Identify this beetle from Bangladesh

Identify this beetle from Bangladesh

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Location: Bandarban, Bangladesh

This appears to be some species of beetle (order Coleoptera) in the Bostrichidae family.

  • Commonly called auger beetles, false powderpost beetles, or horned powderpost beetles.

Xylopsocus capucinus

Identification to species can be difficult, but fortunately for you Sittichaya et al (2009) created an illustrated key for IDing species associated with South Asia.

Based on their key and visual characteristics of your specimen, my guess is that your specimen is Xylopsocus capucinus

Source: Sittichaya et al 2009 . a. lateral view; b. elytral declivity; c. intercoxal process of the 1st abdominal ventrite; d. dorsal view.

  • Description: 3-5.5 mm long; 1.4-1.7 mm wide. Shape is cylindrical and similar in general appearance to other common species of the family Bostrichidae.

  • Distribution: throughout South and Southeast Asia from India to the Indonesian archipelago.

  • Hosts: apparently polyphagous attacking almost any woody plant in suitable condition.

  • Phenology: adults emerge mainly between May and November.

Like most false powderpost beetles, the head cannot be seen from above as it is downwardly directed and hidden by the thorax:

Source: UFL IAFS


  • Sittichaya, W., Beaver, R., & Ngampongsai, A. (2009). An illustrated key to powder post beetles (Coleoptera, Bostrichidae) associated with rubberwood in Thailand, with new records and a checklist of species found in Southern Thailand. ZooKeys, 26, 33.

  • Woodruff RE, Gerberg EJ, Spilman TJ (2005). A False Powder-post Beetle, Xylopsocus capucinus (Fabricius) (Insecta: Coleoptera: Bostrichidae). University of Florida, IFAS Extension Publication No. EENY-179. Institute of Food and Agricultural Sciences, University of Florida, 4 pp

This appears to be a Powder-Post Beetle (order Coleoptera, family Bostrichidae); a local reference is needed for further identification.

Identification, biology and management of Colorado potato beetle

This year, I’ve had a handful of people ask me about managing Colorado potato beetle (CPB) in Iowa. Read more about CPB identification and biology here. These conversations were with producers growing over 5 acres of potatoes and having a difficult time reducing CPB pressure. With the help of Dr. Ian MacRae at the University of Minnesota, I put together an IPM plan for this devastating pest. Colorado potato beetle (CPB) is a major pest of potato that is native to America and Mexico. It's been in Iowa for over 150 years and has a long history of devastating outbreaks. This article provides some basic information on CPB identification life cycle and damage to potato.

Colorado potato beetle. Photo by Wikipedia.


Adult CPBs are oval in shape and 3/8 inch long. They have a yellow-orange prothorax (the area behind the head) and yellowish white wing covers with 10 narrow black stripes. Females lay clusters of bright yellowish-orange oval eggs on the underside of leaves. Eggs turn dark red just before hatching. When larvae first hatch from eggs, they have brick red bodies with black heads. Older larvae are pink to salmon colored with black heads. All larvae have two rows of dark spots on each side of their bodies.

Colorado potato beetle eggs. Photo by Wikipedia.

Adult CPBs overwinter in potato fields, field margins and windbreaks. They become active in the spring at about the same time potatoes emerge (sometime in May). Adults feed for a short time in the spring and then begin to mate and lay clusters of 10-30 eggs on the undersides of leaves. Each female can lay up to 350 eggs over 3-5 weeks.

Eggs begin to hatch within two weeks, depending upon temperatures. Larvae remain aggregated­ near the egg mass when young but begin to move throughout the plant as they eat the leaves. Larvae can complete development in as little as 10 days if average temperatures are in the mid 80’s. It will take over a month if temperatures average near 60°F. Larvae mature through four instars before they drop from the plant, burrow into the soil and pupate. There can be two generations in Iowa. Because eggs are laid over time, all life stages of CPB can be present at the same time in a potato field by July.

Colorado potato beetle first instars. Photo by Wikipedia.

Both larvae and adults feed on the foliage of potatoes and, if left untreated, can completely defoliate plants. In addition to potato, they may also feed on eggplant, tomato, pepper, and other plants in the nightshade family (Solanaceae). Old larvae (i.e., 4th instars) are responsible for as much as 75% of feeding damage. Potatoes can usually tolerate substantial defoliation, up to 30%, when they are in the vegetative stage, but they are much more sensitive to the effects of defoliation when tubers are beginning to bulk and they can only tolerate about 10% defoliation. Tuber bulking begins soon after flowering, making this time critical for CPB management.

Colorado potato beetle third instar. Photo by Wikipedia.

In general, CPB are very difficult to suppress because of their biology. Successful suppression will take an integrated approach with a focus on being proactive. I often recommend using IPM (integrated pest management) to help with field crop pests. It should include some or all of the following tactics:

1. Cultural. Much of what farmers/gardeners do to grow plants favors insect development. If we can mix up our growing conditions to interfere with the food and habitat pests need to survive, that will greatly discourage them from devastating our crops. For example:

  • Sanitation, or removing potential food and habitat, is an effective starvation technique. If potatoes or solanaceous plants (e.g., eggplant, peppers, tomatoes), are not available when CPB adults first emerge in the spring, they will seek out alternate hosts, such as nightshade and ground cherry. Remove old plants and weeds in and around potato plots before, during and after the season to eliminate food sources.
  • For small plantings, hand removal can be effective. Drop CPB adults and larvae in a pail filled with soapy water. Also remove or crush the eggs on the underside of leaves. Adults can fly into plots so be sure to check your potatoes regularly. Hand removal may be less practical in larger plots.
  • Crop rotation, or only growing potatoes only every other year, may help reduce beetle populations if no potatoes are being grown within a radius of ¼-½ mile away and temperatures are not excessively warm. Moving the area that is planted with potato, is largely ineffective because the CPB can fly long distances when temperatures exceed 70°F.
  • Date of planting will probably not be that effective tactic like for other pests. Most potatoes will not germinate until the soil temp is 40°F and the adult tend to emerge 2-4 weeks later. Planting seed pieces too early can actually cause germination issues, like diseases and rotting.
  • Tillage is not effective unless deep tillage is used (and even then they can survive). The adults overwinter in soil and leaf litter and are quite capable of digging into and out of the soil after being buried.
  • Exclusion with cages or row covers is not recommended for CPB because the supplies are expensive and time consuming to maintain.

