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Species Identification - small insect

Species Identification - small insect


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I found a few of these walking in the middle of the night (about 3 AM) on a wall in my room near the floor. They are at most a few millimetres long (no more than 2 I would say). I am living in Poland, in a block of flats.

EDIT: Since I first wrote, I have seen few more - very small - less then 1 millimetre long. They seemed to be coming from behind the baseboard. This time I've seen them during daytime.


I think @fileunderwater is definitely right, these are psocids (of the family Psocoptera, AKA booklice).

Specifically, the body shape looks very similar to that of Dorypteryx domestica or some closely related species.

See here for an additional photo.

According to this site, the species is present in many countries including Poland.

The source cited at this site (Kučerová 1998) mentions that this species can be brachypterous (i.e., having very reduced wings. I can't tell from the quality of the photo whether your specimens has tiny wings or not. Your top picture looks like it might.

Citations:

Kučerová, Z. (1998). Wing polymorphism in Dorypteryx domestica (Smithers) (Psocoptera: syllipsocidae). Insect Systematics & Evolution, 29(4): 451-57.


It could be booklice (Psocoptera), but the picture is a bit unclear. Booklice are harmless, but could indicate moist conditions and/or mold in the building (see first link for more on their biology). Termites are generally at least twice as large as booklice (>4mm), and the size you indice rather points to booklice.

Here is an example of what booklice look like:

(picture from Penn State University: Dept. of Entomology)


Artificial intelligence puts focus on the life of insects

Insect monitoring cameras in a remote area in East Greenland. Credit: Toke T. Høye

Scientists are combining artificial intelligence and advanced computer technology with biological know how to identify insects with supernatural speed. This opens up new possibilities for describing unknown species and for tracking the life of insects across space and time

Insects are the most diverse group of animals on Earth and only a small fraction of these have been found and formally described. In fact, there are so many species that discovering all of them in the near future is unlikely.

This enormous diversity among insects also means that they have very different life histories and roles in the ecosystems.

For instance, a hoverfly in Greenland lives a very different life than a mantid in the Brazilian rainforest. But even within each of these two groups, numerous species exist each with their own special characteristics and ecological roles.

To examine the biology of each species and its interactions with other species, it is necessary to catch, identify, and count a lot of insects. It goes without saying that this is a very time-consuming process, which to a large degree, has constrained the ability of scientists to gain insights into how external factors shape the life of insects.

A new study published in the Proceedings of the National Academy of Sciences shows how advanced computer technology and artificial intelligence quickly and efficiently can identify and count insects. It is a huge step forward for the scientists to be able to understand how this important group of animals changes through time—for example in response to loss of habitat and climate change.

Close up insect (hoverfly) visiting flower. Credit: Hjalte M.R. Mann

"With the help of advanced camera technology, we can now collect millions of photos at our field sites. When we, at the same time, teach the computer to tell the different species apart, the computer can quickly identify the different species in the images and count how many it found of each of them. It is a game-changer compared to having a person with binoculars in the field or in front of the microscope in the lab who manually identifies and counts the animals," explains senior scientist Toke T. Høye from Department of Bioscience and Arctic Research Centre at Aarhus University, who headed the new study. The international team behind the study included biologists, statisticians, and mechanical, electrical and software engineers.

The methods described in the paper go by the umbrella term deep learning and are forms of artificial intelligence mostly used in other areas of research such as in the development of driverless cars. But now the researchers have demonstrated how the technology can be an alternative to the laborious task of manually observing insects in their natural environment as well as the tasks of sorting and identifying insect samples.

"We can use the deep learning to find the needle in the hay stack so to speak—the specimen of a rare or undescribed species among all the specimens of widespread and common species. In the future, all the trivial work can be done by the computer and we can focus on the most demanding tasks, such as describing new species, which until now was unknown to the computer, and to interpret the wealth of new results we will have" explains Toke T. Høye.

And there is indeed many tasks ahead, when it comes to research on insects and other invertebrates, called entomology. One thing is the lack of good databases to compare unknown species to those which have already been described, but also because a proportionally larger share of researchers concentrate on well-known species like birds and mammals. With deep learning, the researchers expect to be able to rapidly advance knowledge about insects considerably.

Automatic moth trap with moths detected by computer vision. Credit: Toke T. Høye and Kim Bjerge

Long time series are necessary

To understand how insect populations change through time, observations need to be made in the same place and in the same way over a long time. It is necessary with long time series of data.

Some species become more numerous and others more rare, but to understand the mechanisms that causes these changes, it is critical that the same observations are made year after year.

An easy method is to mount cameras in the same location and take pictures of the same local area. For instance, cameras can take a picture every minute. This will give piles of data, which over the years can inform about how insects respond to warmer climates or to the changes caused by land management. Such data can become an important tool to ensure a proper balance between human use and protection of natural resources.

"There are still challenges ahead before these new methods can become widely available, but our study points to a number of results from other research disciplines, which can help solve the challenges for entomology. Here, a close interdisciplinary collaboration among biologists and engineers is critical," says Høye.


Contents

Diptera occur all over the world except in regions with permanent ice-cover. They are found in most land biomes (all 14 WWF major habitat types) including deserts and the tundra. Insects are the most diverse group of Arctic animals (about 3,300species), of which about 50% are Diptera. Palearctic habitats include meadows, prairies, mountain passes, forests, desert oases, seashores, sandy beaches, coastal lagoons, lakes, streams and rivers, bogs, fens, areas (including waters polluted by rotting waste, industrial emissions), urban areas, cattle, horse and poultry farms.

Cave Diptera Edit

see also List of fauna of Batu Caves The Diptera fauna of caves includes species in Sphaeroceridae, Mycetophilidae, Psychodidae, Phoridae, Tipulidae, Trichoceridae, Heleomyzidae, Mycetophilidae and Culicidae. The main sources of food for cave Diptera are other insects, carrion and guano. Most are perhaps only troglophiles. [1]

Desert Diptera Edit

Desert diptera include specialised species of Psychodidae, Nemestrinidae, Therevidae, Scenopinidae and Bombyliidae. These groups are most diverse where dry sandy soils provide a suitable habitat for the larvae.

Freshwater Diptera Edit

Larval stages of Diptera can be found in almost any aquatic or semiaquatic habitat They form an important fraction of the macro zoobenthos of most freshwater ecosystems. families are Chironomidae (very significant), Stratiomyidae, Ephydridae, Dixidae and Tipulidae.

Food habits of most species are largely unknown but broad statements may be made. Diptera are important pollinators and plant pests.

Detritivores Edit

Many Diptera are detritivores. Typical are Dryomyza anilis and, notably, Musca domestica.

Flower feeders Edit

Many adult Brachycera feed on flowers notably Syrphidae which obtain all their protein requirements by feeding on pollen. The Calyptratae exhibit flower feeding in all families except Hippoboscidae Nycterebidae and Glossinidae and in the Acalyptratae the Conopidae are well known flower feeders. Other flower feeding Brachycerous families are Empididae, Stratiomyidae and the Acroceridae like various members of the Nemestrinidae, Bombyliidae and Tabanidae are nectar feeders with exceptionally long proboscises, sometimes longer than the entire bodily length of the insect. Flower feeding Nematocera include Bibionidae and some species in Tipulidae and other families. [2] [3]

Bombylius- note the proboscis

Predators Edit

Adult Asilidae, Empididae and Scathophagidae feed on other insects, including smaller Diptera, Dolichopodidae and some Ephydridae feed on a variety of animal prey.

Tachypeza nubila: Empididae with prey

Dysmachus fuscipennis: Asilidae with beetle prey

Cordilura: Scathophagidae hunting

Ochthera an Ephydridae with raptorial forelegs

Both male and female mosquitoes feed on nectar and plant juices, but in many species the mouthparts of the females are adapted for piercing the skin of animal hosts and feed on blood as ectoparasites. The most important function of blood meals is to obtain proteins as materials for egg production. For females to risk their lives on blood sucking while males abstain, is not a strategy limited to the mosquitoes it also occurs in some other families, such as the Tabanidae. Most female horse flies feed on mammal blood, but some species are known to feed on birds, amphibians or reptiles. Other bloodfeeding Diptera are Ceratopogonidae Phlebotominae Hippoboscidae, Hydrotaea and Philornis downsi (Muscidae), Spaniopsis and Symphoromyia Rhagionidae. There are no known acalyptrates that are obligate blood-feeders.

Haemotopota pluvialis feeding

Phlebotomus pappatasi after a blood meal

Sheep ked, Melophagus ovinus, a highly specialised blood-feeding dipteran ectoparasite

Larvae Edit

The larvae of Diptera feed on a diverse array of nutrients often these are different from those of adults, for instance the larvae of Syrphidae in which family the adults are flower-feeding are saprotrophs, eating decaying plant or animal matter, or insectivores, eating aphids, thrips, and other plant-sucking insects.

Larval Diptera feed in leaf-litter, in leaves, stems, roots, flower and seed heads of plants, moss, fungi, rotting wood, rotting fruit or other organic matter such as slime, flowing sap, and rotting cacti, carrion, dung, detritus in mammal bird or wasp nests, fine organic material including insect frass and micro-organisms. Many Diptera larvae are predatory, sometimes on the larvae of other Diptera.

