bite of infected biting midges include African Horsesickness virus in equines that is confined primarily to Africa and Epizootic Hemorrhagic Disease virus in ruminants found in North America and principally having lethal effects on deer. Some equines experience allergic reactions to the bites, resulting in equine allergic dermatitis, affecting the withers, mane, tail and ears of the animal.

Management and Prevention (Back to Top)

Historically, management methods included diking and drainage of marshlands to reduce the habitats used by the immature stages. The insecticide DDT was used to target the adult stage. Currently, larval habitats are not targeted in control efforts because of the extensive amount of area that the habitats may cover, some negative environmental impacts resulting from changing water flow patterns of large areas, and the spotty spatial distribution of larvae within a given habitat.

Applications of insecticides targeting the adult stage are not efficient. While this type of application may kill biting midges active on a given night, they are continually dispersing from the larval habitat and entering areas of human activity. It would require insecticide applications on a daily basis in some areas, and this is not efficient or environmentally sound. Many government agencies that provide mosquito control services receive complaint calls about biting midges. However, most of the programs are not mandated or allowed to respond by providing control measures.

On a large scale, removal trapping is conducted using CO2 as an attractant to lure the biting midges to an insecticide-treated target where they are killed. Research from the the University of Florida, Institute of Food and Agricultural Sciences Florida Medical Entomology Laboratory showed that biting midge populations were reduced in test areas of Vero Beach and Boynton Beach, FL, and Castaway Cay, Bahamas. This method of control is more appropriate for islands and specific inland areas where pest control personnel can make a long term commitment to this technique.

Homeowners can install proper screening for windows and patios to prevent no-see-ums from entering residences and outdoor areas used for leisure and entertaining. Most biting midges can pass through 16-mesh insect wire screen and netting, so a smaller mesh size is required. The small mesh size does limit air flow through the screens. Additionally, because no-see-ums are so small and are weak fliers, ceiling and window fans can be used at high speeds to keep no-see-ums out of small areas.

Repellents containing DEET (N,N-diethyl-meta-toluamide) typically used as mosquito repellents are also labeled for use against no-see-ums and can be applied prior to exposure to the biting midges. It is important that the directions for application that are printed on the label are followed for any product used as a repellent.

Coastal areas provide primary habitat for biting midges. Tourists and potential home and land owners can consult local maps prior to visiting or purchasing property in coastal areas, to determine the proximity to biting midge producing areas. It is prudent to research the area of geographic interest prior to making decisions that can lead to an unpleasant vacation or unhappy homeowners. Knowing the habitats, and that large scale control operations are not feasible, one can be prepared with repellents or make decisions to build, or visit, elsewhere.

Selected References (Back to Top)

  • Blanton FS, Wirth WW. 1979. The sand flies (Culicoides) of Florida (Diptera: Ceratopogonidae). Arthropods of Florida and Neighboring Land Areas Volume 10. Florida Department of Agriculture and Consumer Services. Gainesville, FL. 204 pp.
  • Day, JF, Duxbury, CG, Glasscock, S and Paganessi, JE. 2001. Removal trapping for the control of coastal biting midge populations. Technical Bulletin of the Florida Mosquito Control Association. 4th Workshop on Salt Marsh Management and Research. Florida Mosquito Control Association, Ft. Myers, FL. 3: 15-16.
  • Eldridge, BF and Edman, JD, Eds. 2000. Medical Entomology: A Textbook on Public Health and Veterinary Problems Caused by Arthropods. Kluwer Academic Publishers, Dordrecht, The Netherlands.
  • Foote RH, Pratt HD. 1954. The Culicoides of the eastern United States (Diptera, Heleidae). Public Health Monograph No. 18. Publication No. 296. U. S. Department of Health, Education and Welfare, Public Health Service. 53 pp.
  • Holbrook FR. 1996. Biting midges and the agents they transmit. In Beaty BJ, Marquardt WC (Eds), The Biology of Disease Vectors. University Press of Colorado, Niwot, CO. p. 110-116.
  • Mullen G. Biting midges (Ceratopogonidae). In Mullen G, Durden L (Eds). 2002. Medical and Veterinary Entomology. Elsevier Science, San Diego, CA. p. 163-183.
  • Rutledge CR, Day JF. 2002. Mosquito Repellents. EDIS. University of Florida/IFAS. (15 June 2016)

Author: C. Roxanne Connelly, University of Florida, Entomology and Nematology Department
Photographs: C. Roxanne Connelly, University of Florida, Entomology and Nematology Department Ed T. Schmidtmann, United States Department of Agriculture, Agricultural Research Services
Web Design: Don Wasik, Jane Medley
Publication Number: EENY-349
Publication Date: May 2005. Latest Revision: August 2013. Reviewed: June 2019.


Crickets are small to medium-sized insects with mostly cylindrical, somewhat vertically flattened bodies. The head is spherical with long slender antennae arising from cone-shaped scapes (first segments) and just behind these are two large compound eyes. On the forehead are three ocelli (simple eyes). The pronotum (first thoracic segment) is trapezoidal in shape, robust, and well-sclerotinized. It is smooth and has neither dorsal nor lateral keels (ridges). [4]

At the tip of the abdomen is a pair of long cerci (paired appendages on rearmost segment), and in females, the ovipositor is cylindrical, long and narrow, smooth and shiny. The femora (third segments) of the back pair of legs are greatly enlarged for jumping. The tibiae (fourth segments) of the hind legs are armed with a number of moveable spurs, the arrangement of which is characteristic of each species. The tibiae of the front legs bear one or more tympani which are used for the reception of sound. [4]

The wings lie flat on the body and are very variable in size between species, being reduced in size in some crickets and missing in others. The fore wings are elytra made of tough chitin, acting as a protective shield for the soft parts of the body and in males, bear the stridulatory organs for the production of sound. The hind pair is membranous, folding fan-wise under the fore wings. In many species, the wings are not adapted for flight. [1]

The largest members of the family are the 5 cm (2 in)-long bull crickets (Brachytrupes) which excavate burrows a metre or more deep. The tree crickets (Oecanthinae) are delicate white or pale green insects with transparent fore wings, while the field crickets (Gryllinae) are robust brown or black insects. [1]

Crickets have a cosmopolitan distribution, being found in all parts of the world with the exception of cold regions at latitudes higher than about 55° North and South. They have colonised many large and small islands, sometimes flying over the sea to reach these locations, or perhaps conveyed on floating timber or by human activity. The greatest diversity occurs in tropical locations, such as in Malaysia, where 88 species were heard chirping from a single location near Kuala Lumpur. A greater number than this could have been present because some species are mute. [1]

Crickets are found in many habitats. Members of several subfamilies are found in the upper tree canopy, in bushes, and among grasses and herbs. They also occur on the ground and in caves, and some are subterranean, excavating shallow or deep burrows. Some make home in rotting wood, and certain beach-dwelling species can run and jump over the surface of water. [1]