2. Genetic. Planting an early-maturing variety will allow you to escape much of the damage caused by adults emerging in midsummer. Check seed catalogs for varieties that mature in less than 80 days. Yields on early-maturing varieties are not as large, and often these varieties do not store as well as the popular Russet Burbank potato.

3. Biological. There are few natural enemies of CPB, like predatory stink bugs and beetles. There is also a naturally-occurring fungus, Beauveria bassiana, that will kill larvae and adults. Unfortunately, biological control has minimal impact on CPB populations compared to other crop pests. I wouldn’t recommend planting flowering crops around or within potato plots to attract natural enemies, or releasing predators to suppress CPB.

Colorado potato beetle infected with Beauveria bassiana, an insect killing fungus.

4. Scouting. To focus your suppression efforts, I highly recommend the use of tracking air temperature. Insects develop based on accumulating heat units, or degree days (DD) just like most plants and other invertebrates. The lower developmental threshold, a point at which no development will occur, for CPB is 52ºF. The very first adult you see in the spring sets the “biofix” where accumulating DD begins:

  • 120 dd post biofix - eggs are maturing, look for the beginning of 1st instars
  • 185 dd post biofix - 1st instars completing, look for the beginning of 2nd instars
  • 240 dd post biofix - 2nd instars completing, look for the first 3rd instars
  • 300 dd post biofix - 3rd instars completing, look for the first 4th instars
  • 400 dd post biofix - 4th instars completing, larvae will start dropping to ground to pupate

*You would want to target your scouting and suppression efforts on the first generation, 120-200 DD post biofix. Every summer is a bit different, so it is important to monitor each season rather than depend on calendar dates.

5. Thresholds. Even when implementing proactive IPM tactics, like cultural tools, there will be seasons when CPB still have outbreaks. In general, I recommend applying insecticides to protect yield based on CPB leaf defoliation: 20%-30% before flowering, 5%-10% at flowering, and 15%-20% post flowering. Estimating defoliation by eye takes practice because people tend to overestimate leaf area removed. There could be some seasons where these thresholds are not met and insecticides are not required.

Scouting for Colorado potato beetle eggs and young larvae, in addition to estimating defoliation, will inform your pest suppression plan and protect the crop!

6. Chemical. Depending on the size of the potato plots and the historical outbreaks on the farm, insecticides may be required to protect the crop. This method can be used with other IPM tactics and in moderation (e.g., a last resort when other tactics are not working). I highly recommend scouting and using thresholds to determine if applications are needed (see previous sections). There is a wide spectrum of chemical control products that target CPB, ranging from organic to restricted-use pesticides. It is important to read the label and follow all the directions, particularly how soon you can harvest potatoes after applications. Keep the following considerations in mind:

  • It is easier to kill young larvae compared to older larvae and adults.
  • Products are more effective when making direct contact with the body. Getting small droplets to contact CPB can be challenging given they feed and aggregate on the undersides of leaves.
  • Targeting suppression efforts of the first generation can greatly reduce the impact of the second generation when plants are more sensitive to defoliation.
  • Most chemicals are broad spectrum, meaning they will kill most insects that make contact with the application (e.g., pollinators, predators, etc.). This is true of organic and synthetic insecticides.

Microbial insecticides (the following products are approved for organic production and considered low risk to humans):

Common and Unusual Identifications - Beetles

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We get a lot of questions about beetles. This is probably due to the fact that beetles are the largest group of animals on Earth. There are more than 30,000 known species in Australia and many more yet to be discovered. Conservative estimates of beetles worldwide is 350 000 species. Beetles can vary in size from tiny, just a fraction of a millimetre to huge, 160 millimetres long. Their sheer numbers, diversity and beauty make them an often sighted animal.

Beetles belong to the order Coleptera, which means 'sheath wings'.

Like almost all insects, beetles have 3 body parts- a head, thorax and abdomen. they have 3 pairs of legs and 2 pairs of wings. Beetles' forewings are hardened into sheath like protective coverings called elytra that protect their delicate hindwings.

Beetles tough exoskeleton prevents water loss allowing them to live in almost every environment. They can be found on snow covered mountains, in harsh deserts, deep dark caves, hot springs, underwater, backyards and urban environments.

All beetles have a life cycle that includes a grub, pupa, and adult stage.

Check how many of these beetles you have come across.

Botany Bay weevil

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Image: Bruce Hulbert
© Bruce Hulbert

Commonly known as the Botany bay or Diamond weevil, Chrysolopus spectabilis, was collected by Joseph Banks on his voyage with captain cook and was the first insect to be scientifically described and named from Australia. It is commonly found on wattles in coastal woodlands and forests where the adults feed on the foliage.

Christmas beetle

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Image: Stanley Breeden
© Australian Museum

Christmas beetle, Anoplognathus sp., are often seen as their name suggests, at the close of the year from November to January. The adults feed on both young and mature gum tree foliage, while the larvae are underground feeding on organic matter and the plant roots of grasses. The mated females burrow 5-10 centimetres into soil to lay about 40 eggs in small cavities. These beetles come in a range of colours according to species and usually have a metallic o green or golden sheen on their bodies.

Discover the wonderful diversity of Australia's most famous beetles with Australian Museum's free mobile app - Xmas Beetle ID Guide iOS app, Xmas Beetle ID Guide Android app developed by Australian Museum entomologists Dr Chris Reid and Mike Burleigh.

Fiddler beetle

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Fiddler beetles, Eupoecila australasiae, get their name from the patterns on the body resembling those of a fiddle or violin. They can be found feeding on flower nectar and pollen of flowering gum trees and other native trees, as well as rotting fruit during the summer months. They are active day flyers the females seek out areas where there are rotten wood, mulch or other vegetable compost to lay their eggs which will hatch into curled larvae.

The spotted flower chafer, Neorrhina punctatum, (below) is a related species.

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Image: Peter Firus
© Peter Firus

Cowboy beetle

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Image: Bruce Hulbert
© Bruce Hulbert

Cowboy beetles, Chondropyga dorsalis, are relatively common in Eastern Australia. Found in dry forests, woodlands and suburban areas, they are most common in coastal areas. These orange- brown to mid brown beetles have a wide black stripe down the middle of the thorax and abdomen, the patterns are variable in shape and size. Females are slightly larger than the males and the adults emerge from their pupal cell in early summer in all the eastern states and territories.