Many Agromyzidae are leaf miners. Some Tephritidae are leaf miners or gall formers. The larvae of all Oestridae oestrids are obligate parasites of mammals. (Oestridae include the highest proportion of species whose larvae live as obligate parasites within the bodies of mammals. Most other species prone to cause myiasis are members of related families, such as the Calliphoridae. There are roughly 150 known species worldwide.) Tachinidae larvae are parasitic on other insects. Conopidae larvae are endoparasites of bees and wasps or of cockroaches and calyptrate Diptera, Pyrgotidae larvae are endoparasites of adult scarab beetles. Sciomyzidae larvae are exclusively associated with freshwater and terrestrial snails, or slugs. They feed on snails as predators, parasitoids, or scavengers. Females search out snails for oviposition. Known Odiniidae larvae live in the tunnels of wood-boring larvae of Coleoptera, Lepidoptera, and other Diptera and function as scavengers or predators of the host larvae. Oedoparena larvae feed on barnacles. The larvae of Acroceridae and some Bombyliidae are hypermetamorphic.


Turfgrass Insects

This publication provides turfgrass management professionals and property owners with information to help them 1) properly identify the most common billbug species associated with turfgrass in Indiana and adjacent states, 2) understand billbug biology, 3) recognize billbug damage and 4) formulate safe and effective billbug management strategies. For information on turfgrass identification, weed, disease and fertility management, visit the Purdue Turfgrass Science Website ( http://www.turf.purdue.edu) or call Purdue Extension (765-494-8491).

BILLBUG SPECIES ASSOCIATED WITH TURFGRASS IN THE MIDWEST

Billbugs represent a complex of weevils in the genus Sphenophorus that are increasingly being recognized as major pests of managed turfgrass around the world. The larvae of these insects damage a variety of warm- and cool-season grasses by feeding on or inside the stems, crowns, roots, stolons, and rhizomes. There are at least 4 species of billbugs associated with turfgrass in the Midwest. These include the bluegrass billbug Sphenophorus parvulus Gyllenhal (Fig. 1A), the hunting billbug Spenophorus venatus Say (Fig. 1B), the lesser billbug Sphenophorus minimus Hart (Fig. 1C) and the unequal billbug Sphenophorus inaequalis Say (Fig. 1D). The distribution of these four species overlaps significantly. It is not uncommon to find mixed populations of two or more species at a single location. In the Midwest, bluegrass billbug is the most prevalent species infesting cool-season turfgrasses such as Kentucky bluegrass, perennial ryegrass, fine fescue, and tall fescue. Hunting billbug is the most frequently encountered billbug pest of warm-season turfgrasses such as zoysiagrass and Bermudagrass. Lesser and unequal billbugs may infest warm-or cool-season turfgrasses, but infestations of these two species usually occur at comparatively low densities.

Figure 1a. Adults of four billbug species associated with turfgrass in the Midwest. (A) bluegrass billbug.

Figure 1b. Adults of four billbug species associated with turfgrass in the Midwest. (B) hunting billbug.

Figure 1c. Adults of four billbug species associated with turfgrass in the Midwest. (C) lesser billbug.

Figure 1d. Adults of four billbug species associated with turfgrass in the Midwest. (D) unequal billbug.

IDENTIFICATION AND SEASONAL BIOLOGY

Bluegrass Billbug

Like all billbug species, bluegrass billbug adults can be easily recognized by the presence of a long snout on the front of the head. They are 7-8 mm long and gray to black in color, but they are sometimes coated with soil making them appear brown or beige. Upon closer examination, the region directly behind the head (pronotum) is adorned with small, evenly spaced punctures of uniform size. The rest of the body is covered with alternating rows of small and large punctures that give them a striped appearance. The larvae are white, legless, soil- and crown-inhabiting insects with a chestnut colored head (Fig. 2).

Figure 2. Bluegrass billbug larva and damaged crown of a Kentucky bluegrass plant.

Adult bluegrass billbugs spend the winter in the thatch, cracks and crevices in the soil, plant debris or around structures such as sidewalks, driveways and buildings. They become active in April or May as soil temperatures at the surface warm to about 65°F (Fig 3.). Adults feed by chewing holes in grass stems, but cause no significant damage to the turf. Adult females insert eggs into the feeding holes they create (Fig. 4) and these eggs hatch into small larvae. Larvae bore inside the stems until they deplete the resources within (Fig. 5). By mid-June, larger larvae begin feeding on plant crowns just below the soil surface. Feeding by these larger larvae causes significant damage and may kill plants. By mid-July, larvae start to pupate. New adults begin to appear by August. These adults generally feed for a short time and find a suitable place to overwinter, but some may lay eggs resulting in a partial second generation of larvae. Larvae of this second generation do not survive the winter. Activity and development of bluegrass billbug can be tracked online using the Growing Degree-Day Tracker: http://www.gddtracker.net/?zip=49001&offset=0&model=12.

Figure 3. Seasonal biology of the bluegrass billbug and windows of opportunity for three different management strategies using chemical insecticides: (1) adult preventive, (2) larval preventive, (3) larval curative. Seasonal biology of the hunting billbug is currently under study at Purdue University.

Figure 4. Bluegrass billbug egg inside the stem of a Kentucky bluegrass plant.

Figure 5. Bluegrass billbug larvae inside stem of a Kentucky bluegrass plant.

Hunting Billbug

Although adult hunting billbugs are also easy to recognize by their characteristic snout, they differ somewhat from bluegrass billbug in both appearance and biology. Hunting billbug adults vary in size from about 8-11 mm and are usually dark reddish-brown in color although they may also be coated with soil giving them a dirty appearance. In contrast to the bluegrass billbug, the area behind the head (pronotum) is covered with unevenly spaced punctures that are not uniformly sized. The area behind the head also exhibits a raised Y-shape that is surrounded on each side with a parenthesis (Y). Like other billbugs, hunting billbug larvae are white and legless, with a chestnut colored head (Fig. 6).

Figure 6. Hunting billbug larva in the root zone of zoysiagrass.

Both adults and larvae of this species overwinter at least as far north as West Lafayette, IN (40.5°N). Overwintering adults become active in April and immediately begin laying eggs in leaf sheaths and near the crowns of plants. Overwintering larvae resume feeding on plant crowns, roots, stolons, and rhizomes in the spring. These larvae soon pupate and emerge as adults resulting in a prolonged period of adult activity during the spring and early summer. Larger larvae resulting from overwintering adults are generally present in the soil from mid-June, into September. Some of these larvae apparently pupate and emerge as adults before winter while others spend the winter as larvae. Larval activity may be accompanied by significant damage to turfgrass whenever they are present, but particularly from July through September. Adults of this species may damage turf when populations are high.

Other Billbug Species Associated with Turfgrass

Although not uncommon, much less is known about the biology of two additional billbug species associated with turfgrass in the Midwest the lesser and unequal billbugs. These species often occur in mixed populations with the bluegrass and hunting billbugs and probably have a seasonal biology similar to the bluegrass billbug (one generation of larvae each year). The unequal billbug is a bit broader than the bluegrass billbug, but about the same length (7-8 mm). The area behind the head is covered with unevenly spaced punctures that are not uniformly sized, as well as a smooth, raised area resembling an elongated diamond shape. It apparently feeds on both warm- and cool-season grasses, but its status as a pest has not been confirmed.

The lesser billbug is a bit smaller than the bluegrass billbug (6-7 mm). The area behind the head is more sparsely punctured and the punctures are very obviously not of uniform size. Otherwise, there are no other distinguishing marks. Like the uneven billbug, it apparently feeds on both warm- and cool-season grasses and its status as a pest has not been confirmed.

DAMAGE AND DIAGNOSIS

Billbugs are the most commonly misdiagnosed insect-related turfgrass disorder in North America. The list of ailments for which billbug damage is confused includes compacted soil, drought or summer dormancy, nematode damage, spring dead spot and dollar spot disease. Damage caused by billbugs is often incorrectly attributed to other insects such as white grubs. As a result, these insects can become a perennial problem leading to seriously degraded stands of turfgrass that are easily overrun by weeds. Billbugs affect roughly half of all home lawns in Indiana making them the most common turfgrass-infesting insect in our region.

Billbug larvae primarily damage turfgrass by feeding on plant crowns and roots, and damage is similar regardless of billbug species. The first indications of billbug feeding are usually visible by mid-June when individual plants start to decline as a result of crown-feeding by the larvae. This early phase of damage appears as dead spots about 2-3” in diameter. As damage proceeds, these spots may come together forming large, irregular patches of dead and damaged turf (Fig. 7). In areas where hunting billbug is present, patches of turf may exhibit delayed green-up during the spring as a result of feeding by overwintered larvae. Damage from adult hunting billbugs feeding on stems and leaves has been reported where populations densities are high.

Figure 7. Kentucky bluegrass (A) high-cut zoysiagrass (B) and short-cut zoysiagrass (C) showing typical symptoms of billbug damage.