Defence Edit

Crickets are relatively defenceless, soft-bodied insects. Most species are nocturnal and spend the day hidden in cracks, under bark, inside curling leaves, under stones or fallen logs, in leaf litter, or in the cracks in the ground that develop in dry weather. Some excavate their own shallow holes in rotting wood or underground and fold in their antennae to conceal their presence. Some of these burrows are temporary shelters, used for a single day, but others serve as more permanent residences and places for mating and laying eggs. Crickets burrow by loosening the soil with the mandibles and then carrying it with the limbs, flicking it backwards with the hind legs or pushing it with the head. [5]

Other defensive strategies are the use of camouflage, fleeing, and aggression. Some species have adopted colourings, shapes, and patterns that make it difficult for predators that hunt by sight to detect them. They tend to be dull shades of brown, grey, and green that blend into their background, and desert species tend to be pale. Some species can fly, but the mode of flight tends to be clumsy, so the most usual response to danger is to scuttle away to find a hiding place. [5]

Chirping Edit

Most male crickets make a loud chirping sound by stridulation (scraping two specially textured body parts together). The stridulatory organ is located on the tegmen, or fore wing, which is leathery in texture. A large vein runs along the centre of each tegmen, with comb-like serrations on its edge forming a file-like structure, and at the rear edge of the tegmen is a scraper. The tegmina are held at an angle to the body and rhythmically raised and lowered which causes the scraper on one wing to rasp on the file on the other. The central part of the tegmen contains the "harp", an area of thick, sclerotinized membrane which resonates and amplifies the volume of sound, as does the pocket of air between the tegmina and the body wall. Most female crickets lack the necessary adaptations to stridulate, so make no sound. [6]

Several types of cricket songs are in the repertoire of some species. The calling song attracts females and repels other males, and is fairly loud. The courting song is used when a female cricket is near and encourages her to mate with the caller. A triumphal song is produced for a brief period after a successful mating, and may reinforce the mating bond to encourage the female to lay some eggs rather than find another male. [7] An aggressive song is triggered by contact chemoreceptors on the antennae that detect the presence of another male cricket. [8]

Crickets chirp at different rates depending on their species and the temperature of their environment. Most species chirp at higher rates the higher the temperature is (about 62 chirps a minute at 13 °C (55 °F) in one common species each species has its own rate). The relationship between temperature and the rate of chirping is known as Dolbear's law. According to this law, counting the number of chirps produced in 14 seconds by the snowy tree cricket, common in the United States, and adding 40 will approximate the temperature in degrees Fahrenheit. [7]

In 1975, Dr. William H. Cade discovered that the parasitic tachinid fly Ormia ochracea is attracted to the song of the cricket, and uses it to locate the male to deposit her larvae on him. It was the first known example of a natural enemy that locates its host or prey using the mating signal. [9] Since then, many species of crickets have been found to be carrying the same parasitic fly, or related species. In response to this selective pressure, a mutation leaving males unable to chirp was observed amongst a population of Teleogryllus oceanicus on the Hawaiian island of Kauai, enabling these crickets to elude their parasitoid predators. [10] A different mutation with the same effect was also discovered on the neighboring island of Oahu (ca. 100 miles (160 km) away). [11] Recently, new "purring" males of the same species in Hawaii are able to produce a novel auditory sexual signal that can be used to attract females while greatly reducing the likelihood of parasitoid attack from the fly. [12]

Flight Edit

Some species, such as the ground crickets (Nemobiinae), are wingless others have small fore wings and no hind wings (Copholandrevus), others lack hind wings and have shortened fore wings in females only, while others are macropterous, with the hind wings longer than the fore wings. In Teleogryllus, the proportion of macropterous individuals varies from very low to 100%. Probably, most species with hind wings longer than fore wings engage in flight. [4]

Some species, such as Gryllus assimilis, take off, fly, and land efficiently and well, while other species are clumsy fliers. [1] In some species, the hind wings are shed, leaving wing stumps, usually after dispersal of the insect by flight. In other species, they may be pulled off and consumed by the cricket itself or by another individual, probably providing a nutritional boost. [13]

Gryllus firmus exhibits wing polymorphism some individuals have fully functional, long hind wings and others have short wings and cannot fly. The short-winged females have smaller flight muscles, greater ovarian development, and produce more eggs, so the polymorphism adapts the cricket for either dispersal or reproduction. In some long-winged individuals, the flight muscles deteriorate during adulthood and the insect's reproductive capabilities improve. [14]

Diet Edit

Captive crickets are omnivorous when deprived of their natural diet, they accept a wide range of organic foodstuffs. Some species are completely herbivorous, feeding on flowers, fruit, and leaves, with ground-based species consuming seedlings, grasses, pieces of leaf, and the shoots of young plants. Others are more predatory and include in their diet invertebrate eggs, larvae, pupae, moulting insects, scale insects, and aphids. [15] Many are scavengers and consume various organic remains, decaying plants, seedlings, and fungi. [16] In captivity, many species have been successfully reared on a diet of ground, commercial dry dog food, supplemented with lettuce and aphids. [15]

Crickets have relatively powerful jaws, and several species have been known to bite humans. [17]

Reproduction and lifecycle Edit

Male crickets establish their dominance over each other by aggression. They start by lashing each other with their antennae and flaring their mandibles. Unless one retreats at this stage, they resort to grappling, at the same time each emitting calls that are quite unlike those uttered in other circumstances. When one achieves dominance, it sings loudly, while the loser remains silent. [18]

Females are generally attracted to males by their calls, though in nonstridulatory species, some other mechanism must be involved. After the pair has made antennal contact, a courtship period may occur during which the character of the call changes. The female mounts the male and a single spermatophore is transferred to the external genitalia of the female. Sperm flows from this into the female's oviduct over a period of a few minutes or up to an hour, depending on species. After copulation, the female may remove or eat the spermatophore males may attempt to prevent this with various ritualised behaviours. The female may mate on several occasions with different males. [19]

Most crickets lay their eggs in the soil or inside the stems of plants, and to do this, female crickets have a long, needle-like or sabre-like egg-laying organ called an ovipositor. Some ground-dwelling species have dispensed with this, either depositing their eggs in an underground chamber or pushing them into the wall of a burrow. [1] The short-tailed cricket (Anurogryllus) excavates a burrow with chambers and a defecating area, lays its eggs in a pile on a chamber floor, and after the eggs have hatched, feeds the juveniles for about a month. [20]

Crickets are hemimetabolic insects, whose lifecycle consists of an egg stage, a larval or nymph stage that increasingly resembles the adult form as the nymph grows, and an adult stage. The egg hatches into a nymph about the size of a fruit fly. This passes through about 10 larval stages, and with each successive moult, it becomes more like an adult. After the final moult, the genitalia and wings are fully developed, but a period of maturation is needed before the cricket is ready to breed. [21]