An Australian native, the commonly named cowboy beetle, is a species of scarab beetle. They are not dangerous and not even considered as pests. The larvae feed on dead and decaying wood and the adults feed on nectar.

We often are asked to identify the grubs and adults of this common native beetle.

In a plant pot this species at the grub stage of its life cycle will cause growth reduction to your plants as they will eat the roots and are quite destructive. The Female adult beetles are attracted to the organic matter in the potting mix and will lay her eggs there. The eggs will pupate underground. The large white C shaped grubs will then feed on decaying wood using strong mouth parts on its tiny orange head.

In a garden bed this would be fine, but in a pot, these grubs eat all the food supply and then start on the roots. You will notice the soil in pot will looked turned up. In terms of control, If the pots are of a convenient size, tip all the plant contents out onto a large plastic sheet or tray. Save aside all the plants in a pile. Then sort through the soil and pick out all the grubs (release them to a section of your yard allowing them to emerge as beetles). Replace everything back into the pot and replant. You will be amazed in the sudden growth as the grubs will have broken down all the potting mix, making it ideal plant food and now the plant roots can grow again.

The most common host in the Black Hills is ponderosa pine. This tree occurs on more than 1 million acres of forestland in western South Dakota. Ponderosa pine is also extensively planted in shelterbelts and landscapes. Ponderosa pine can be separated from other pines by its bundles of needles in two’s and three’s (most other pine trees will have needles in bundles of only two or only three). The needles are typically 5 to 11 inches long.

Lodgepole, sugar and western white pine, though far more common in western states, are also susceptible wherever they are found. The pines we frequently use in the ornamental landscape, Scots (Scotch) and Austrian pine, are generally not hosts due to their smaller size and distance from mountain pine beetle infestations. However, these trees are highly susceptible to attack and large trees near infestations are vulnerable. Spruce, fir or Douglas-firs may be attacked if in the vicinity of infestations but these attacks are rarely successful.

Identify this beetle from Bangladesh - Biology

Oryctes rhinoceros (L.), the coconut rhinoceros beetle, is a pest species occurring throughout many tropical regions of the world. Adults can cause extensive damage to economically important wild and plantation palms.

Figure 1. Palm damaged by Oryctes rhinoceros. Photograph by Mark Benavente.

Oryctes rhinoceros is one of the most damaging insects to palms in Asia and the Pacific Islands. Adults eat the leaves and burrow into the crown, stunting plant development (Giblin-Davis 2001).

Distribution (Back to Top)

Oryctes rhinoceros is native to Asia, between India and Indonesia. It has since spread to Yemen, Reunion, and Hawaii. Throughout this distribution, the coconut rhinoceros beetle is most closely associated with its preferred host plant, Cocos nucifera L., the coconut palm (Hinckley 1973).

Figure 2. Distribution of Oryctes rhinoceros, based on published distribution records. Image by Mike Dornberg, Florida Department of Agriculture and Consumer Services, Division of Plant Industry.

Description (Back to Top)

Although Oryctes rhinoceros is found in several regions of the world, its shape, size and color are generally consistent (Manjeri et al. 2013). Adult beetles range from 1.2 to 2.5 inches in length (3.0 to 6.3 cm) and are dark brown or black. The ventral surface (underside) of males and females has reddish-brown hairs, but the female has a fuzzy grouping of these hairs at the tip of the abdomen. Both males and females possess a similarly sized horn used for leverage when moving within tightly-packed leaves or within the cavities they create in the crown of palms, the horn length is longer on average for males (Doane 1913).

Figure 3. Female (left) and male (right) Oryctes rhinoceros. In this picture, the head of the female is up while the head of the male is down, displaying an exaggerated difference in horn length. Photograph by Mike Dornberg, Florida Department of Agriculture and Consumer Services, Division of Plant Industry.

Oryctes rhinoceros larvae (grubs) are milky white with red heads. The body is C-shaped, has three pairs of segmented legs, and is grayish posteriorly. Over the course of three instars, or phases between molts, they grow to 4.0 inches long (10.0 cm).

Figure 4. Oryctes rhinoceros larva. Photograph by Aubrey Moore, University of Guam.

Biology (Back to Top)

Adult females deposit eggs inside dead palms, decaying plant material, soil with high organic matter content, and, occasionally, wooden structures (Manjeri et al. 2014). In approximately 11 days, eggs hatch into larvae which begin feeding on surrounding organic material. Eleven to 15 weeks later, the larvae will have grown up to 16 times larger and have stopped eating, after which they enter the pupal stage and are immobile for approximately six weeks (Hickley 1973). Upon emerging, adults fly to a new tree, feed, and mate, sometimes mating just after their first feeding. Adults spend most of their time feeding on fresh leaves. Adult females live up to nine months, over which period they can lay up to 100 eggs. Thus adult progeny may be present with the mother and the population consists of overlapping generations (Manjeri et al. 2014).

Multiple overlapping generations are common under favorable conditions, e.g. when no diapause is needed. Since coconuts occur in regions where there is no cold season and a minimal dry season, the beetles can be active and reproductive throughout the year.

Host Plants (Back to Top)

As with many beetles, adults and larvae have different feeding preferences. In the case of Oryctes rhinoceros, damage to plants is caused by adults (especially young adults) and not larvae, which feed on already rotting material (Giblin-Davis 2001).

Larvae live in decaying material including: Cocos nucifera, Artocarpus sp. (breadfruit), Calophyllum inophyllum (Alexandrian laurel), Mangifera sp. (mango), and Pandanus sp. (Gressitt 1953).

Adults are a major pest of Cocos nucifera (coconut palm) and Elaeis guineensis (African oil palm) (Giblin-Davis 2001) but are a minor pest of many other palms and plant species. By feeding on healthy leaves, Oryctes rhinoceros causes physical damage, which can stunt growth and lead to secondary infections from bacteria or fungi (Hinckley 1973).