Detection and Monitoring

Diagnosing billbug damage is a relatively straight-forward process. In areas where damage is suspected, plant crowns can be examined during June July and August by pulling on dead or damaged stems. If stems easily dislodge or break-off at the soil surface, the bottom ends should be examined for the presence of fine, powdery, sawdust-like material (frass). The presence of this material is diagnostic for billbug larval feeding (Fig. 8). In warm-season turfgrasses such as zoysia or Bermudagrass, a similar technique can be employed, but inspection of plant stolons and rhizomes may also be useful (Fig. 9). Larvae can be detected directly using of a golf course cup cutter or a sturdy knife to cut a core or wedge into the sod about 3 inches deep. The soil can then be broken apart and carefully examined for the presence of larvae in the crowns and roots

Figure 8. A tug-test can be used to examine the bottom-ends of Kentucky bluegrass tillers that pull easily from the sod and are filled with fine sawdust-like frass indicating billbug damage.

Figure 9. Zosiagrass shoots hollowed-out by hunting billbug ( Photo Credit: A.J. Patton).

Scouting for adult billbugs can provide an early indication of a potential billbug infestation. Since adults are not capable of sustained flight, they often use driveways, sidewalks, cart paths and curbs to disperse in the spring and late summer (Fig. 10). Experienced turfgrass managers will keep an eye on these areas for adult billbug activity as this can serve as an early indicator of billbug presence. Pitfall trapping can also be used to monitor adult activity and linear pitfall traps can be particularly effective for this purpose (Fig. 11). However, adult activity only indicates that billbugs are present and does not necessarily predict damage by larvae. Unless billbug damage has been previously diagnosed, control may not be warranted.

Figure 10. Billbugs often used sidewalks and curbs to disperse. Observing these areas can serve as a simple monitoring tool. ( Photo Credit: H.D. Niemczyk).

Figure 11. Plans for constructing a linear pitfall trap used to monitor adult billbug activity (A). Installed trap (B).

BILLBUG MANAGEMENT

Billbug management relies on a combination of cultural, biological and chemical tools aimed at keeping populations below damaging levels. Although detection of adults can be an indication of a potential problem, it is usually the larvae that damage plants. Larval populations densities of 10/ft2 are not uncommon and most turfgrasses can tolerate such densities without suffering significant damage.

Cultural Tools

The primary challenge for turfgrass managers is striking a balance between the functional and aesthetic requirements of the turf and maintaining an environment that is suitable for beneficial organisms and the services they provide. Sound cultural practices that include, 1) selection of turfgrass species and cultivars that are well adapted for a specific site or use and 2) proper mowing, fertilization, irrigation, thatch management and cultivation to promote healthy, vigorous turf. Such turf is capable of tolerating or quickly recovering from insect feeding and serves as the foundation of “integrated pest management” (IPM).

Resistant Turfgrasses

Resistant turfgrass varieties play an important role in integrated pest management because they are less likely to suffer damage and quicker to recover if damage should occur. When complimented by proper mowing, irrigation and fertilization, planting resistant varieties can reduce or eliminate the need for chemical insecticides.

Endophyte-Enhanced Varieties

Endophyte-enhanced (E+) turfgrasses, including many cultivars of perennial ryegrass, tall fescue and creeping red fescue provide resistance to billbug adults and larvae. These grasses harbor symbiotic fungi ( Neotyphodium spp.) (Fig. 12) that deter feeding and development of above-ground insects and provide improved tolerance to environmental stresses such as heat and drought. A stand of turfgrass composed of at least 40% E+ plants is generally recommended for providing billbug resistance and overseeding E+ grasses into an otherwise susceptible stand can achieve excellent results. However, reliable estimates of endophyte infection must be assessed in living plants. Infection rates measured in the seed only provide an estimate of initial infection and viable infection may actually be much lower. Low infection rates limit the utility of endophyte-enhanced turfgrasses in IPM. For a list of E+ turfgrass cultivars and initial endophyte infection rates measured in the seed, see http://www.ntep.org/endophyte.htm. Endophyte infection rates in living stands of turfgrass can be assed using a commercial endophyte detection service available from Agrinostics Ltd. Co. ( http://www.agrinostics.com).

Figure 12. The fungal endophyte Neotyphodium coenophlalum in tall fescue. Note the darker stained fungal hyphae growing between the plant cells.

Resistant Kentucky Bluegrasses

Several varieties of Kentucky bluegrass exhibit resistance or tolerance to bluegrass billbug, probably due to their finer texture and narrower leaves and stems that are not preferred for egg-laying. Among these varieties are Arista, Barvette HGT, Delta, Eagleton, Kenblue, Midnight, NuDwarf, Park, Prosperity, Ram I, South Dakota certified, Unique, Wabash, Washington, Wildwood and 4-Season. Susceptible varieties include Broadway, Canterbury, Classic, Georgetown and Nassau.

Resistant Warm-Season Turfgrasses

Bermudagrass and zoysiagrass are considered favored hosts for hunting billbug, but resistant varieties of these two warm-season turfgrasses are available. Little is known about resistance in more cold-tolerant varieties of Bermudagrass and the only variety showing good resistance to hubting billbug is TifEagle, with Celebration and Tifdwarf showing moderate resistance. TifWay Bermudagrass is considered highly susceptible to hunting billbug. In zoysiagrass, Z. martella varieties are generally more resistant than Z. japonica varieties. The most resistant varieties of Z. martella included Diamond, and Zorro, whereas Cavalier and Royal are considered susceptible. The most resistant varieties of Z. japonica include De Anzo and El Toro whereas Pallisades, Meyer and Crowne are considered susceptible.

Biological Controls

Although a host of pathogens predators and parasites will attack and kill billbug adults and larvae, commercially available, effective biological controls are limited primarily to the insect-parasitic nematodes Heterorhabditis bacteriophora and Steinernema carpocapsae (Fig. 13). H. bacteriophora is effective against billbug larvae once they have entered the root zone, whereas S. carpocapsae is more effective against adult billbugs. When used properly, these products can provide adequate control and are generally safer than chemical insecticides. However, special considerations must be made when using insect parasitic nematodes.

Figure 13. Infective juvenile of the insect-parasitic nematode Steinernema carpocapsae a biological control for billbug adults.

Nematode products should be refrigerated upon arrival and stored as briefly as possible. Nematode viability should be checked prior to application by examining a small amount of the spray solutions with a magnifying glass to ensure the nematodes are active and moving about. After mixing, nematodes should be applied immediately and not allowed to sit in the tank for more than a few hours without agitation. Applications should be made in the early morning or evening to limit exposure to UV radiation and irrigation should immediately follow application in order to wash the nematodes off of the turf canopy and into the soil. Screens should be removed from the spray nozzles and spray equipment should be pressurized to a maximum of 50 psi. CO2 should not be used to pressurize spray equipment as nematodes may be asphyxiated.

Chemical Insecticides

There are three basic strategies for using insecticides to target billbugs. Table 1 provides a list of insecticides recommended for each of these strategies. These recommendations are based primarily on knowledge of bluegrass billbug seasonal biology. The seasonal biology of hunting billbug in the Midwest has not been well studied and efforts to understand its seasonal biology are currently underway at Purdue University. Until a better understanding of hunting billbug biology has been gained, special considerations (as outlined at the bottom of this section) may be required to manage this insect.

Strategy 1: Preventive Control of Adults

This approach relies on the use of a surface applied, contact insecticide to target adults as they emerge from overwintering and before they have a chance to deposit eggs. This is also the most effective time for using the insect-parasitic nematode Steinernema carpocapsae. Because adult activity during this time of year can fluctuate with unpredictable spring weather patterns, the primary challenge for using this approach is timing of application. This is especially true when using products with moderate- to short-term residual activity like pyrethroids, carbamates organophosphates or nematodes. However, as a rule of thumb, bluegrass billbugs historically become active by late-April (southern Indiana) or early May (northern Indiana) with hunting billbug activity beginning about 2 weeks earlier. Activity of adult bluegrass billbugs can also be predicted using a degree-day model. With this approach, heat units can be tracked using average daily temperatures starting on March 1 and a developmental threshold of 50 o F. Adult bluegrass billbugs first become active around 280 DD50. No degree-day models exist for predicting hunting billbug adult activity. After making an application targeting adult billbugs, liquid materials should be left on the surface or only lightly irrigated in order to achieve maximum contact activity. Granular materials should always be lightly irrigated in order to wash the active ingredient from the granule.

Strategy 2: Preventive Control of Larvae

This approach targets the larvae inside the plant after adults have begun depositing eggs, but before damage is visible. It is a somewhat more flexible approach since it relies on the use of systemic insecticides (neonicotinoids or diamides) that are taken up by the plant and distributed throughout the plant tissues over an extended period of time. In this way, the active ingredients are able to reach the larvae inside plant stems. The neonicotiods in particular also have good contact activity against adult billbugs so they can be used to target adults while providing residual plant-systemic activity against the larvae inside the stems. Optimal timing for using this approach is roughly the same as for the adult preventive strategy with efficacy decreasing by mid-June. Post-application Irrigation is recommended to wash material into the root zone where it can be taken up by the plants. Bluegrass billbug larvae usually begin tunneling inside stems at 650 DD50.