Inbreeding avoidance Edit

Some species of cricket are polyandrous. In Gryllus bimaculatus, the females select and mate with multiple viable sperm donors, preferring novel mates. [22] Female Teleogryllus oceanicus crickets from natural populations similarly mate and store sperm from multiple males. [23] Female crickets exert a postcopulatory fertilization bias in favour of unrelated males to avoid the genetic consequences of inbreeding. Fertilization bias depends on the control of sperm transport to the sperm storage organs. The inhibition of sperm storage by female crickets can act as a form of cryptic female choice to avoid the severe negative effects of inbreeding. [24] Controlled-breeding experiments with the cricket Gryllus firmus demonstrated inbreeding depression, as nymphal weight and early fecundity declined substantially over the generations' [25] this was caused as expected by an increased frequency of homozygous combinations of deleterious recessive alleles. [25] [26]

Predators, parasites, and pathogens Edit

Crickets have many natural enemies and are subject to various pathogens and parasites. They are eaten by large numbers of vertebrate and invertebrate predators and their hard parts are often found during the examination of animal intestines. [5] Mediterranean house geckos (Hemidactylus turcicus) have learned that although a calling decorated cricket (Gryllodes supplicans) may be safely positioned in an out-of-reach burrow, female crickets attracted to the call can be intercepted and eaten. [18]

The entomopathogenic fungus Metarhizium anisopliae attacks and kills crickets and has been used as the basis of control in pest populations. [5] The insects are also affected by the cricket paralysis virus, which has caused high levels of fatalities in cricket-rearing facilities. [27] Other fatal diseases that have been identified in mass-rearing establishments include Rickettsia and three further viruses. The diseases may spread more rapidly if the crickets become cannibalistic and eat the corpses. [5]

Red parasitic mites sometimes attach themselves to the dorsal region of crickets and may greatly affect them. [5] The horsehair worm Paragordius varius is an internal parasite and can control the behaviour of its cricket host and cause it to enter water, where the parasite continues its lifecycle and the cricket likely drowns. [28] The larvae of the sarcophagid fly Sarcophaga kellyi develop inside the body cavity of field crickets. [29] Female parasitic wasps of Rhopalosoma lay their eggs on crickets, and their developing larvae gradually devour their hosts. Other wasps in the family Scelionidae are egg parasitoids, seeking out batches of eggs laid by crickets in plant tissues in which to insert their eggs. [5]

The fly Ormia ochracea has very acute hearing and targets calling male crickets. It locates its prey by ear and then lays its eggs nearby. The developing larvae burrow inside any crickets with which they come in contact and in the course of a week or so, devour what remains of the host before pupating. [30] In Florida, the parasitic flies were only present in the autumn, and at that time of year, the males sang less but for longer periods. A trade-off exists for the male between attracting females and being parasitized. [31]

The phylogenetic relationships of the Gryllidae, summarized by Darryl Gwynne in 1995 from his own work (using mainly anatomical characteristics) and that of earlier authors, [a] are shown in the following cladogram, with the Orthoptera divided into two main groups, Ensifera (crickets sensu lato) and Caelifera (grasshoppers). Fossil Ensifera are found from the late Carboniferous period (300 Mya) onwards, [32] [33] and the true crickets, Gryllidae, from the Triassic period (250 to 200 Mya). [1]

Cladogram after Gwynne, 1995: [32]

Gryllidae (true crickets)

Tettigonioidea (katydids, bush crickets, weta)

A phylogenetic study by Jost & Shaw in 2006 using sequences from 18S, 28S, and 16S rRNA supported the monophyly of Ensifera. Most ensiferan families were also found to be monophyletic, and the superfamily Gryllacridoidea was found to include Stenopelmatidae, Anostostomatidae, Gryllacrididae and Lezina. Schizodactylidae and Grylloidea were shown to be sister taxa, and Rhaphidophoridae and Tettigoniidae were found to be more closely related to Grylloidea than had previously been thought. The authors stated that "a high degree of conflict exists between the molecular and morphological data, possibly indicating that much homoplasy is present in Ensifera, particularly in acoustic structures." They considered that tegmen stridulation and tibial tympanae are ancestral to Ensifera and have been lost on multiple occasions, especially within the Gryllidae. [34]

"Cricket" Families Edit

Several families and other taxa in the Ensifera may be called "crickets", including:

    – "true crickets" – scaly crickets – "spider-crickets" and their allies - sword-tail crickets and wood or ground-crickets.
  • other families in the infraorder Gryllidea may be included:
      – mole crickets – ant crickets.
    • – the bush crickets or katydids – which are quite distinct and unrelated, with 4-segmented tarsi (at least in the middle and hind legs) [3] and females with flattened ovipositors. Also note:
      • within this family is the genus Anabrus – the "mormon crickets"
      • "bush crickets" (American usage) include members of the subfamily Trigonidiinae – which are "true crickets".

      Folklore and myth Edit

      The folklore and mythology surrounding crickets is extensive. [35] The singing of crickets in the folklore of Brazil and elsewhere is sometimes taken to be a sign of impending rain, or of a financial windfall. In Álvar Núñez Cabeza de Vaca's chronicles of the Spanish conquest of the Americas, the sudden chirping of a cricket heralded the sighting of land for his crew, just as their water supply had run out. [36] In Caraguatatuba, Brazil, a black cricket in a room is said to portend illness a grey one, money and a green one, hope. [36] In Alagoas state, northeast Brazil, a cricket announces death, thus it is killed if it chirps in a house. [37] In Barbados, a loud cricket means money is coming in hence, a cricket must not be killed or evicted if it chirps inside a house. However, another type of cricket that is less noisy forebodes illness or death. [38]

      In literature Edit

      Crickets feature as major characters in novels and children's books. Charles Dickens's 1845 novella The Cricket on the Hearth, divided into sections called "Chirps", tells the story of a cricket which chirps on the hearth and acts as a guardian angel to a family. [39] Carlo Collodi's 1883 children's book "Le avventure di Pinocchio" (The Adventures of Pinocchio) featured "Il Grillo Parlante" (The Talking Cricket) as one of its characters. [40] George Selden's 1960 children's book The Cricket in Times Square tells the story of Chester the cricket from Connecticut who joins a family and their other animals, and is taken to see Times Square in New York. [41] The story, which won the Newbery Honor, [42] came to Selden on hearing a real cricket chirp in Times Square. [43]

      Souvenirs entomologiques, a book written by the French entomologist Jean-Henri Fabre, devotes a whole chapter to the cricket, discussing its construction of a burrow and its song-making. The account is mainly of the field cricket, but also mentions the Italian cricket. [44]

      Crickets have from time to time appeared in poetry. William Wordsworth's 1805 poem The Cottager to Her Infant includes the couplet "The kitten sleeps upon the hearth, The crickets long have ceased their mirth". [45] John Keats's 1819 poem Ode to Autumn includes the lines "Hedge-crickets sing and now with treble soft / The redbreast whistles from a garden-croft". [46] The Chinese Tang dynasty poet Du Fu (712–770) wrote a poem that in the translation by J. P. Seaton begins "House cricket . Trifling thing. And yet how his mournful song moves us. Out in the grass his cry was a tremble, But now, he trills beneath our bed, to share his sorrow." [47]