Minor host plant species include:

Acanthophoenix rubra (barbel palm) Corypha umbraculifera (talipot palm) Pandanus tectorius (Tahitian screwpine)
Agave sisalana (sisal agave) Corypha utan (buri palm) Phoenix dactylifera (date palm)
Agave americana (American agave) Cyathea sp. (tree fern) Phoenix sylvestris (wild date palm)
Aiphanes horrida (ruffle palm) Dictyosperma album (red palm) Pinanga sp.
Ananas comosus (pineapple) Dypsis pinnatifrons Pinanga insignis
Areca sp. (areca palm) Heterospathe elata var. palauensis Pritchardia pacifica (Fiji fan palm)
Areca catechu (betel-nut palm) Hydriastele palauensis Raphia farinifera (raffia palm)
Arenga sp. (arenga palm) Hyophorbe lagenicaulis (bottle palm) Raphia vinifera (bamboo palm)
Arenga pinnata (sugar palm) Latania sp. Roystonea regia (royal palm)
Borassus sp. (borassus palm) Livistona chinensis (Chinese fan palm) Saccharum sp. (sugarcane)
Borassus flabellifer (palmyra palm) Metroxylon amicarum (Caroline ivory-nut palm) Syagrus romanzoffiana (queen palm)
Caryota urens (fish-tail palm) Metroxylon sagu (sago palm) Thrinax sp. (thatch palm)
Casuarina equisetifolia (Australian pine) Musa sp. (banana) Verschaffeltia splendida (Seychelles stilt palm)
Clinostigma samoense Normanbya sp. Wodyetia bifurcata (foxtail palm)
Colocasia sp. (taro) Nypa fruticans (nipa palm)
Corypha sp. (gebang palm) Oncosperma sp.

(Gressitt 1953 Lever 1969 Elfers 1988 Giblin-Davis 2001 Quitugua 2010)

Economic Importance (Back to Top)

Malaysia: Adult Oryctes rhinoceros cut through leaves and bore holes into palm crowns. Damage is exacerbated by the tendency of beetles to aggregate. Production loss in plantations in Malaysia has averaged 40%, but has reached 92% (Manjeri et al. 2013).

Paulau: Oryctes rhinoceros was found in Palau in 1942 and caused an overall tree mortality of 50% (Gressitt 1953).

Guam and Hawaii: In 2007 Oryctes rhinoceros was declared established in Guam. As of 2012, the coconut palm was Guam's second most abundant tree (Moore 2012). Moore (2007) had previously concluded that "accidental transport of other scarab beetles from Guam to Hawaii is well documented." In 2013, the coconut rhinoceros beetle was found in Hawaii (Hara 2014).

North America: Coconut rhinoceros beetle is not established in the mainland United States however, the risk of accidental transport remains in our increasingly connected world. If you suspect that you have found this beetle, immediately contact your local state agency. In Florida, contact the Florida Department of Agriculture and Consumer Services 1-800-HELP-FLA (1-800-436-7352).

Management and Control (Back to Top)

Detection can be difficult due to the beetle&rsquos nocturnal activity and residence within trees. Visual signs such as holes bored at the base of leaves and V-shaped feeding damage help locate this beetle. Recently, acoustic detection was used to find Oryctes rhinoceros in live and dead palms on Guam (Mankin and Moore 2010). Once detected, management and control are required to mitigate the economic impact of a beetle infestation.

Figure 5. Typical V-shaped damage to coconut leaves by Oryctes rhinoceros. Photograph by Ben Quichocho, USDA-APHIS.

Control: Historical control of scarab beetle pests has included chemical and biopesticides, biological control (predators, parasitoids, and pathogens), and trapping with lures (Jackson and Klein 2006). Traps for the coconut rhinoceros beetle contain 4-methyloctanoate, an aggregation pheromone produced by the male beetle. In breeding sites, the fungus Metarhizium anisopliae may be applied for larval control and is distributed by adults. This fungus acts as a biopesticide on immature stages of the beetle (Bedford (2014).

Viruses in the genus Nudivirus have also been associated with Oryctes rhinoceros and may play a role in controlling the beetle in regions where they are invasive (Bedford 2014). Infection by Oryctes rhinoceros nudivirusdeforms and may kill larvae, and hinders oviposition by females (Bedford 2014). However, the nudivirus may harm other species and genera of scarabs. In Korea, farmers of Allomyrina dichotoma (Japanese rhinoceros beetle) face a potential disaster if the nudivirus were to infect their populations, which consist of hundreds of larvae grown together in large plastic containers. These beetles are cultivated for sale as pets and to be used in gambling. The result of a Korean study in which Allomyrina dichotoma larvae were infected with the nudivirus was that 60% died in six weeks. There is also concern that the nudivirus may be transmitted to wild populations of Allomyrina dichotoma. There is no clear indication that the Oryctes rhinoceros nudivirus is the major pathogen responsible for losses of Allomyrina dichotoma in Korea and tests are ongoing (Lee et al. 2015).

Management: Managing the coconut rhinoceros beetle involves removing or destroying organic material that supports larval development such as decaying logs and stumps, removing dead palms, and removing piles of leaves and grass (Schmaedick 2005). A study on burning the downed and decomposing trunks of oil palms has shown that only partial burning of sites is ineffective in managing population levels of Oryctes rhinoceros (Abidin et al. 2014).

Selected References (Back to Top)