Strategy 3: Curative Control of Larvae

This strategy targets larvae in the crowns and soil after damage has become apparent. In this regard, it is a reactive strategy aimed at mitigating damage that is ongoing. Most soil insecticides labeled for use in turfgrass can be used in this manner, including neonicotinoids, carbamates and organophosphates. The insect-parasitic nematode Heterorhabditis bacteriophora is also most effective when used in this capacity. The window for using this approach is fairly narrow since the appearance of damage is highly dependent on weather conditions (especially rainfall) and can manifest quickly. Larvae of bluegrass billbug start to appear in the soil at 926 DD50 In order to reach soil dwelling larvae, insecticide applications should be followed with rainfall or irrigation within 24 hours.

Special Considerations for Hunting Billbug in Warm-Season Turf

Because hunting billbugs overwinter as larvae and adults, a prolonged period of adult activity may result during spring and early summer. Overwintering larvae will develop into adults and emerge during this late spring/early summer time frame. These two separate, but overlapping cohorts of adults can make proper timing of insecticide applications difficult to achieve. A combination of the strategies outlined above, with the second application of insecticide 6-8 weeks after the first application may sometimes be required to prevent damage to turf.

Table 1. Active ingredients of Synthetic insecticide products recommended for use against billbugs in turfgrass.
Insecticide*
(Trade Name/Manufacturer)
Insecticide Class Adult Stage
(Strategy 1)
Larval Stage in Stems
(Strategy 2)
Larval Stage in Soil
(Strategy 3)
Beta-cyfluthrin
(Tempo/Bayer)
Pyrethroid X
Bifenthrin
(Talstar/FMC)
Pyrethroid X
Carbaryl
(Sevin/Bayer)
Carbamate X X
Chlopyrifos a
(Dursban/Dow)
Organophosphate X
Chlorantraniliprole
(Acelepryn/Syngenta others)
Diamide X X
Cyantraniliprole
(Ference/Syngenta)
Diamide X X
Clothianidin
(Arena/Nufarm others)
Neonicotinoid X X X
Deltamethrin
(DeltaGard/Bayer others)
Pyrethroid X
Dinotefuran
(Zylam/PBI-Gordon)
Neonicotinoid X X
Imidacloprid
(Merit/Bayer, others)
Neonicotinoid X X X
Lambda-cyhalothrin
(Scimitar/Syngenta)
Pyrethroid X
Thiamethoxam
(Meridian/Syngenta)
Neonicotinoid X X X
Trichlorfon
(Dylox/Bayer)
Organophosphate X X
Zeta-cypermethrin
(Talstar Xtra/FMC)
Pyrethroid X
*Always consult label directions for specific timing and application recommendations.
a Labeled only for use on turfgrass grown for sod or seed.

Table 2. Active ingredients of BIOLOGICAL/BIORATIONAL insecticide products recommended for use against billbugs in turf-grass.
Insecticide*
(Trade Name/Manufacturer)
Insecticide Class Adult Stage
(Strategy 1)
Larval Stage in Stems
(Strategy 2)
Larval Stage in Soil
(Strategy 3)
Heterorhabditis bacteriphora
(Nemasys G/BASF others)
Parasitic nematode X
Steinernema carpocapsae
(Millenium/BASF others)
Parasitic nematode X
*Always consult label directions for specific timing and application recommendations.
aLabeled only for use on turfgrass grown for sod or seed.

READ AND FOLLOW ALL LABEL INSTRUCTIONS. THIS INCLUDES DIRECTIONS FOR USE, PRECAUTIONARY STATEMENTS (HAZARDS TO HUMANS, DOMESTIC ANIMALS, AND ENDANGERED SPECIES), ENVIRONMENTAL HAZARDS, RATES OF APPLICATION, NUMBER OF APPLICATIONS, REENTRY INTERVALS, HARVEST RESTRICTIONS, STORAGE AND DISPOSAL, AND ANY SPECIFIC WARNINGS AND/OR PRECAUTIONS FOR SAFE HANDLING OF THE PESTICIDE.

It is the policy of the Purdue University Cooperative Extension Service that all persons have equal opportunity and access to its educational programs, services, activities, and facilities without regard to race, religion, color, sex, age, national origin or ancestry, marital status, parental status, sexual orientation, disability or status as a veteran. Purdue University is an Affirmative Action institution. This material may be available in alternative formats.

This work is supported in part by Extension Implementation Grant 2017-70006-27140/ IND011460G4-1013877 from the USDA National Institute of Food and Agriculture.


Insects of Cyprus

Acontia lucida
Acrotylus insubricus
Agapanthia gemella
Ameles cypria
Anthaxia (Cratomerus) diadema
Autographa gamma
Blepharopsis mendica
Celastrina argiolus
Charaxes jasius
Chilades trochylus
Chrysis judaica
Chrysis mavromoustakisi
Chrysura sulcata
Cigaritis acamas cypriaca
Colias croceus
Coniocleonus excoriatus
Crocothemis erythraea
Exodrymadusa inornata
Glaucopsyche paphos
Libelloides macaronius
Lycaena thersamon
Maniola cypricola
Ophiusa tirhaca
Orthetrum chrysostigma
Pelopidas thrax
Perotis susannae
Phytoecia (Helladia) humeralis
Polyommatus icarus
Quercusia quercus
Sympetrum striolatum
Trachyderma philistina
Trithemis festiva
Tylopsis lilifolia
Zerynthia cerisyi cypria

Agapanthia dahli
Agapanthia gemella
Agapanthia nicosiensis
Aporus bicolor
Arhopalus ferus
Arhopalus syriacus
Axinopalpis barbarae
Calamobius filum
Callergates gaillardoti
Cerambyx nodulosus
Cerambyx welensii
Certallum ebulinum
Chlorophorus sartor
Coptosia ganglbaueri
Deilus fugax
Euchlanis contorta
Eutheia formicetorum
Glaphyra bassettii
Grammoptera baudii
Haplothrix subtilis
Helladia adelpha
Hylotrupes bajulus
Isocerus balearicus
Isocerus purpurascens
Lampropterus femoratus
Leiopus syriacus
Leiopus andreae
Leprosoma stali
Nathrius brevipennis
Niphona picticornis
Pedostrangalia adaliae
Pedostrangalia raggii
Penichroa fasciata
Phoracantha semipunctata
Phymatodes testaceus
Phytoecia geniculata
Phytoecia croceipes
Poecilium fasciatum
Pogonocherus anatolicus
Purpuricenus nicocles
Purpuricenus nudicollis
Rhaesus serricollis
Rhodopsis pusilla
Saperda punctata
Sphenoptera oertzeni
Stenhomalus bicolor
Stenhomalus bicolor
Stromatium unicolor
Trichoferus fasciculatus
Trichoferus antonioui
Trichoferus georgiui
Trichoferus griseus
Xylotrechus antilope


National Science Foundation - Where Discoveries Begin

A conversation about conserving and naming species


More than half of all marine species may be on the brink of extinction by 2100, says UNESCO.


January 4, 2013

A little more than 39 years ago, on Dec. 28, 1973, the Endangered Species Act was enacted to conserve threatened and endangered species and their ecosystems. To honor this anniversary, Daphne Fautin of the National Science Foundation (NSF) answered questions about biodiversity.

As a marine biologist, Fautin has literally gone to the ends of the Earth--from the poles to the tropics--to study marine life. She is currently a program manager at NSF, a professor of ecology and evolutionary biology at the University of Kansas, and a commissioner with the International Commission on Zoological Nomenclature, which produces rules on giving scientific names to animals.

What is biodiversity?

Biodiversity--short for "biological diversity"--is the variety and abundance of plants, animals and other living things on Earth and in particular locations. Biodiversity is absolutely essential to ecosystem health. And human survival depends on the health of our planet's ecosystem.

Rain forests and coral reefs are known for their biodiversity. Why is so much biodiversity concentrated in these types of ecosystems?

More than 25 percent of the world's fish species and between nine and 12 percent of all of the world's fisheries are associated with coral reefs. More than half of the world's plant and animal species live in rain forests.

We don't know for sure why rain forests and coral reefs harbor so much biodiversity. One idea is that these ecosystems occur in tropical climates, and so they are quite climatically consistent year-round.

According to this idea, tropical organisms diverged because they don't have to deal with the climatic extremes that organisms at higher latitudes (and altitudes) do. A rabbit, for example, that lives in a non-tropical place must be able to eat certain plants in the summer and certain other plants in the winter. Therefore, it must remain a generalist to survive.

By contrast, a rabbit that lives in the tropics may specialize in eating certain plants that are available year-round at the same time, other species of rabbits (or other organisms) may evolve that specialize in eating other plants. Such specialization promotes diversity.