      As pets and fighting animals Edit

      Crickets are kept as pets and are considered good luck in some countries in China, they are sometimes kept in cages or in hollowed-out gourds specially created in novel shapes. [48] The practice was common in Japan for thousands of years it peaked in the 19th century, though crickets are still sold at pet shops. [49] It is also common to have them as caged pets in some European countries, particularly in the Iberian Peninsula. Cricket fighting is a traditional Chinese pastime that dates back to the Tang dynasty (618–907). Originally an indulgence of emperors, cricket fighting later became popular among commoners. [50] The dominance and fighting ability of males does not depend on strength alone it has been found that they become more aggressive after certain pre-fight experiences such as isolation, or when defending a refuge. Crickets forced to fly for a short while will afterwards fight for two to three times longer than they otherwise would. [51]

      As food Edit

      In the southern part of Asia including Cambodia, Laos, Thailand, and Vietnam, crickets commonly are eaten as a snack, prepared by deep frying soaked and cleaned insects. [52] In Thailand, there are 20,000 farmers rearing crickets, with an estimated production of 7,500 tons per year [53] and United Nation's FAO has implemented a project in Laos to improve cricket farming and, consequently, food security. [54] The food conversion efficiency of house crickets (Acheta domesticus) is 1.7, some five times higher than that for beef cattle, and if their fecundity is taken into account, 15 to 20 times higher. [55] [56]

      Cricket flour may be used as an additive to consumer foods such as pasta, bread, crackers, and cookies. The cricket flour is being used in protein bars, pet foods, livestock feed, nutraceuticals, and other industrial uses. The United Nations says the use of insect protein, such as cricket flour, could be critical in feeding the growing population of the planet while being less damaging to the environment. [57]

      Crickets are also reared as food for carnivorous zoo animals, laboratory animals, and pets. [5] [58] They may be "gut loaded" with additional minerals, such as calcium, to provide a balanced diet for predators such as tree frogs (Hylidae). [59]

      Common expressions Edit

      By the 19th century "cricket" and "crickets" were in use as euphemisms for using Christ as an interjection. The addition of "Jiminy" (a variation of "Gemini"), sometimes shortened to "Jimmy" created the expressions "Jiminy Cricket!" or "Jimmy Crickets!" as less blasphemous alternatives to exclaiming "Jesus Christ!" [60]

      By the end of the 20th century the sound of chirping crickets came to represent quietude in literature, theatre and film. From this sentiment arose expressions equating "crickets" with silence altogether, particularly when a group of assembled people makes no noise. These expressions have grown from the more descriptive, "so quiet that you can hear crickets," to simply saying , "crickets" as shorthand for "complete silence." [61]

      In popular culture Edit

      Cricket characters feature in the Walt Disney animated movies Pinocchio (1940), where Jiminy Cricket becomes the title character's conscience, and in Mulan (1998), where Cri-Kee is carried in a cage as a symbol of luck, in the Asian manner. The Crickets was the name of Buddy Holly's rock and roll band [62] Holly's home town baseball team in the 1990s was called the Lubbock Crickets. [63] Cricket is the name of a US children's literary magazine founded in 1973 it uses a cast of insect characters. [64] The sound of crickets is often used in media to emphasize silence, often for comic effect after an awkward joke, in a similar manner to tumbleweed.

      What is the name of this small black insect? - Biology

      Scientific Name: Gryllus pennsylvanicus
      Common Name: Field Cricket

      (Information for this species page was gathered in part by Alicia Fitzgerald for the Spring 2006, Biology 220W at Penn State New Kensington)

      The field cricket (Gryllus pennsylvanicus) is found abundantly in a great variety of habitats (including fields and lawns, forest edges, mature forests, caves, damp basements, around plumbing, and even in outhouses) and occurs over a wide geographic range which includes most of the eastern and midwestern United States north of Florida. This wide range of habitat selection is reflective of the species&rsquo extremely broad tolerance ranges to key environmental factors. In natural habitats, G. pennsylvanicus may be found in shallow burrows and also in the matrix of dead or living vegetation above the level of the soil.

      Adult field crickets are black and brown in color and are between one half and one inch long. They have six legs, long antennae, and prominent cerci at the end of their abdomens. Their hindmost legs are very enlarged and are used by the cricket for powerful and rapid jumping. The hind wings of the field cricket are large and brightly pigmented. Not all field cricket individuals, though, are capable of flight. The non-flying G. pennsylvanicus individuals have substantially reduced flight muscle masses and may be able to more efficiently allocate energy to other biological needs (flightless female field crickets, for example, tend to be more fertile than flying females).

      Sound Production
      All field crickets are able to make the universally recognizable cricket, &ldquochirping&rdquo sounds. Males, though, are able to make the loudest and most noticeable sounds. The chirping is generated by the movement of &ldquoscrapers&rdquo found on the edge of the left forewing across a row of teeth-like structures located on the underside of the right forewing. The male field cricket generates a three note, highly trilled song which is answered by a more simplified, two note female song. The rate of chirping is directly influenced by temperature. Counting the number of chirps a male field cricket makes in 13 seconds, and then adding 40 to that number generates an approximate index of the environmental temperature (in degrees Fahrenheit).

      Field crickets are omnivorous. They eat dried organic materials, fresh plant matter, small fruits, seeds, and, at extreme need, both living and dead insects. Plants such as crabgrass, ragweed, and chicory seem to be highly favored food sources. Large populations of G. pennsylvanicus can cause significant damage to agricultural crops, and when this species enters houses (typically in the late summer and early fall) wool, cotton, silk, nylon, rubber, and leather materials may be consumed. Population explosions in this species typical come after rainfall relieving prolonged drought conditions. The crickets feed at night and spend most of the daylight hours in warm, dark refugia. A field cricket must eat its body weight or more in food every day.

      Predation, Disease and Parasites
      Field crickets are preyed upon by a wide range of predators. Most bird species (including cardinals, turkeys, blackbirds, and even some hawks) will either preferentially or opportunistically eat field crickets. Red fox, box turtles, American toads, and many other mammalian, reptilian, and amphibian predators also vigorously consume field crickets. Field crickets are also subject to many diseases and parasites. There is a virus which causes body paralysis, fungal infections and protist colonizations of the intestines, ricketsia infections, mermithid worms (nematodes), and ectoparasitic mite infestations all of which beset these animals. There are also species of parasitic wasps which sting and paralyze field crickets and then lay their eggs in the still living cricket&rsquos body. The larvae of these wasps feed upon the cricket as they grow and develop.