  • Abidin CMRZ, Ahmad AH, Salim H, Hamid NH. 2014. Population dynamics of Oryctes rhinoceros in decomposing oil palm trunks in areas practicing zero burning and partial burning. Journal of Oil Palm Research 26: 140-145.
  • Bedford GO. 2014. Advances in the control of rhinoceros beetle, Oryctes rhinoceros in oil palm. Journal of Oil Palm Research 26: 183-194.
  • Doane RW. 1913. How Oryctes rhinoceros, a dynastid beetle, uses its horn. Science, New Series 38: 883.
  • Elfers SC. 1998. Abstract for Casuarina equisetifolia, Australian pine. The Nature Conservancy: 9-10.
  • Giblin-Davis R. 2001. Borers of palms. Insects on palms. CABI Publishing, Wallingford Great Britain: 297-300.
  • Gressitt JL. 1953. The coconut rhinoceros beetle (Oryctes rhinoceros) with particular reference to the Palau Islands. Bulletin of the Bernice P. Bishop Museum 212: 157.
  • Hara AH. 2014. Coconut rhinoceros beetle, Oryctes rhinoceros: a major threat to Hawaii&rsquos coconut and palm trees. University of Hawaii at Manoa Crop Production Services Seminar & Tradeshow.
  • Hinckley AD. 1973. Ecology of the coconut rhinoceros beetle, Oryctes rhinoceros (L.) (Coleoptera: Dynastidae). Biotropica 5: 111-116.
  • Jackson TA, Klein MG. 2006. Scarabs as pests: a continuing problem. Coleopterists Society Monographs 5: 102-119.
  • Lee S, Park KH, Nam, SH, Kwak KW, Choi JY. 2015. First report of Oryctes rhinoceros nudivirus (Coleoptera: Scarabaeidae) causing severe disease in Allomyrina dichotoma in Korea. Journal of Insect Science 15: 26.
  • Lever RJAW. 1969. Pests of the coconut palm. Food and Agriculture Organization of the United Nations: 125-133.
  • Manjeri et al. 2013. Morphometric analysis of Oryctes rhinoceros (L.) (Coleoptera: Scarabaeidae) from oil palm plantations. The Coleopterists Bulletin 67: 194-200.
  • Manjeri et al. 2014. Oryctes rhinoceros Beetles, an oil palm pest in Malaysia. Annual Research and Review in Biology 4: 3430-3439.
  • Mankin RW, Moore A. 2010. Acoustic detection of Oryctes rhinoceros (Coleoptera: Scarabaeidae: Dynastinae) and Nasutitermes luzonicus (Isoptera: Termitidae) in palm trees in urban Guam. Journal of Economic Entomology 103: 1135-1143.
  • Moore A. 2007. Assessment of the rhinoceros beetle infestation on Guam. University of Guam, Western Pacific Tropical Research Center.
  • Moore A. 2012. Guam as a source of new insects for Hawaii. Pacific Entomology Conference. University of Hawaii, Cooperative Extension Services Western Pacific Research Center.
  • Quitugua R. 2010. Rhino beetles take aim at new palm species, an interview with Eradication Project Logistics Manager for the Department of Agriculture, Roland Quitugua. Kuam News Network, Guam.
  • Schmaedick M. 2005. Coconut Rhinoceros Beetle. American Samoa Community College Community & Natural Resources Cooperative Research. Pests and Diseases of American Samoa number 8.

Author: Mike Dornberg, Division of Plant Industry, Florida Department of Agriculture and Consumer Services
Photographs: Mark Benavente, Mike Dornberg, Division of Plant Industry, Aubrey Moore, University of Guam, Ben Quichocho, United States Department of Agriculture, Animal and Plant Health Inspection Service.
Graphics: Mike Dornberg, Division of Plant Industry
Publication Number: EENY-629
Publication Date: July 2015.

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

Japanese beetle, Popillia japonica Newman, is an invasive insect to the United States. This beetle was first discovered in New Jersey in the early 1900s and has since successfully spread across much of the eastern United States. In its invaded range, Japanese beetle has become a significant insect pest of turfgrass and ornamental, horticultural and agricultural plants. The significance of Japanese beetle is increasing throughout more recently infested areas. Currently, the United States is the world’s top producer of both corn and soybean, with most of this production occurring in the Midwest. Due to the recent activity of Japanese beetle in the Midwest and this insect’s ability to injure soybean and corn, this review will focus on relevant information regarding Japanese beetle identification and biology, and its impacts and management in corn and soybean.

Japanese beetle, Popillia japonica Newman, has become a significant insect pest of turfgrass and ornamental, horticultural and agricultural plants in the eastern United States ( Potter and Held 2002). The pest status of Japanese beetle is due in part to its generalist nature, feeding on more than 300 plant species, as well as the ability to form large aggregations ( Smith and Hadley 1926, Fleming 1972, Potter and Held 2002).

As larvae, Japanese beetles are destructive to turfgrass roots, including lawns, golf courses, and athletic fields ( Potter 1998, Vittum et al. 1999). Adults feed mainly on leaves of plants, eating between the veins and leaving a characteristic skeletonized appearance. Once established, Japanese beetles can be a difficult and expensive insect pest to control, estimated at approximately $450 million each year in the United States for turfgrass management alone ( USDA-NASS 2016). The significance of this invasive species in the Midwestern United States is increasing ( Hudson 2017, Severson 2017), as modeling shows suitable climate space for Japanese beetle remains unoccupied in the area ( Zhu et al. 2017).

The United States is the world’s top producer of both corn, Zea mays L. (Poales: Poaceae) and soybean, Glycine max (L.) Merr. (Fabales: Fabaceae), yielding 14.6 and 4.39 billion bushels, respectively, in 2017, with most of this production occurring in the Midwest ( USDA-NASS 2018). Due to recent activity of Japanese beetle in the Midwest and this insect’s potential to injure soybean and corn ( Hammond 1994, Edwards 1999), this review will focus on relevant information regarding its identification and biology, and its impacts and management in corn and soybean.

History and Current Distribution in the United States

Japanese beetle is native to northern Japan ( Fleming 1976), where it is considered a minor agricultural pest due to the combination of coevolved natural enemies and unsuitable terrain for larval development ( Clausen et al. 1927). In the United States, Japanese beetle was first found in 1916 at a nursery near Riverton, New Jersey and is speculated to have arrived via imported rhizomes of Japanese iris, Iris ensata Thunb. (Asparagales: Iridaceae) ( Dickerson and Weiss 1918). Since its discovery, the beetle spread westward across the country with great success, presumably due to favorable groundcover, such as turfgrass, for larval development, adequate rainfall and human-assisted movement ( Potter and Held 2002). Currently, Japanese beetle is considered established in 28 states and five Canadian provinces, with six other states partially infested ( CFIA 2016, USDA-APHIS 2018 Fig. 1).

Map of Japanese beetle distribution in the United States as of 2018 and Canada as of 2016. Graphic by Hailey Shanovich, adapted from CFIA 2016 and USDA-APHIS 2018.

Map of Japanese beetle distribution in the United States as of 2018 and Canada as of 2016. Graphic by Hailey Shanovich, adapted from CFIA 2016 and USDA-APHIS 2018.


Newly deposited eggs are laid singly and can be generally found at a depth of up to 4 inches (10 cm) ( Dalthorp et al. 2000). Eggs vary slightly in size and shape. Most often, eggs are spherical with a diameter of 1 /16 inch (1.5 mm), but they can be elliptical, measuring 1 /16 inch (1.5 mm) long and < 1 /16 inch (1 mm) wide. Color ranges from translucent to creamy white ( Fig. 2a) ( EMPPO 2006, USDA-APHIS 2015).

Life cycle of the Japanese beetle, including (a) eggs David Cappaert, Michigan State University,, (b) larvae (grubs) David Cappaert, Michigan State University,, (c) pupa(USDA-APHIS), and (d) adult (Theresa Cira, University of Minnesota).