But some evidence does refute this idea--such as the fact that not all groups of plants and animals demonstrate more diversity in the tropics than at higher latitudes. So, many other ideas have also been proposed to explain the extraordinary biodiversity of the tropics.

Insects account for a large proportion of the biodiversity on Earth. Why?

Many statistics bear out the biodiversity of insects. For example, more than 850,000 insect species have been named. And the total estimated weight of just ants in the Amazon is four times the estimated weight of all land vertebrates in the Amazon--including all mammals, birds, reptiles and amphibians!

We don't really know why insects account for so much of the Earth's biodiversity. That is one of the questions that is being studied by entomologists--the people who research insects.

One idea is that insects began to diversify when flowering plants evolved on Earth, and so insects evolved along with flowering plants because they are so important in the pollination of plants.

Insects tend to be small and specialized: There may be a certain insect that sucks out the cell sap from the stems of a particular plant other insects that eat that plant's leaves other insects that feed on that plant's nectar and other insects that feed on that plant's pollen and pollinate the plant in the process. So as flowers evolved, many insects evolved as well.

This idea about the evolutionary connections between flowering plants and insects is consistent with what we see in the oceans: Relatively few species of flowering plants and relatively few species of insects live in the oceans.

(By the way, NSF recently issued a press release that identified some interesting reasons why humans need insects--even pesky ones.)

How many species have been described and named by scientists, so far?

According to some reliable estimates, there are 1.9 million known eukaryotic organisms. (Eukaryotic organisms are those that are made of one or more cells with a nucleus bacteria and viruses are not eukaryotic organisms.)

How many species exist on Earth?

Estimates range from 2 million to 10 million species.

A recent estimate of 8.7 million species received a lot of press, in part, I suspect, because of its supposed accuracy and because it corresponds quite well to the often-bandied figure that 80 percent of the Earth's biodiversity has yet to be discovered/named. Some scientists estimate that there are "at least four times" the number of known species exist on Earth. If there are indeed almost 2 million known organisms, this estimate would translate to at least 8 million species.

A paper estimating the number of marine species (which I contributed to) was recently published. According to this paper, 226,000 species that live in the ocean have been named and described by scientists, and 72,000 additional species are in collections waiting to be named and described.

But who knows what hasn't been collected yet? And of course, as I previously mentioned, oceans have few insects, but insects account for the bulk of biodiversity.

How can scientists estimate the total number of species on Earth when it is obviously impossible to count what has not yet been counted? In other words, how can we know what we don't know?

People have used various creative methods to estimate the total number of organisms on Earth. For example, there was a very large estimate made years ago by a scientist who went to the jungles of Panama and used insecticide to spray a tree in the jungle. Then, lots of insects died and fell from the tree to the ground. And the scientist and his colleagues identified and counted as many of these fallen insects as they could, and the rest were counted as unknown. Then, the scientist extrapolated from the proportion of species in that one tree that were known versus unknown to produce a global estimate of known versus unknown species.

It was a good first try. But a lot of people point to the fact that in many parts of the world, the proportion of known species to unkown species is higher than it is in Panama. So this fact would suggest that the estimate may be excessive.

In 2011, an NSF-funded researcher provided the first empirical evidence of what had been long suspected: that biodiversity promotes water quality. What are some of the other reasons why we need biodiversity?

We need biodiversity to eat. We need to preserve species that we use as food, including fish from the sea. We also need to preserve those species that serve as food for the fish we eat, so that our food supply persists. And we also need to preserve all the species that create the habitat that enables all of these needed species to live, spawn and raise their young.

So, there are all of these connections in the great "web of life" that we don't even know yet. And these connections support all of the species on Earth, including species that provide us with food and clothing.

Also, 50 percent of the oxygen we breathe is produced by microscopic plants that live in the ocean and the other 50 percent is produced by plants that live on land.

(If you want to learn more about the ways in which the various species of plants help humans survive, watch this dynamic, upbeat video produced by NSF.)

The planet's ecosystem is sometimes compared to an airplane. You can lose one rivet from an airplane, and the airplane will probably fly. You can lose two rivets for an airplane and the airplane will still probably fly. But eventually, if you lose too many rivets--and nobody knows how many--the plane will crash.

The same principle applies to ecology: You can lose some species without major harm. But no one knows how many species can be lost before the planet's ecosystem will crash.

What does it mean to discover a new species?

A new species is one that hasn't yet been formally described and named according to scientific procedures--not one that is newly evolved.

People on the street or people in the jungle may have a name for it. But if we haven't followed the internationally recognized rules of nomenclature for describing and naming a species, it doesn't exist for certain scientific purposes.

When we have discovered a new species, it means we have finally found and gone through the procedures of formally describing it (distinguishing it from other species) and giving it a name following the rules of nomenclature.

How many new species are named each year?

Between 15,000 and 20,000 new species are named each year.

A species may be discovered and collected before it is described and named. But it isn't recognized as a new, distinct species until it is described and named.

How many species go extinct each year?

We don't know. The World Wildlife Fund's website says that experts have calculated that between .01 percent and .10 percent of all species on Earth go extinct each year.

But because we don't know how many species there are, we don't know how many species those percentages actually represent. And so, if the low estimate of the number of species on Earth is true--if there are around 2 million species on our planet--then between 200 and 2,000 extinctions occur each year.

But if, in fact, there really are 10 million species on Earth, then between 10,000 and 100,000 extinctions occur each year.

What would you say to naysayers who argue that newly discovered species offset species losses, and so there really is no extinction crisis?

"New" species are not newly evolved. They evolved a long time ago. They are simply being newly discovered by science. They may be very well known to the people living in the areas where they live. So they aren't new in that sense they are only new to those of us who name them.

What does the process of naming a species involve?

It can be a long, protracted and difficult process that can take many years. First, you want to be sure that the animal or plant hasn't been named before.

This can be difficult for a variety of reasons. For one thing, many descriptions that were prepared in the early days were very vague. And usually, only small groups of experts have the specialized expertise to know what has and hasn't been described before.

And in order to name a species, you also have to describe it. To do this, you have to know what kinds of features are used to identify species, and figure out what distinguishes the "new" species from known species. This is important, because at least according to the rules of zoological nomenclature, when you publish a description of a new species, you have to write out what makes it different from everything that is already known--including organisms that are not closely related to it, but that look like it anyway.

For example, suppose you have an organism that has a red spot then you have to distinguish your "new" species from everything that is red spotted, even if those other red-spotted species are not closely related to your species. That way, when somebody comes across your species, they can say, "Aha! This is another one of those red-spotted organisms that is covered by that new name it's not another thing with red spots."

Other rules in the codes of nomenclature require you to name species in Latin or make them sound like Latin. You also have make sure a specimen of your species is deposited in a natural history collection (typically in a museum or herbarium). If the "new" species is an animal, you may also have to register the name in ZooBank (The official Registry of Zoological Nomenclature).

And then you have to publish your description of the species in a scientific journal, so that other scientists can look at it and agree that it is correct.

It is interesting to note that names that are accepted are frequently later "sunk" for various reasons. For example, when the exhaustive homework that is required for describing and naming a "new" species is not conducted in a comprehensive and thorough way, it may ultimately turn out that the "new" species has already been described and named.

Alternatively, a name for a "new" species may be sunk because the difference that was thought to distinguish it from others does not hold up. For example, I have a colleague who described several coral reef fish as "new"--only to ultimately discover that the "new" species was a member of a species for which only one sex had previously been identified. So the "new" species was really just a female (or male) of a known species!

Do you have to be a professional taxonomist to identify and name new species?

About half of the species that are named each year are named by people who aren't employed as taxonomists--whose job isn't in a museum or university. In fact, some of the people with the most expertise and time to do this are not professionals in the field.

For example, I knew a dentist who is one of the world's foremost authorities on tiger beetles. He had earned a master's degree in entomology. And then he realized that if he had gone into academic entomology, he would be spending his time teaching, writing grant proposals and doing administrative work--but he wanted to catch tiger beetles.

And so he went into dentistry so that he could make enough money to take time off each year to catch tiger beetles. He probably thereby ended up being able to spend more time chasing tiger beetles as a dentist than he would have if he had become a professional entomologist.

A new species of frog was recently identified by an NSF-funded researcher right smack in New York City. Is that common for new species to be discovered in such populated places?

I think that it is quite common for new species to be found in populated areas.

In the summer of 2012, an NSF-funded researcher named a new coral reef crustacean after Bob Marley, the singer. Is that unusual for species to be named after celebrities?

It may be less common than it used to be.

Many years ago, it would be common for a patron to fund the travels of a scientist to exotic places, and then species that were found during those travels would be named for the patron.

Tell me about one of the big problems that is reducing biodiversity in the oceans?

We have depleted the oceans of many of the big fish that we eat.

Part of the reason we are able to over fish is due to technology. We have fish-finders, various types of tracking devices, including sonar equipment, and airplanes that are used to help find schools of fish. And we now know enough about marine biology to predict where fish and crustaceans will be under particular conditions. Fishing is no longer about a fisherman just saying, I'll drop a line here or there." Fishing has become very scientific and methodical.