      Mating and Reproduction
      The male field cricket&rsquos song is his proclamation of his readiness to mate. The males make energetic and highly conspicuous song displays, and those that sing the loudest tend to attract the most females. Those singing the loudest, though, also attract the most predators! There is, then, an extremely delicate balance between the biological success of a singing male cricket (i.e. successful reproduction) and his sudden termination by a hungry predator. This balance must have many subtle contributory factors which influence the ultimate fate of each individual. Consequences of "group singing&rdquo and the possibility of non-singing &ldquolurkers&rdquo (phenomenon observed in studies of singing frogs and toads) are interesting areas for possible study in field cricket populations.

      After mating, the females lay their eggs in moist sand or soil. The cerci of females are modified into digging ovipositors. Typically, eggs are laid in groups of fifty. A female will lay up to four hundred eggs. These eggs incubate in the soil for 15 to 25 days and then hatch into nymphs. These nymphs eat very vigorously and grow rapidly undergoing eight moults as they grow into the adult forms. These nymphs and the subsequent life stages up to adult all look fundamentally alike and differ only in their relative body sizes. This type of development is called &ldquosimple metamorphosis.&rdquo The adult stage is reached in about twelve weeks, but very few individuals actually reach this level: the average life span of a field cricket is only one week.

      Ecological Role
      Field crickets are important agents in the decomposer communities of many ecosystems. They consume large quantities of often highly resistant, cellulose rich plant materials and produce fecal pellets that are easily decomposed by bacteria and fungi. Their activity, then, greatly accelerates the energy and nutrient flows in an ecosystem and provides plants with a much more abundant reservoir of highly available, essential growth factors. Field crickets also consume the seeds of many significant &ldquoweed&rdquo species thus reducing the potential of these rapidly growing, invasive plants to dominate both natural and human generated (i.e. lawns and gardens) ecosystems. Crabgrass in particular is a &ldquoweed&rdquo whose abundance can be reduced by the feeding activities of the field cricket. As mentioned above, though, when populations of the field cricket become excessively large, they can cause damage to agricultural crops. They can also do significant damage to clothing, furniture, rugs, and even rubber materials when they invade a house in large numbers.

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      What is the name of this small black insect? - Biology

      This grasshopper is well known in the southeastern USA, and elsewhere, due to its large size and widespread use in biology classrooms for dissection exercises. Also, it can be of economic importance in Florida. It is one of a few species of grasshoppers in Florida that occurs in large enough numbers to cause serious damage to citrus, vegetable crops, and landscape ornamentals. Unfortunately, the scientific community uses two different scientific names for the same species, and Romalea microptera (Palisot de Beauvois) is also called Romalea guttata (Houttuyn). Technically, the latter is probably the correct name (Kevan 1980), but because the former designation was used for many years, this proposed &lsquocorrection&rsquo has introduced unnecessary confusion (Cohn 1999), so most scientists continue to call it Romalea microptera.

      The eastern lubber is quite clumsy and slow in movement and mostly travels by walking and crawling feebly over the substrate. The &ldquolubber&rdquo designation is interesting because it aptly describes this grasshopper. &ldquoLubber&rdquo is derived from an old English word &ldquolobre&rdquo which means lazy or clumsy. This term has come to mean a big, clumsy, and stupid person, also known as a lout or lummox. In modern times, it is normally used only by seafarers, who term novices &ldquolandlubbers&rdquo. Eastern lubber is one of only four species in the family Romaleidae found north of Mexico, but there are many other species in South America (Rehn and Grant 1961), and many are winged and agile, so although some other species in this family may be called lubbers, the &ldquolubber&rdquo designation is not appropriate for the entire family.

      Distribution (Back to Top)

      The eastern lubber grasshopper is limited to the southeastern region of the United States. It is found from the North Carolina south through South Carolina, Georgia and Florida, and west through Alabama, Mississippi, and Louisiana to central Texas (Capinera et al. 2004).

      Figure 1. Distribution of Romalea microptera, the eastern lubber grasshopper.

      Description (Back to Top)

      Eastern lubber grasshopper is surely the most distinctive grasshopper species found in the southeastern USA. Adults are colorful, but the color pattern varies. Often the adult eastern lubber is mostly yellow or tawny, with black on the distal portion of the antennae, on the pronotum, and on the abdominal segments. The forewings extend two-thirds to three-fourths the length of the abdomen. The hind wings are short and incapable of providing lift for flight. The forewings tend to be pink or rose in color centrally whereas the hind wings are entirely rose in color. Darker forms of this species also exist, wherein the yellow color becomes the minor rather than the major color component, and in northern Florida a predominantly black form is sometimes found. Adults attain a large size, males measuring 43-55 mm in length and females often measuring 50-70 mm, sometimes 90 mm. Not only is this large, heavy-bodied grasshopper unable to fly, but is poor at leaping as well, so mostly it is observed walking. However, it is a good climber, and often climbs trees to feed on juvenile foliage at the tips of branches.

      Both sexes stridulate (make noise) by rubbing the forewing against the hind wing. When alarmed, lubbers will spread their wings, hiss, and secrete foul-smelling froth from their spiracles. They can expel a fine spray of toxic chemicals for a distance of 15 cm. The chemical discharge from the tracheal system is believed to be an anti-predator defense, and to consist of chemicals both synthesized and sequestered from the diet. The variation in toxins assimilated from the diet make it difficult for predators to adapt to the toxins (Chapman and Joern 1990). Many vertebrate, but not invertebrate, predators are affected (Jones et al. 1987, 1989 Whitman et al. 1992). Their bright color pattern is believed to be a warning to vertebrate predators that lubbers are not palatable. Their tendency to aggregate and to climb vegetation, especially at night, is a component of their defensive behavior.

      Eggs. The eggs of lubber grasshoppers are yellowish or brown in color. They are elongate elliptical in shape and measure about 9.5 mm in length and 2.5 mm in width. They are laid in neatly arranged clusters, or pods, which consist of rows of eggs positioned parallel to one another, and held together by a secretion. Normally there are 30-50 eggs in each pod. Ovipositing females are reported to prefer mixed broadleaf tree-pine habitats with intermediate soil moisture levels, avoiding both lowland, moist, compact soil and upland, dry, sandy soil (Watson 1941, Kuitert and Connin 1953). The female deposits the pod in the soil at a depth of 3-5 cm and closes the oviposition hole with a frothy secretion or plug. The plug allows the young grasshoppers easy access to the soil surface when they hatch. Egg pods tend to be clustered, with females preferentially ovipositing where eggs have already been deposited (Stauffer et al. 1998). According to Hunter-Jones (1967), females each produce 3-5 egg clusters in structures called pods. The pod is not much more than tightly packed eggs surrounded by rigid, frothy material, with most of the froth deposited at the tip of the pod closest to the surface. The froth allows an easy exit for the young hoppers as they can readily wiggle through this as they hatch. The interval between egg pod production by a female is about 2 weeks. Hunter-Jones reported egg production of 30-80 eggs per pod, averaging about 60 eggs per pod. Egg production was greater under solitary than crowded conditions. On the other hand, under field conditions Stauffer and Whitman (2007) reported egg production of 25-50 eggs per pod, with only 1-3 pods per female. Thus, egg production in the laboratory was greater than in the field. The eggs require a cool period (e.g., 20°C for 3 months) but then will hatch when exposed to warmer temperatures. Typically, egg hatch occurs in the morning.