Life cycle of the Japanese beetle, including (a) eggs David Cappaert, Michigan State University,, (b) larvae (grubs) David Cappaert, Michigan State University,, (c) pupa(USDA-APHIS), and (d) adult (Theresa Cira, University of Minnesota).


Larvae are C-shaped white grubs with a yellowish-brown head ( Fig. 2b). Larvae have chewing mouthparts, three pairs of thoracic legs, and 10 abdominal segments ( EMPPO 2006). The bodies of the larvae are covered with brown hairs concentrated on the dorsal (top) side and at the tip of the abdomen ( Potter et al. 2006). The ventral (bottom) side of the last abdominal segment bears two diagnostic V-shaped rows of six or seven spine-like hairs, which may be used to distinguish larvae of this species from other scarab species ( Sim 1934 Fig. 3). There are three larval stages (or instars) of Japanese beetle. Newly-hatched larvae, or first instars, are up to 1 /8 inch (3 mm) long, while fully-grown larvae, or third instars, are about 1 3 /16 inches (30 mm) long ( Isaacs et al. 2003). The second and third instars can be distinguished by head capsule size. The second instar’s head capsule is about ½ inch (1.2 mm) long and ¾ inch (1.9 mm) wide, while that of the third instar is 5 /64 inch (2.1 mm) long and 1 /8 inch (3.1 mm) wide ( EMPPO 2006).

Spines on the tip abdomen are important for white grub diagnostics, (a) as shown here (John C. French Sr., Retired, Universities: Auburn, GA, Clemson and U of MO, (b) Japanese beetle grubs have spines in a characteristic V-shaped pattern (Jeff Hahn and Hailey Shanovich, University of Minnesota) (c) Northern masked chafers have an irregular pattern of spines (Mike Reding and Betsy Anderson, USDA-ARS) and (d) May/June beetles have a zipper-like arrangement of spines (Jeff Hahn and Hailey Shanovich University of Minnesota).

Spines on the tip abdomen are important for white grub diagnostics, (a) as shown here (John C. French Sr., Retired, Universities: Auburn, GA, Clemson and U of MO, (b) Japanese beetle grubs have spines in a characteristic V-shaped pattern (Jeff Hahn and Hailey Shanovich, University of Minnesota) (c) Northern masked chafers have an irregular pattern of spines (Mike Reding and Betsy Anderson, USDA-ARS) and (d) May/June beetles have a zipper-like arrangement of spines (Jeff Hahn and Hailey Shanovich University of Minnesota).


Pupae are exarate (having legs that are not held closely to the body and resembling the adult). Body color ranges from cream to tan and eventually metallic-green just before adult emergence ( Fig. 2c). The pupae are about ½ inch (14 mm) long and ¼ inch (7 mm) wide ( EMPPO 2006).


Adult Japanese beetles have brightly colored metallic-green bodies with coppery-bronze elytra (i.e., wing coverings) ( Fig. 2d). Along the sides of the body are tufts of white setae (hair) and two spots of white setae on the back end. Their bodies are oval in shape and vary in size from 5 /16 to 7 /16 inch (8 to 11 mm) long and 13 /64 to ¼ inch (5 to 7 mm) wide ( Hammond 1994, Edwards 1999). The females are generally larger than the males ( USDA-APHIS 2015). Males and females can be differentiated from each other by the shape of the tibia (part of the leg) and tarsus (foot) on the pair of legs nearest the head ( Fig. 4a). Males have more sharply-pointed tibial spurs and shorter tarsi than females ( Fig. 4b). Adults may be confused with other species of beetles that can co-occur on many of their host plants ( EMPPO 2006 Fig. 5).

Distinguishing tibial spines of Japanese beetle (a) males and (b) females. Photos by Tom Hillyer.

Distinguishing tibial spines of Japanese beetle (a) males and (b) females. Photos by Tom Hillyer.

Japanese beetle look-alikes, including (a) six-spotted tiger beetle (David Cappaert, Michigan State University, (b) dogbane beetle (David Cappaert, Michigan State University, (c) false Japanese beetle (Erin Hodgson) (d) northern masked chafer (Mike Reding and Betsy Anderson, USDA-ARS, (e) May beetle (Emmy Engasser, USDA-APHIS PPQ, (f) shiny leaf chafer (Whitney Cranshaw, Colorado State University) and (g) June beetle (Steven Katovich, USDA Forest Service,

Japanese beetle look-alikes, including (a) six-spotted tiger beetle (David Cappaert, Michigan State University, (b) dogbane beetle (David Cappaert, Michigan State University, (c) false Japanese beetle (Erin Hodgson) (d) northern masked chafer (Mike Reding and Betsy Anderson, USDA-ARS, (e) May beetle (Emmy Engasser, USDA-APHIS PPQ, (f) shiny leaf chafer (Whitney Cranshaw, Colorado State University) and (g) June beetle (Steven Katovich, USDA Forest Service,


Seasonal Biology and Reproduction

Throughout most of its range in the United States, Japanese beetle has an annual life cycle (one generation per year). In the Midwest, adults begin emerging from the soil in mid-to-late June to early July ( Hammond 1994, Edwards 1999, Hodgson 2018, MDA 2018), with females probably emerging a few days earlier than males ( Van Timmerman et al. 2001). Emerging females carry an average of 20 mature eggs and are thought to release a sex pheromone, (Z)-5-(1-decenyl)dihydro-2(3H)-furanone, to attract males, mate and lay eggs before initiating feeding ( Tumlinson et al. 1977, Ishida and Leal 2008).

Following the initial oviposition, females fly to host plants and begin feeding ( Barrows and Gordh 1978). Throughout their adult lifespan of 4 to 6 wk, females will continually alternate between feeding, mating and ovipositing eggs. They will enter the soil a dozen or more times, laying up to 60 individual eggs ( Fleming 1972). If the host-plant females are feeding on is adjacent to a suitable oviposition site, the females will usually lay eggs there ( Gould 1963, Fleming 1972) otherwise, they will disperse to more suitable sites ( Régnière et al. 1983). Sites with short grass cover ( Hawley 1944, Smitley 1996, Szendrei and Isaacs 2005, Wood et al. 2009) along with high soil moisture, moderate soil texture ( Allsopp et al. 1992), low organic matter content ( Dalthorp et al. 2000), and sunlight ( Dalthorp et al. 1999) are preferred by females for oviposition, although eggs may also be laid within crop fields, with soybean seemingly preferred to corn ( Gould 1963, Hammond 1994, Edwards 1999). In addition, tillage practices ( Smith et al. 1988) influence the number of eggs laid by females in crop fields, with higher densities in reduced-tillage systems ( Hammond and Stinner 1987), and determine the proximity of the oviposition site to the feeding site ( Régnière et al. 1983). Oviposition cues, therefore, are detected by females on the surface as well as within the soil, providing further indication of habitat suitability ( Szendrei and Isaacs 2005, 2006 Wood et al. 2009).