When we trawl, we drag nets along the ocean floor. And whatever else gets swept by these nets in addition to the target fish is called "bycatch." This bycatch gets thrown back into the ocean because we are not licensed to take it or because it's not profitable to take.

But how many organisms can manage to survive after being caught in a big net, pulled up and then thrown back into the ocean? What's more, trawling destroys habitat after the ocean floor has been trawled, it may no longer be a suitable home for what is thrown back.

The analogous situation on land would be if we flew an airplane that dragged a big net across the ground to catch grazing cattle. And we would draw up the net periodically--and keep the cattle, but toss back the dogs, trees and everything else that we happened to net in addition to the cattle. That is similar to what we do to the oceans.

Many people assume that it is better to consume farmed fish and shrimp than wild caught fish and shrimp. But many fish and shrimp farming practices are also harmful to the environment. Just because it is farmed doesn't mean that it is environmentally neutral or preferred over wild caught.

Can you cite any "good news" stories in biodiversity?

I read the other day that we have lost 97 percent of wild tigers in just over a century. Only about 3,200 tigers currently remain in the wild. We can infer that the populations of many smaller species are plummeting just like populations of many large species are plummeting.

But I am happy to say that a few species have come off the endangered species list, like the wolf and the bald eagle, because plans for their recovery were enacted.

Also, there are "good news" stories in the history of whales. Many of them were hunted to the brink of extinction, and then they were listed as endangered. It therefore became legally, socially and economically difficult to harvest whales, and so populations of many whale species have fortunately recovered.

These kinds of successes show that if we stop harvesting species, and if their habitat is conserved, life is resilient and endangered species may recover.

Additional Resources

To learn more about biodiversity and help promote conservation:

  • Read about the Endangered Species Act on the websites of the U.S. Fish & Wildlife Service and the National Oceanic and Atmospheric Administration.
  • Join citizen science groups that work to identify and track species of birds, butterflies, ladybugs, plants and other creatures, advance our understanding of nature, and increase habitat for wildlife.
  • Restrict your seafood purchases in restaurants and stores to ocean-friendly products. Resources provided by the Monterey Bay Aquarium's Seafood Watch Program may help you do so.


Daphne Fautin searches for new organisms while conducting a marine survey.
Credit and Larger Version

Investigators
Daphne Fautin


Daphne Fautin searches for new organisms while conducting a marine survey.
Credit and Larger Version


Green milkweed grasshopper

Scientific name: Phymateus viridipes

Distribution: Southern Africa

Size: 70 mm (2.75 inches) long

This large African grasshopper secretes a noxious fluid from the thorax when alarmed. The fluid is derived from the poisonous milkweed plants upon which it feeds as an immature nymph or adult. The colored hind wings, which are normally hidden when the grasshopper is at rest, can also be flashed to deter potential predators.


Turfgrass Insects

This publication provides turfgrass management professionals and property owners with information to help them 1) properly identify the most common white grub species associated with turfgrass in Indiana and adjacent states, 2) understand white grub biology, 3) recognize white grub damage and 4) formulate safe and effective white grub management strate­gies. For information on turfgrass identification, weed, disease and fertility management, visit the Purdue Turfgrass Science Website https://turf.purdue.edu or call Purdue Extension (765-494-8491).

WHITE GRUB SPECIES ASSOCIATED WITH TURFGRASS IN THE MIDWEST

White grubs represent a complex of beetle larvae in the fam­ily Scarabaeidae that are common pests of agricultural and horticultural systems. Often called scarab beetles, the family consists of over 30,000 species world-wide. The larvae, or grubs, of several species are common pests of turfgrass. These species include the Japanese beetle, masked chafers (2 species), European chafer, Asiatic garden beetle, Oriental beetle, green June beetle, May/June beetles (several spe­cies), and black turfgrass ataenius. White grubs damage a variety of warm- and cool-season grasses while feeding in the soil matrix on organic matter, thatch and plant roots. The distribution of these species overlaps significantly and it is not uncommon to find mixed populations of two or more species at a single location.

IDENTIFICATION AND SEASONAL BIOLOGY

Proper identification and basic understanding of the varying life cycles of different white grub species can help turfgrass managers monitor, plan for and manage infestations. White grubs are white, C-shaped insects with a chestnut colored head and 3 pairs of legs that are clearly visible (Fig. 1). The rear end is slightly larger in diameter than the rest of the body and may appear darker in color due to the soil and organic matter they ingest. Size may vary considerably depending on the species and age, but older larvae will generally range from 1/4 to 1-1/2 inches in length. White grubs can be identified to genus or species based on the conformation of the raster pattern. The raster pattern is composed of a series of short hairs and spines on the underside of the tip of the abdomen (Fig. 2). A hand lens, magnifying glass or microscope may be required to see the pattern clearly. The life cycles of these insects can be grouped broadly into three categories (annual, semi-annual and multi-annual) based on the amount of time required to complete development.

Figure 1. A typical white grub. Notice that the body is C-shaped and 3 pairs of legs are present. The yellow arrow indicates the location of the raster pattern that is useful for identification.

Figure 2. Raster patterns of annual and multi-annual white grub species common in the Midwest.

Annual White Grubs

Annual white grubs are the most common pest species, pro­ducing one generation every year. Several annual white grubs species (Japanese beetle, European chafer, Asiatic garden beetle and Oriental beetle) are considered exotic, invasive species, but others are native to North America (masked chafers, green June beetle). These species overwinter in the larval stage, pupating in the soil during late spring or early summer (Fig. 3). Adults emerge and fly during early- to mid-summer and begin laying eggs in the soil. The adults of some species (e.g., Japanese beetle, Asiatic garden beetle) can be serious pests of ornamental plants during this time. Eggs hatch by the end of July producing small larvae that begin feeding in the root zone. Large larvae are present by September, but damage may appear anytime between August and November. Late instar larvae migrate down into the soil to spend the winter. Larvae migrate back up into the root zone to feed again in the spring before pupation and damage to turf may also occur during this time.

Figure 3. Life cycle of a typical annual white grub and relative timing of three different chemical or biological management strategies preventive (strategy 1), early curative (strategy 2) and late curative (strategy 3).

Semi-Annual White Grubs

The Black turfgrass ateanius produces two generations of larvae each year and is the primary species of semi-annual white grub affecting turfgrass in the Midwest. Adults are small black beetles (3.5-5.5 mm in length) that overwinter in the thatch and soil along the edges of golf course fairways. Adults migrate from overwintering areas during spring, about the time redbuds (Cercis canadensis) are in bloom (April), and begin laying eggs when Vanhoutte spirea (Spirea x vanhouttei) is in full bloom (May) (Fig. 4). Eggs are laid in the soil in small clusters that typically hatch by late May. First generation larvae feed until mid-July, but damage may be visible by late June. These larvae pupate producing new adults by early August. These adults typically lay eggs producing a second generation of grubs by mid-August in all but the most northerly parts of the Midwest. Damage from this second generation is not uncommon. Second generation larvae pupate by September and emerging adults usually leave the fairways for overwintering sites by October. This species is not known to damage lawns, but is capable of causing serious damage to closely mowed golf course turf.

Figure 4. Life cycle of a typical semi-annual white grub, black turfgrass ataenius, and relative timing of three different chemical or biological management strategies preventive (strategy 1), early curative (strategy 2) and late curative (strategy 3).

Multi-Annual White Grubs

Several species of May/June beetles in the genus Phyllophaga are occasionally associated with damage to turfgrass. These insects require 2-3 years to complete development, depending on latitude. As their name implies, overwintering adults emerge during May and June. These beetles mate and lay eggs in the soil. The resulting white grubs feed during the summer and fall, then migrate deeper into the soil to overwinter. The following spring, larvae migrate back up into the root zone to feed for another season. Because of their larger size, May/ June beetle larvae cause their most severe injury to turfgrass during this second season of their life cycle. Larvae again overwinter in the soil and complete development the following spring and early summer. These larvae stop feeding, pupate and transform into the adult that remains inactive in the soil until the following spring.

DAMAGE AND DIAGNOSIS

White grubs are capable of causing serious damage to turf­grass. Their feeding damages plant roots, causing the turf to wilt and die. Early indications of grub damage may include patchy areas of wilting, discolored or stressed turf that does not respond to irrigation. Turf eventually collapses resulting in dead or extremely thin patches that may range in size from a few meters to large contiguous areas (Fig. 5). This kind of damage, called primary damage, may result in sod that easily pulls-up or becomes dislodged from the soil, revealing the white grubs beneath (Fig. 6). One species in particular, the green June beetle, produces small mounds of soil in the turf marking the entrance to their burrows in spring and late summer (Fig. 7). Secondary damage from raccoons, skunks, or turkeys foraging for white grubs is also common and can sometimes be the first obvious indication of an infestation (Fig. 8).

Figure 5. Damage to a golf course putting green caused by white grubs feeding in the soil.

Figure 6. Turfgrass damaged by white grubs can some­times be peeled back revealing the white grubs beneath.

Figure 7. Small mounds of soil resulting from the activity of green June beetle grubs.