      Nymphs. The immature eastern lubber grasshopper differs dramatically in appearance from the adults (Capinera et al. 1999, 2001). Their color pattern is so different from the adult stage that the nymphs commonly are mistaken for a different species than the adult form. Nymphs (immature grasshoppers) typically are almost completely black, but with a distinctive yellow, orange, or red stripe located dorsally (though occasionally they are reddish brown). The hopper&rsquos face, edge of the pronotum, and abdominal segments also may contain reddish accents. Often the reddish accents change to yellow over the course of development. When they first molt, the young hoppers may be brownish, but they soon darken to black.

      As they mature, the nymphs change slightly in appearance the instar can be determined by examination of the developing wings. Normally there are 5 instars, though occasionally 6 instars occur. The early instars can be distinguished by a combination of body size, the number of antennal segments, and the form of the developing wings. The nymphs measure about 10-12, 16-20, 22-25, 30-40, and 35-45 mm in length during instars 1-5, respectively. Antennal segments, which can be difficult to distinguish even with magnification, number 12, 14-16, 16-18, 20, and 20 segments per antenna during instars 1-5, respectively. The shapes of the plates immediately behind the pronotum (the future wings) change slightly with each molt. During the first instar the ventral surface is broadly rounded during the second instar the ventral edges begin to narrow slightly and point slightly posteriorly, and also acquire slight indication of venation during the third instar the ventral edges of the plates are markedly elongate, point strongly posteriorly, and the veins are pronounced. At the molt to the fourth instar the orientation of the small, developing wings shifts from pointing downward to pointing upward and posteriorly. In instar 4 the small forewings and hind wings are discrete and do not overlap, though the forewings may be completely or partly hidden beneath the pronotum. In instar 5, the slightly larger wings overlap, appearing to be but a single pair of wings. Even in the fifth instar, however, the wing buds do not cover the tympanum. In adults, however, the wings overlap and cover the tympanum, extending posteriorly to cover 3-4 abdominal segments.

      Young nymphs are highly gregarious, and remain gregarious through most of the nymphal period, though the intensity dissipates with time. Especially at night, they tend to aggregate, and may climb vegetation to rest for the evening.

      Figure 2. Young lubber, Romalea microptera (Beauvois), instar one. The segments above the second and third legs bear wing buds, though at this stage they are hardly visible. Photograph by Lyle J. Buss, University of Florida.

      Figure 3. Young lubber, Romalea microptera (Beauvois), instar two. The beginnings of the wing veins can be seen now. Photograph by Lyle J. Buss, University of Florida.

      Figure 4. Young lubber, Romalea microptera (Beauvois), instar three. Now you can better see the developing veins and a slight backward (posterior) extension of the wing buds. Photograph by Lyle J. Buss, University of Florida.

      Figure 5. Young lubber, Romalea microptera (Beauvois), instar four. Now you can see something that actually looks like a wing, although it is quite small. Note that it is pointed upward (dorsally). Also note its length relative to the oval, reddish-colored tympanum on the first abdominal segment. Photograph by Lyle J. Buss, University of Florida.

      Figure 6. Young lubber, Romalea microptera (Beauvois), instar five. The wing is longer now, extending further posteriorly relative to the tympanum. Next it will molt to the adult, though you cannot tell from the nymph what color form will be assumed by the adult. Photograph by John Capinera, University of Florida.

      Figure 7. Lubber nymph Romalea microptera (Beauvois), molting, leaving behind its old (darker) body covering. Photograph by Lyle J. Buss, University of Florida.

      Adults. Adult males and females are usually 6.0 and 8.0 cm. long, respectively. The body is quite robust while the legs remain relatively slender. The general color of adults is dull yellow with varying degrees of black spots and markings. The front pair of wings (tegmina) are yellow with numerous scattered black dots, while the hind wings when exposed reveal a bright red/rose coloration with a black border. The color of adult lubbers also varies throughout most of the insect&rsquos range. One phase is nearly entirely black (melanic) with a few marks of yellowish. The adults of this melanic phase seem to resemble the nymph, but only in color. Individuals in the same geographic area may be yellowish, melanic, or somewhere in between. However, despite their different appearances, these different-appearing grasshoppers are the same species and will mate successfully.

      Figure 8. Adult eastern lubber grasshopper, Romalea microptera (Beauvois), light color phase. Photograph by John Capinera, University of Florida.

      The color of adult lubbers also varies throughout most of the insect's range. One phase is nearly entirely black with a few marks of yellowish tawny. The adults of this phase seem to resemble the nymph. However, the different phases are indeed the same species.

      Figure 9. Adult eastern lubber grasshopper, Romalea microptera (Beauvois), intermediate color phase. Photograph by John Capinera, University of Florida.

      Figure 10. Adult eastern lubber grasshopper, Romalea microptera (Beauvois), dark color phase. Photograph by John Capinera, University of Florida.

      Figure 11. Black color form of adult eastern lubber grasshopper, Romalea microptera (Beauvois). Photograph by Lyle J. Buss, University of Florida.

      Life Cycle (Back to Top)

      There is one generation per year, with the egg stage overwintering. Apparently there is not an obligatory diapause (required period of dormancy) in the egg stage they simply have a long period of development (about 200 days) when held at low temperatures (Hunter-Jones 1967). These grasshoppers are long-lived, and either nymphs or adults are present throughout most of the year in the southern portions of Florida. In northern Florida and along the Gulf Coast they may be found from March-April to about October-November. In Florida, the highest number of adults can be observed during the months of July and August. Eggs are produced about a month after emergence of the adults. Duration of the egg stage is 6-8 months.

      After mating, females will begin laying eggs during the summer months. The male usually guards the ovipositing female, sometimes for more than a day. The timing of oviposition is highly variable, but ovipositing females select open, sunny areas of higher elevation, then use the tip of the abdomen to dig a small hole into a suitable patch of soil. Usually at a shallow depth, but sometimes up to a depth of about 5 cm, she will deposit her eggs within a light foamy froth. These eggs will remain in the soil through late fall and winter and then begin hatching in Spring. The young grasshoppers crawl up out of the soil upon hatching and congregate near suitable food sources. Lubbers are often found in damp or wet habitats, but seek drier sites for egg-laying.

      Figure 12. Mating adults of two color forms. Photograph by Lyle J. Buss, University of Florida.

      Populations cycle up and down, possibly due to the action of parasites. The tachinid fly Anisia serotina (Reinhard) attains high levels of parasitism, sometimes 60-90% (Lamb et al. 1999). We have also found the sarcophagids Blaesoxipha opifera (Coquillett) and Blaesoxipha hunter (Hough) parasitizing this grasshopper, sometimes at high incidences of parasitism (unpublished identified by G.A. Dahlem, Northern Kentucky University). Pathogens known from Romalea microptera include Boliviana floridensis (Stauffer and Whitman 2007) and Encephalitozoon romaleae (Lange et al. 2009).