Eggs typically hatch within 10 to 14 d, and development of the first and second instars requires 2 to 3 wk and 1 wk, respectively, and larvae reach the third instar in autumn ( Fleming 1976). The larvae are restricted to feeding on plant roots and decaying vegetation wherever they hatch, due to their limited mobility through the soil ( Potter and Held 2002). Therefore, the adult female’s oviposition site selection is crucial for larval survival and determining subsequent populations, which can lead to patchy distributions with mean population densities low in some areas and high in others throughout a given landscape ( Dalthorp et al. 1999). The development and survival of newly hatched larvae are greatly dependent on soil moisture and temperature, with larvae being susceptible to extreme heat, cold, and drought ( Régnière et al. 1981, Petty et al. 2015). Third instars will feed into October and begin to move deeper into the soil profile, typically up to 6 inches below the soil surface, for overwintering ( Fleming 1976). Overwintering larvae are considered susceptible to freezing ( Fleming 1976). Diapause ends the following spring when soil temperatures in the upper 6 inches (15 cm) exceed 50°F (10°C), and grubs begin to move back upward in the soil profile to continue feeding for another 4 to 8 wk before pupating ( Vittum 1986). The pupal stage lasts 7 to 17 d and the newly-molted adults remain in the soil for another 2 to 14 d prior to emergence, which is also highly dependent on soil temperature ( Fleming 1976, Régnière et al. 1981).

Chemical Ecology and Host Suitability

Japanese beetle is a generalist herbivore that attacks foliage, flowers, and fruits of more than 300 wild and cultivated plant species in 79 families ( Fleming 1972 Ladd 1987b, 1989). Feeding on different species and/or cultivars of host plants can dramatically increase the longevity and fecundity of adults ( Ladd 1987a, Spicer et al. 1995). Despite its extreme generalist feeding, Japanese beetle shows distinct preferences for certain plant species, whereas other plant species are rarely or never fed upon.

Host-plant quality, characterized by levels of primary metabolites and presence of secondary metabolites, determines feeding by an insect ( Jaenike 1990, Bernays and Chapman 1994, Patton et al. 1997). With Japanese beetle, non-host plants are thought to be identified mainly by the presence of feeding deterrents rather than feeding stimulants in host plants ( Potter and Held 2002, Adesanya et al. 2016). This may also determine resistance in closely-related plants. However, specific mechanisms of resistance among varieties need more exploration. There is building evidence for Japanese beetles’ selectivity among varieties documented in soybean ( Chandrasena et al. 2012) grape, Vitis L. spp. (Vitales: Vitaceae) ( Gu and Pomper 2008, Hammons et al. 2010) Prunus L. spp. (Rosales: Rosaceae) ( Patton et al. 1997) Asian elm, Ulmus L. spp. (Rosales: Ulmaceae) ( Paluch et al. 2006) maple, Acer L. spp. (Sapindales: Sapindaceae) (Seagraves et al. 2013) and birch, Betula spp. (Fagales: Betulaceae) ( Gu et al. 2008). Specific secondary plant defense compounds have been identified as feeding deterrents for the Japanese beetle in tree fruit and are valuable for the development of cultivars but have not yet been identified or explored in field crops ( Fulcher et al. 1996, Patton et al. 1997). However, there have been a few resistance genes and quantitative trait loci identified that confer some resistance to Japanese beetle in soybean (reviewed in the management section) ( Yesudas et al. 2010).

Other factors are also known to influence the level of plant susceptibility to feeding by adult Japanese beetles, including flower color ( Fleming 1972) and the amount of sunlight the plant receives ( Potter et al. 1996 Rowe and Potter 1996, 2000). In addition, Japanese beetles tend to aggregate and feed most in the upper canopy of plants, defoliating them from the top down ( Rowe and Potter 1996). Therefore, Japanese beetle feeding tends to occur on full-sun plants with a top–down feeding pattern (Rowe and Potter 1996, 2000 Zavala et al. 2009), which may in part be due to the sugar content of upper leaves. Plants with higher amounts of sugar have been found to have greater damage from Japanese beetles than those with lower sugar concentrations ( Ladd 1986, Patton et al. 1997). Sugar concentrations vary considerably with levels of light and typically increase during the day therefore, sunlit plants experience higher natural sugar content ( Bernays and Chapman 1994), increasing their chance for attack.

It has been observed that Japanese beetle feeding aggregations are initiated by ‘pioneer’ females feeding on the leaves, with males joining later in the process, leading to later gregarious mating and feeding, causing damage to plants ( Kowles and Switzer 2011). Feeding-induced plant volatiles, not female sex pheromones, are thought to attract more beetles to the feeding aggregations on plants ( Ladd 1970 Loughrin et al. 1995, 1996a, 1996b, 1997, 1998). Feeding aggregations have also been found to be male-biased, which may be created by females leaving aggregations after a shorter period of time than males to oviposit and colonize new plants afterward ( Switzer et al. 2001, Tigreros and Switzer 2009, Kowles and Switzer 2011). However, both male and female beetles are thought to move around frequently, flying to nearby plants or to different locations on the same plant ( Fleming 1972), with one study estimating that about one-third of the beetles alighting on a tree left the tree during the same day ( Van Leeuwen 1932).

Impact on Crops

Because of their generalist nature, Japanese beetle adults frequently feed on many crops, and have the potential to damage corn and soybean ( Hammond 1994, Edwards 1999). In the Midwestern United States, Japanese beetle has not historically been an economically important pest. However, as they increase in number and expand their range in the Midwest, farmers may need to consider scouting and managing this emerging pest that can feed on a number of plant parts on various crop species. In soybean, for example, Japanese beetle has become an important member of the guild of defoliating pests ( Hammond 1994, Steffey 2015, Hurley and Mitchell 2016).