Figure 8. Damage caused by vertebrate ani­mals foraging for white grubs in turfgrass.

Detection and Monitoring

In order to establish that a white grub infestation is present, a golf course cup-cutter or a sturdy knife can be used to cut a core or wedge from the sod to a depth of 3 inches (Fig. 9). The soil can then be carefully broken apart and examined for the presence of white grubs, which may be located high in the soil profile. Relatively high densities (5-10 grubs/ft 2 ) are usually required to cause significant damage, so a few scattered white grubs are not necessarily cause for concern. Areas experiencing damage are likely candidates for future infestation, so close attention should be paid to these problem areas or &lsquohot spots&rsquo.
Monitoring can be useful anytime white grub damage is sus­pected. Strategies that use monitoring to inform treatment decisions should focus soil sampling efforts during July and August for annual white grubs. Golf course superintendents concerned about black turfgrass ataenius grubs should moni­tor vulnerable areas during May-June and August-September. White grub population densities of up 5 ft 2 are not uncom­mon and most turfgrasses can easily tolerate such densities without suffering visible damage. Detection of flying adults at lights or in traps should not be used to predict future white grub infestations or damage.

Figure 9. White grubs can be monitored by extracting a se­ries of soil cores from the turf and carefully breaking them apart to find the grubs.

WHITE GRUB MANAGEMENT

White grub management relies on a combination of cultural, biological and chemical tools aimed at keeping populations below damaging levels.

Cultural Tools

The primary challenge for turfgrass managers is striking a balance between the functional and aesthetic requirements of the turf and maintaining an environment that is suitable for beneficial organisms and the services they provide. Sound cultural practices that include, 1) selection of turfgrass species and cultivars that are well adapted for a specific site and use and 2) proper mowing, fertilization, irrigation, thatch manage­ment and cultivation to promote healthy, vigorous turf. Such turf is capable of tolerating or quickly recovering from most insect-feeding and serves as the foundation of &ldquointegrated pest management&rdquo (IPM).

Biological Controls

Although a host of pathogens, predators and parasites attack and kill white grubs, commercially available, effective biological controls are limited. When used properly, these products can provide reasonable levels of control and are generally safer than chemical insecticides. However, special considerations must sometimes be made when using biological insecticides. Biological insecticides tend to be more expensive than chemi­cal insecticides and they are more variable in the level and speed of control provided. Figure 10 provides a summary of the efficacy of various biological insecticides against white grubs based on the time of application and Table 1 provides a list of strategies for which these products are recomended.

Figure 10. Chart showing the relative efficacy of different biological insecticides against white grub based on the time of application.

Insect-Parasitic Nematodes:Heterorhabditis bacteriophora is a parasitic nematode that attacks and kills white grubs by vectoring a bacterial pathogen. It should be refrigerated upon arrival and used as soon as possible as possible. Nematode viability should be checked prior to application by examining a small amount of the spray solutions with a magnifying glass to ensure the nematodes are active and moving about. After mixing, nematodes should be applied immediately and not allowed to sit in the tank for more than a few hours without agitation. Applications should be made in the early morning or evening to limit exposure to UV radiation, and irrigation should immediately follow application in order to wash the nematodes off of the turf canopy and into the soil. Screens should be re­moved from the spray nozzles, and spray equipment should be pressurized to a maximum of 50 psi. CO2 should not be used to pressurize spray equipment as nematodes may be asphyxiated. Larger larvae are the best targets for nematode applications so they can be used most effectively in early and late curative strategies targeting all but the smallest white grubs (Figs. 3 & 4, Table 1, see list of management strategies under "chemical Insecticides").

Insect-Pathogenic Bacteria:Bacillus thuringiensis galleriae is a strain of naturally occurring soil bacteria that produces toxins capable of killing insects. It can be mixed and applied using methods similar to those employed for chemical insecticides. Post-application irrigation is recommended in order to wash the material into the root zone where white grubs are feed­ing. This product appears to work equally well against small and large white grubs making it useful in both early and late curative strategies (Figs. 3 & 4, Table 1). Although efficacy appears to vary between white grubs species, levels of control ranging from 55-100% can be expected.

Paenibacillus popilliae, also known as milky spore, is a bacte­rial pathogen of white grubs. Although strains of this bacte­rium that infect and kill other white grub species are known, commercially available formulations are only active against Japanese beetle grubs. It is usually applied as a granule or dry formulation, but commercially available products have not proven to be effective. As with other biological insecticides, post-application irrigation is recommended.

Entomophagous Fungi:Metarhizium brunneum (formerly Metarhizium anisopliae) is a soil-born fungal pathogen of many insect species including white grubs. It is commercially available in liquid and granule formulations. Efficacy can vary widely, but the most consistent levels of control are obtained with fall applications targeting later instar grubs. Such ap­plications routinely provide 40-70% control. For this reason, Metarhizium brunneum is most compatible with early or late curative strategies (Figs. 3&4, Table 1). Post-application ir­rigation is recommended to wash the material into the soil where white grubs are actively feeding.

Chemical Insecticides

There are three basic strategies for using chemical insecticides against white grubs. Table 2 provides a list of insecticides recommended for each of these strategies. Figure 11 provides a summary of the efficacy of various chemical insecticides against white grubs based on the time of application. Post-application irrigation or rainfall is recommended for most chemical insecticide applications in order to wash material into the soil where grubs are feeding.

Strategy 1: Preventive

This strategy relies on the use of insecticide formulations that remain active in the soil for an extended period of time. Given the extended residual activity of the insecticides appropriate for this strategy, applications may be made during a broad window ranging from several weeks in advance of egg laying activity to egg hatch. Areas that have a history of white grub infestation and highly manicured playing surfaces, such as golf course fairways, are the most common candidates for this type of approach either because white grub damage is recurring, or because these areas are too valuable to risk damage. This approach is also appropriate when more than one pest species has become a management concern. For example, the application of neonicotinoid or diamide insec­ticides targeting early-season insects such as bluegrass billbug (May) can provide enough residual activity to protect turf from white grubs that become active later in the season (July). This &ldquomultiple targeting&rdquo approach can eliminate the need for repeated applications targeting one pest species at a time and reduce total insecticide use.

Strategy 2: Early Curative

This strategy targets early or late instar white grubs in areas where densities are high enough to be a concern, but before damage is visible. Any registered white grub insecticide is ap­propriate for this approach. Monitoring white grub populations in the soil is a cornerstone of this strategy since the goal is to prevent damage while avoiding unnecessary applications. Population densities of less than 5 grubs/ft 2 rarely require treatment and population densities as high as 20 grubs/ft 2 may not necessarily cause noticeable damage. This is due to differences in size and feeding behavior among white grub species. Thresholds also can vary with turf type, turf health and cultural practices. Although European chafer is capable of causing damage at lower densities (5 grubs/ft 2 ), other spe­cies, such as Japanese beetle, Asiatic garden beetle and black turfgrass ataeneus, usually require higher densities (≥10 grubs/ ft 2 ) to cause visible damage. Because of the burrowing and mounding activity of green June beetle larvae, unacceptable levels of damage may occur at even lower densities.

Strategy 3: Late Curative (Rescue)

This strategy is often referred to as a rescue strategy because it targets white grubs after damage has been noticed. Damage may either be a direct result of white grub feeding (primary damage) or a result of animals destroying the turf while foraging for the grubs (secondary damage). Chemical options for this strategy are somewhat more limited because they must kill, or cause the grubs to stop feeding relatively quickly. Ideally, insecticides used in this capacity will provide an opportunity for the turf to recover and resume growth before winter. The efficacy of late curative applications will be greatly reduced if grubs have stopped feeding or moved deeper into the soil to overwinter.

Deterring Foraging Animals

As previously mentioned, animals foraging for white grubs can be a serious concern for turf managers because of the damage caused as they dig for the grubs. Animals such as raccoons, skunks, armadillos and turkeys routinely forage for and consume white grubs that infest turfgrass even when primary damage from the grubs themselves is not apparent. Although trapping and hunting these foraging animals may provide a long-term solution for turf managers, such activities can be time consuming and are not always compatible with the goals of property managers. One recent study suggests the use of Milorganite organic fertilizer can deter foraging animals, substantially reducing secondary damage to turf. The application of Milorganite to areas damaged by foraging animals at a rate of 0.02 lbs/ft 2 can reduce further damage by 75% or more over the short-term. Lower application rates (0.007 lbs/ft 2 ) can reduce further damage by more than 50%. The long-term residual effectiveness of Milorganite remains unclear, but reactive use appears to be effective at reducing further damage over the short-term.

Figure 11. Chart showing the relative efficacy of different chemical insecticides against white grub based on the time of application.

Table 1. Active ingredients of biological insecticide products recommended for use against white grubs in turfgrass.

*Always consult label directions for specific timing and appication recommendations.

a Effective only against Japanese beetle grubs.

Table 2. Active ingredients of chemical insecticide products recommended for use against white grubs in turfgrass.

*Always consult label directions for specific timing and appication recommendations.

a Effective only against Japanese beetle grubs.