      Overall, the natural enemies of lubber grasshoppers are poorly documented. Vertebrate predators such as birds and lizards learn to avoid these insects due to the production of toxic secretions by the adult hoppers, though this is not absolute (Chapman and Joern 1990). Naïve vertebrates often gag, regurgitate, and sometimes die following consumption of lubbers (Yousef and Whitman 1992). However, loggerhead shrikes, Lanius ludovicianus Linnaeus, capture and cache lubbers by impaling them on thorns and the barbs of barbed wire fence. After 1-2 days the toxins degrade and the dead lubbers become edible to the shrikes (Yousef and Whitman 1992). In addition to parasitic flies, nematodes have been reported from lubbers, and it is possible to infect lubbers experimentally with the grasshopper-infecting nematode Mermis nigrescens.

      Figure 13. One of the species of parasitic flies (Blaesoxipha hunteri) that affects lubbers. The larvae develop within the nymphs, killing their hosts when they emerge. Photograph by Lyle J. Buss, University of Florida.

      Habitat and Hosts (Back to Top)

      Eastern lubber grasshopper has a broad host range. At least 100 species from 38 plant families containing shrubs, herbs, broadleaf weeds, and grasses are reportedly eaten (Whitman 1988), though their mouthparts are best adapted for feeding on forbs (broad-leaf plants), not grasses (Squitier and Capinera 2002). Among the plants observed to be eaten are pokeweed, Phytolaca americana tread-softly, Cnidoscolus stimmulosus pickerel weed, Pontederia cordata lizard&rsquos tail, Saururus sp. sedge, Cyperus and arrowhead, Sagittaria spp. Though its preferred habitat seems to be low, wet areas in pastures and woods and along ditches, lubbers disperse long distances during the nymphal period. They are gregarious and flightless, their migrations sometimes bringing large numbers into contact with crops where they damage vegetables, some field crops (peanuts, cowpeas, corn), fruit trees (citrus, figs, peach), and ornamental plants (Kuitert and Connin 1953).

      The nature of polyphagous insects it to accept at least small amounts of many plants, a strategy that assures they will not starve. However, not all plants are accepted equally. For example, although lubbers display broad preference among vegetable crops, some plants such as pea, lettuce, kale, beans, and cabbage are relatively preferred whereas eggplant, tomato, pepper, celery, okra, fennel, and sweet corn are less preferred (Capinera 2014). They commonly defoliate amaryllis, Amazon lily, crinum, narcissus, and related plants (family Amaryllidaceae) in flower gardens. Among ornamental plants, they also will feed readily on oleander, butterfly weed, peregrina, Mexican petunia, and lantana (Capinera 2014). Preferred weeds include painted leaf, Poinsettia cyathophora tread softly, Cnidoscolus stimulosus chamber bitter, Phyllanthus urinaria Florida beggarweed, Desmodium tortuosum Old-world diamond-flower, Oldenlandia corymbosa Florida pusley, Richardia scabra and smooth crabgrass, Digitaria ischaemum (Capinera 2014).

      Damage (Back to Top)

      Lubber grasshoppers are defoliators, consuming the leaf tissue of numerous plants. They climb readily, and because they are gregarious they can completely strip foliage from plants. More commonly, however, they will eat irregular holes in vegetation and then move on to another leaf or plant. Lubber grasshoppers are not as damaging as their size might suggest they consume less food than smaller grasshoppers (Griffiths and Thompson 1952). Damage is commonly associated with areas that support weeds or semi-aquatic plants such as irrigation and drainage ditches, end edges of ponds. Grasshoppers developing initially in such areas will disperse to crops and residential areas, where they cause damage. Thus, as is the case with many grasshoppers, monitoring and treatment of areas where nymphal development occurs is recommended to prevent damage to economically important plants. Also helpful is to keep vegetation mowed, as short vegetation is less supportive of grasshoppers.

      Management (Back to Top)

      Management of these insects tends to rely on capture (physical removal) when only a few hoppers are present. When there are too many to be controlled by hand-picking, insecticides can be applied. Insecticides can be applied to the foliage or directly to the grasshopper. However, due to their large size and ability to detoxify natural toxins associated with food plants, they often prove difficult to kill, especially by spraying the foliage. Insecticides that will kill lubber grasshoppers include carbaryl, bifenthrin, cyhalothrin, permethrin, esfenvalerate, and spinosad (note: these are the technical names of insecticides, not the trade names these names appear in the &lsquoingredients&rsquo section of the label). Spinosad is particularly interesting because it is a biologically based, relatively safe product unfortunately, it is rather slow acting on grasshoppers, so it may take a few days to see results of treatment. Insecticide treatment is more effective for young grasshoppers, which may necessitate scouting for hoppers in weedy areas, and treatment of them before they move into gardens and crops. An alternative is to treat the margins of cropland, perhaps the initial 1-20 meters, so that as hoppers disperse through the crop from the edges they encounter treated vegetation and perish after sampling it. Because they are dispersive, and may continue to invade an area even after it is treated with insecticide, it can be difficult to provide protection to plants without diligent monitoring and retreatment.

      If insecticides are used, be sure to apply them according to the directions on the label of the container. Especially if insecticides are applied to food crops or near water, it is important to follow directions. Most of the insecticides listed above are toxic to fish.

      Insecticide-containing baits are sometimes formulated for grasshopper control normally bran is the substrate to which the insecticide is applied, and it is sprinkled on the soil surface near the plants being protected. Lubbers will accept such baits, and insects are readily killed if they ingest the toxicant and bait. However, this tactic works better when the bait is applied to the vicinity of less preferred plants, as the hoppers will tend to eat the favored host plants in preference to the treated bait (Barbara and Capinera 2003, Capinera 2014). Treatment of field margins with baits can help to reduce crop damage from immature and flightless hoppers such as lubbers, though treatment of field margins is less effective with grasshopper species that are strong fliers such as Schistocerca americana (Drury).

      Figure 14. Young nymphs of the eastern lubber grasshopper, Romalea microptera (Beauvois), clustered on a citrus reset (young citrus tree). Photograph by John Capinera, University of Florida.