Japanese beetle adults are easily detected in the field because of their relatively large size, metallic-green color and characteristic damage to plants. In contrast, the inconspicuous soil-dwelling larvae, which feed on roots of short grass species, are not considered as much of a concern as the adults in crops ( Hammond 1994, Edwards 1999). Depending on the level of feeding, root damage can have effects on plant standability and water and nutrient uptake of seedlings ( Potter et al. 2006). Larvae are relatively immobile in the soil therefore, the level of damage depends on their location relative to the plant.

Injury to Corn

Japanese beetle adults are a sporadic pest of corn ( Edwards 1999). Although the adults can feed on corn leaves, the main concern is the clipping of silks ( Fig. 6), which can interfere with pollination, leading to ears with a reduced set of kernels ( Edwards 1999). Feeding on exposed kernels can happen but is also less of a concern ( Edwards 1999). Risk of infestation of cornfields is greater for fields following sod, cover crops, or soybean as soybean is thought to be more attractive for oviposition by females ( Gould 1963, Dewerff et al. 2019a). These environments present ideal conditions for oviposition for the overwintering generation. In addition, nearby soybean fields can serve as a source of Japanese beetles adults in corn ( Edwards 1999).

Japanese beetle injury to corn, including (a) early feeding and (b) severe silk clipping. Photos by Erin Hodgson.

Algae and Other Microorganisms

Volvox (green)Haematococcus (red colored)dinoflagellate (red, green, some multicolored)

Desmids (green)Spirogyra (green)

Anything green and stringy can be classified as an algae.

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Adult corn rootworm identification

Corn rootworm adult emergence is underway in Iowa. The three species of rootworm found in Iowa include the northern corn rootworm (NCR), southern corn rootworm (SCR), and western corn rootworm (WCR). Adults of all three species can be found until the first frost. Knowing how to distinguish the three species is important for making management decisions for future growing seasons.

Southern corn rootworm (left), western corn rootworm (middle), and northern corn rootworm (right). Photo courtesy of Adam J. Varenhorst.

Adult NCR adults are typically one solid color, but their coloration can range from light green to dark green. NCR are approximately 1/4 inch long and do not have markings on their hardened forewings.

Northern corn rootworm. Photo courtesy of Adam J. Varenhorst.

Though mated females lay eggs in cornfields, adult NCR will readily move to other plants. There is one generation of northern corn rootworm per year however, some NCR populations survive as eggs in the soil for multiple years. This characteristic is referred to as extended diapause, and is an adaptation to crop rotation. Injury to first-year corn by extended-diapause NCR occurs most commonly in the western two thirds of Iowa, but extended diapause may be found elsewhere Iowa.

Adult WCR are typically slightly larger than NCR and are yellow in color with three dark stripes running lengthwise on their hardened forewings. These stripes can vary from three distinct lines to one large stripe covering most of the forewing.

Western corn rootworm (the three on the right are males). Photo courtesy of Adam J. Varenhorst.

WCR females primarily lay eggs in cornfields, however there are instances where eggs are deposited in fields containing other crops. This is a resistance mechanism to crop rotation that is commonly referred to as the “soybean variant.” However, the rotation-resistant WCR can lay eggs in corn, soybean, oats, alfalfa, and winter wheat. Populations of rotation-resistant WCR are found in the eastern Corn Belt but remain rare in Iowa.

Adult SCR adults are bright yellow with 12 black spots on their hardened forewings. Sometimes this species is also known as twelve-spotted cucumber beetle. Their heads, legs, and antennae are always black. Adult SCR are the largest of the CRW species found in Iowa.

Southern corn rootworm. Photo courtesy of Adam J. Varenhorst.

SCR overwinter as adults in states to the south of Iowa, and then establish in Iowa each spring. Adults become active in the spring when temperatures exceed 70°F. After emerging, adult females lay their eggs near the base of corn stalks and larvae will feed on the corn roots. Both adult and larval SCR consume a wide variety of plants. Though SCR are commonly found in cornfields, they rarely cause economic injury to corn.

Click to visit the virtual dissection tool.

Open the virtual dissection to get started exploring the beetle. You will be able to use several tools to open, remove, and zoom in on many body parts. Follow along with our Dissection Activity.

Make sure to hover your cursor over all parts of the beetle so you don't miss anything. You can also "glue" the beetle back together to take a step back and explore a new area.

Honey Bees Make Honey … and Bread? | Deep Look

Bees pollinate an estimated 130 different varieties of fruit, flowers, nuts and vegetables in the United States alone. Farmers obviously depend on bees to pollinate crops, such as fruit and nuts, but in recent years thousands of bee colonies have disappeared. This could be a devastating issue for farmers. Can anything be done? Meet two Northern California researchers looking for ways to make sure we always have bees to pollinate our crops at

Lesson Summary

  • Arthropods are the largest phylum in the animal kingdom. Most arthropods are insects. The phylum also includes spiders, centipedes, and crustaceans. The arthropod body consists of three segments with a hard exoskeleton and jointed appendages.
  • Insects are arthropods in the class Hexapoda. They are the most numerous organisms in the world. Most are terrestrial, and many are aerial. Insects have six legs and a pair of antennae for sensing chemicals. They also have several eyes and specialized mouthparts for feeding. Insects are the only invertebrates than can fly. Flight is the main reason for their success. Insects may live in large colonies and have complex social behaviors. Insects spread disease and destroy crops. However, they are essential for pollinating flowering plants.

Lesson Review Questions


1. Identify distinguishing traits of most arthropods.

2. What is molting? Why does it occur?

3. Name three arthropod head appendages and state their functions.

4. Describe two structures that allow arthropods to breathe air.

5. List several traits that characterize insects.

6. State two important advantages of flight in insects.

7. Give examples of insect behavior.

Apply Concepts

8. Assume you see a “bug” crawling over the ground. It has two body segments and lacks antennae. Which arthropod subphylum does the “bug” belong to? Explain your answer.

Think Critically

9. Present facts and a logical argument to support the following statement: Insects dominate life on Earth.

10. Relate form to function in the mouthparts of insects.

11. Explain why distinctive life stages and metamorphosis are helpful.

Points to Consider

The invertebrates described so far in this chapter are protostomes. They differ from the other major grouping of animals, the deuterostomes, in how their embryos develop. The next lesson describes invertebrates that are deuterostomes. These invertebrates are more closely related to vertebrates such as humans. Some of these invertebrates are even placed in the chordate phylum.

Watch the video: Woodworm Identification: 11 Types of Wood Boring Insect (January 2023).