READ AND FOLLOW ALL LABEL INSTRUCTIONS. THIS INCLUDES DIRECTIONS FOR USE, PRECAUTIONARY STATEMENTS (HAZARDS TO HUMANS, DOMESTIC ANIMALS, AND ENDANGERED SPECIES), ENVIRONMENTAL HAZARDS, RATES OF APPLICATION, NUMBER OF APPLICATIONS, REENTRY INTERVALS, HARVEST RESTRICTIONS, STORAGE AND DISPOSAL, AND ANY SPECIFIC WARNINGS AND/OR PRECAUTIONS FOR SAFE HANDLING OF THE PESTICIDE.

It is the policy of the Purdue University Cooperative Extension Service that all persons have equal opportunity and access to its educational programs, services, activities, and facilities without regard to race, religion, color, sex, age, national origin or ancestry, marital status, parental status, sexual orientation, disability or status as a veteran. Purdue University is an Affirmative Action institution. This material may be available in alternative formats.

This work is supported in part by Extension Implementation Grant 2017-70006-27140/ IND011460G4-1013877 from the USDA National Institute of Food and Agriculture.


BugInfo Wasps, Ants, and Bees (Hymenoptera)

Defining the Order. This vast assemblage of insects is second only to Coleoptera (Beetles) in the number of worldwide, described species. Hymenoptera species number some 115,000, and Coleoptera species number some 300,000. Of the 6,000-7,000 new species of insects described annually, Hymenoptera is a large component, especially in the parasitic wasp groups. Nearly all commonly encountered Hymenoptera can be recognized by a narrow "waist." When winged, the wings form two membranous pairs that can be hooked together. Ovipositors of Hymenopteta are usually well developed and modified into a stinger in the higher forms of the order. Because the "stinger" of such forms has developed from the ovipositor of females, male wasps are not able to sting. Many species of Hymenoptera are extremely small and are thus difficult to identify even to family. A publication by Edward Mockford in 1997 recorded discovery of a new species of tiny wasp that is now known as the tiniest existing insect.

Benefits to mankind. This order of insects is considered to be the most beneficial to humankind of all the insects. The strongest benefit performed by most Hymenoptera is active pollination of plants, ensuring the proper development of many fruit and vegetable crops. Many kinds of Hymenoptera are also helpful in their actions of parasitism and predation on pest species of insects.

Ants. These are familiar insects, and are most numerous in tropical forests, where surveys of tree species of insects have consistently shown that individuals of ants compose some 50 percent of the entomofauna. Some species of ants squirt formic acid into wounds. There are more than 8,000 species of ants in the world. Ants are often confused with termites, but have a slender waist and elbowed antennae. In some cases, ants can by pests, especially in such species as the Carpenter Ant, which invades houses near wooded areas. Fire Ants, of course, are a major concern in the Southern United States. Army Ants are perhaps the most fascinating species of ant, capable of preying upon insects, small reptiles, birds, and even small mammals.

Leafcutter ants harvesting and carrying pieces of leaf.
Title: Leafcutter Ants on Leaf.,
Class: A Arthropoda. Order: Insecta. Family: Formicidae.
Genus: Atta. Species: cephalotes.

Wasps. This group of Hymenoptera includes some familiar types, such as Hornets, Spider Wasps, and Hunting Wasps. Sawflies are also a group of wasps, composed of several families, and noteworthy because they have no "waist" that is present in all other Hymenoptera. Tiny parasitic wasps are one of the most beneficial groups of insects, reducing populations of pest species. The Family Ichneumonidae includes vast numbers of parasites also, and is considered one of the largest families of insects.

Bees. The most familiar bee, of course, is the Honeybee, a social insect that was imported from Europe for honey production. Most bees are not social and do not build large nests like the Honeybee. The most colorful of bees are a group of Tropical American insects called Orchid Bees, which have brilliant iridescent colors of green, blue and red. The males visit orchid flowers. Africanized Bees, or Killer Bees, are a major health threat to the population of the Southern United States, and are slowly expanding their geographic range. Entomologists are attempting to find ways to alleviate the invasion of this aggressive strain of bees.

Africanized bee working a flower.

Selected References:

Evans, H. E. & Ebarhard, J. W. 1970. The Wasps. University of Michigan Press, Ann Arbor.

Holldober, B. & Wilson, E. 0. 1990. The Ants. The Belknap Press of Harvard University Press, Cambridge.

Krombein, K. V., Hurd, P. D. Jr., & Smith, D. R. 1979. Catalog of Hymenoptera in America North of Mexico. Volumes 1-3. Smithsonian Press, Washington, D. C.

Michener, C. D. 1974. The Social Behavior of Bees. A Comparative Study. Belknap Press of Harvard University Press.

Michener, C. D., McGinley, R. J. & Danforth, B. N. 1994. The Bee genera of North and Central America (Hymenoptera: Apoidea). Smithsonian Institution Press, Washington and London.

Mitchell, T. B. 1960-1962. "Bees of Eastern United States." Volume 1, Technical Bulletin141, North Carolina Agricultural Experiment Station.

Prepared by the Department of Systematic Biology, Entomology Section,
National Museum of Natural History, in cooperation with Public Inquiry Services,
Smithsonian Institution


The App That Aims To Gamify Biology Has Amateurs Discovering New Species

Bug enthusiast Anna Lindqvist uploads photos like this — of the Ailanthus Webworm Moth (Atteva aurea) to the iNaturalist app. Like a social network for wildlife, her location paired with the photo help both amateur and expert naturalists identify the species. Annika Lindqvist hide caption

Bug enthusiast Anna Lindqvist uploads photos like this — of the Ailanthus Webworm Moth (Atteva aurea) to the iNaturalist app. Like a social network for wildlife, her location paired with the photo help both amateur and expert naturalists identify the species.

It's dusk at a park in Dallas, and white sheets are pinned up next to tall trees, fluttering like ghosts in the wind. They've been lit up with ultraviolet lights to attract moths.

A handful of people are holding up their smartphones, zooming in on the small dark specks that fly to the cloth.

"Bugs have become my obsession," says Annika Lindqvist. "And the more you look, the more you have to look at the tiny things, and when you blow them up you see that they are gorgeous."

Like a lot of bug fanatics, Lindqvist doesn't just take photos of moths — she uploads them to iNaturalist. It's like a social network for wildlife. When you upload a photo of a moth or bird to the app, it posts your location. Then, both amateur and expert naturalists help identify the species.

Lindqvist has uploaded more than 2,000 observations on her profile.

iNaturalist has grown exponentially in the past few years. There are nearly 250,000 users and about 3 million observations. At gatherings like this one, put on by the Texas Parks and Wildlife Department, people who've connected online meet in person to swap stories about giant walking sticks and learn about moths together.

Stalin Murugesapandi, an engineer by day, is one of the citizen scientists here with his smartphone. He points out a moth with feathery antennae that's landed below some mayflies.

Murugesapandi's passion is photography. Some of the moths we're looking at — including one that's meringue yellow and another with bands of olive green — will end up on iNaturalist, next to his pictures of fire ants and turquoise mushrooms.

Stalin Murugesapandi takes photos and makes field sketches to record the different insect species he sees. Lauren Silverman/KERA hide caption

Stalin Murugesapandi takes photos and makes field sketches to record the different insect species he sees.

Sam Kieschnick, an urban biologist with Texas Parks and Wildlife, says an individual photo might not be groundbreaking — and true, you're not getting any PokéCoins or other rewards — but each observation adds to our understanding of biodiversity, like a mosaic or pointillist painting.

"It's just a single dot if you look up close, but when you start to take a step back, you can get to see these patterns that start to develop," Kieschnick says.

There have been major discoveries as a result of photo sharing on iNaturalist.

In 2013, for example, a man in Colombia uploaded a photo of a bright red and black frog. A poison frog expert in Washington, D.C., spotted it and eventually determined it was a brand-new species. The pair co-authored the results in the peer-reviewed journal Zootaxa.

One of the developers behind iNaturalist is Scott Loarie. He says when he partnered with naturalist Ken-ichi Ueda, the initial idea was to use it as a tool to get people engaged with nature, and later, as a tool for science.

He recalls another great discovery, in 2014, when one app user traveling in Vietnam happened to upload a photo of a snail.

"A couple of weeks later, a Vietnamese scientist who was a specialist in snails and slugs was going through and looking at pictures and said 'Wow! I recognize this,' " Loarie says.


The Incredible Ants

Ants are social animals and live in colonies.

Sometimes these colonies may contain only 50 or so individuals… but one supercolony of Formica yessensis on the coast of Japan is reported to have had 1,080,000 queens and 306,000,000 workers in 45,000 interconnected nests.

Ants are highly intelligent, social insects. Here they collaborate by forming a bridge across leaves.

In fact, some Scientist think that 30% of the animal biomass of the Amazon Basin is made up of ants – and that 10% of the animal biomass of the world is ants.

Furthermore they believe another 10% is composed of Termites.

This means that ‘social insects’ could make up an incredible 20% of the total animal biomass of this planet!


Watch the video: Amazing Electron Microscope Images (October 2022).