      Selected References (Back to Top)

      • Barbara, KA, Capinera JL. 2003. Development of a toxic bait for control of eastern lubber grasshopper (Orthoptera: Acrididae). Journal of Economic Entomology 96: 584-591.
      • Blatchley WS. 1920. Orthoptera of Northeastern America. Nature Publishing Company. Indianapolis, Indiana. p. 304-307.
      • Capinera JL. 2014. Host plant selection by Romalea microptera (Orthoptera: Romaleidae). Florida Entomologist 97: 38-49.
      • Capinera JL, Scherer CW, Squitier JM. 1999. Grasshoppers of Florida.
      • Capinera JL, Scherer CW, Squitier JM. 2001. Grasshoppers of Florida. University Press of Florida. 143 pp.
      • Capinera JL, Scott RD, and Walker TJ. 2004. Field guide to the grasshoppers, katydids, and crickets of the United States. Cornell University Press, Ithaca. 249 pp.
      • Chapman RF, Joern A. (eds.) 1990. Biology of Grasshoppers. John Wiley , New York.
      • Cohn TJ. 1999. Current controversies: nomenclatorial stability and the law of priority. Metaleptea 19: 8-9.
      • Griffiths JT, Thompson WL. 1952. Grasshoppers in citrus groves. University of Florida Agricultural Experiment Station Bulletin no. 496.
      • Helfer JR. 1953. How to Know the Grasshoppers, Cockroaches and Their Allies. WM.C. Brown Company Publishers. Dubuque, Iowa. p. 100-101.
      • Hunter-Jones, P. 1967. The life-history of the eastern lubber grasshopper, Romalea microptera (Beauvois), (Orthoptera: Acrididae) under laboratory conditions. Proceedings of the Royal Entomological Society, London (A) 42: 18-24.
      • Johny J, Whitman DW. 2005. Description and laboratory biology of Boliviana floridensisn. sp (Apicomplexa: Eugregarinida) parasitizing the eastern lobster grasshopper, Romalea microptera (Ortoptera: Romalidae) from Florida, USA. Comparative Parasitology 72: 150-156, doi: 10.1654/4164.
      • Jones CG, Hess TA, Whitman DW, Silk PJ, Blum MS. 1987. Effects of diet breadth on autogenous chemical defense of a generalist grasshopper. Journal of Chemical Ecology 13: 283-297.
      • Jones CG, Whitman DW, Compton SJ, Silk PJ, Blum MS. 1989. Reduction in diet breadth results in sequestration of plant chemicals and increases efficacy of chemical defense in a generalist grasshopper. Journal of Chemical Ecology 15: 1811-1822.
      • Kevan DK McE. 1080. Romalea guttata (Houttuyn), name change for well-known &ldquoeastern lubber grasshopper&rdquo (Orthoptera: Romaleidae). Entomological News 91: 139-140.
      • Kuitert LC, Connin RV. 1953. Grasshoppers and their control. University of Florida Agricultural Experiment Station Bulletin no. 516.
      • Lamb MA, Otto DJ, Whitman DW. 1999. Parasitism of eastern lubber grasshopper by Anisia serotina (Diptera: Tachinidae) in Florida. Florida Entomologist 82: 365-371.
      • Lange CE, Johny S, Baker MD, Whitman DW, Solter LF. 2009. A new Encephalitozoon species (Microsporida) isolated from the lubber grasshopper, Romalea microptera (Beauvois) (Orthoptera: Romaleidae). Journal of Parasitology 95: 976-986.
      • Rehn JAG, Grant Jr HJ. 1961. A Monograph of the Orthoptera of North America (North of Mexico). Monographs of the Academy of Natural Sciences of Philadelphia. No. 12. Vol. 1. p.231-240. Wickersham Printing Company. Lancaster, Pennsylvania.
      • Squitier JM, Capinera JL. 2002. Host selection by grasshoppers (Orthoptera: Acrididae) inhabiting semi-aquatic environments. Florida Entomologist 85: 336-340.
      • Stauffer TW, Hegrenes SG, Whitman, DW. 1998. A laboratory study of oviposition site preference in the lubber grasshopper, Romalea guttata (Houttuyn). Journal of Orthoptera Research 7: 217-221.
      • Stauffer TW, Whitman DW. 2007. Divergent oviposition behaviors in a desert vs a marsh grasshopper. Journal of Orthoptera Research 16: 103-114.
      • Watson JR. 1941. Migrations and food preferences of the lubberly locust. Florida Entomologist 24: 40-42.
      • Whitman DW. 1988. Allelochemical interactions among plants, herbivores, and their predators, pp. 11-64 In Barbosa P and Letournearu D (eds.) Novel Aspects of Insect-Plant Interactions. J. Wiley, New York.
      • Whitman DW, Billen JPJ, Alsop D, Blum MS. 1991. Anatomy, ultrastruicture, and functional morphology of the metathoracic tracheal defensive glands of the grasshopper Romalea guttata. Canadian Journal of Zoology 69: 2100-2108.
      • Whitman DW, Jones CG, Blum MS. 1992. Defensive secretion in lubber grasshoppers (Orthoptera: Romalidae): influence of age, sex, diet, and discharge frequency. Annals of the Entomological Society of America 85: 96-102.
      • Yousef R, Whitman D. 1992. Predator exaptations and defensive adaptations in evolutionary balance: no defence is perfect. Evolutionary Ecology 6: 527-536.

      Authors: John Capinera and Clay Scherer, University of Florida
      Photographs: John Capinera and Lyle J. Buss, University of Florida
      Web Design: Don Wasik, Jane Medley
      Publication Number: EENY-6
      Publication Date: October 1996. Latest revision: November 2016.

      An Equal Opportunity Institution
      Featured Creatures Editor and Coordinator: Dr. Elena Rhodes, University of Florida

      Prevention Summaries

      Many of the pests discussed above had similar prevention tactics to keep your home safe. The top ten pest prevention tips include:

      • Secure garbage bins and store them away from your home
      • Store firewood twenty feet from your home and handle with gloves
      • Screen and secure eaves, crawl spaces, and more
      • Screen and/or seal windows and doors, check weather stripping
      • Ensure proper drainage and remove standing water for fixtures like bird baths, change out the water once a week
      • Minimize clutter and dust in your home
      • Do not leave food open and unattended
      • Wipe up spills and crumbs as soon as possible
      • Ensure pets are adequately protected with veterinarian-recommended solutions

      Nature’s Wisdom

      Not everything requires harsh chemicals, and more companies are finding ways to treat pest issues naturally or relocate mice and other rodents. Don’t be afraid to try natural remedies, taking gradual steps in your fight against invaders. Many people like to try with prevention, then natural or ‘soft’ remedies, lighter chemicals, and then heavier pesticides.

      However, it’s important to remember that even some natural remedies can be dangerous for members of your household, especially those with sensitive allergies and medical conditions and our feathered and fluffy family members. Due diligence is key when researching tactics to keep pests out but keep our families and pets safe.

      Strength in Numbers

      Remember, no homeowner faces this battle alone. Unfortunately, the internet is filled with both helpful information and misguided attempts to educate about pest control.

      Researching potential strategies is always wise, and you can often find likeminded homeowners on social media sites who have both successfully and unsuccessfully dealt with pest situations similar to yours.

      When all else fails, professional exterminators or pest control services may be your best bet. Pest control companies are committed to keeping their communities safe and healthy, so you can always turn to the professionals for help. Any legitimate company will have highly-trained professionals that can accurately assess your situation, offer a range of solutions, and help you with prevention once the original threat is eliminated.

      Watch the video: Seeing little black bugs? Heres what they are (January 2022).