My, what big jaws you have my dear: Reddish-brown Stag Beetle, Lucanus capreolus

 

Do the impressive jaws of a male reddish-brown stage beetle strike fear in your heart?

 

Is the pretty female stag beetle a little less scary than its mate?

Well, we all know what happened to Little Red Riding Hood when she uttered similar words to a big, bad wolf posing as grandma. But fear not, the extremely large jaws of the male reddish-brown stag beetle are used to impress a potential mate, or to battle other males for mating rights to a comely and much smaller-jawed female stag beetle. Stag beetles are relatives of rhinoceros beetles we met in a previous episode. Male rhinoceros beetles, like our indigenous Hercules beetle, use prodigious horns on their head to battle competitors for access to females. Stag beetles dwell in damp forest woodlands (and apparently sometimes in my back yard), where females seek soggy tree holes like the one in my ancient maple or moist rotting stumps on the forest floor. Wet decaying wood serves as the favored location for female beetles to deposit eggs.

Watch as this pretty reddish-brown female stag beetle inspects the soggy interior of a large tree hole in my ancient red maple tree. She was rather nonplused about the camera and stopped a few times to glam for the cam. With some luck, perhaps she will find a mate and grace my rotting tree with a fine batch of lucanid grubs.

Larvae of stag beetles are quite similar in appearance to this rather large scarab grub.

Larvae, aka grubs, that hatch from these eggs take two years to develop as they feed on lignified tissues of their woody hosts. Stag beetle larvae house a rich microbial community in their gut. These symbionts release nutrients locked in tough woody tissues, making these goodies available to support the growth and development of stag beetle grubs. Along with several other species of wood digesting insects, lucanid larvae play a critical role in recycling organic matter in forests around the world. Adult lucanid beetles are reported to dine on fermenting exudates of plants. The important role of these forest recyclers is imperiled as forested lands disappear. A close relative of our reddish-brown stag beetle, the charismatic Lucanus cervus, has declined dramatically in some parts of Europe.

Stag beetles are noisy, somewhat clumsy fliers and create quite a buzz as they zoom though the forest or zero in on your porch light at nighttime. If you are a bug geek like me, you might just want to hold one of these reddish-brown stag beetles. Worried about those jaws? Don’t be. I have never had any issues holding males or females other than trying to release them when I was finished examining one. They have very sticky claws at the tips of their feet which enable them to climb trees and grip tightly to nosy humans. According to the Maryland Biodiversity Project, July is an excellent month to spot these fascinating creatures here in the DMV. Head for the forest to catch a glimpse of these giants of the beetle world.

Watch this amazing National Geographic video to see how stag beetles use their supersized jaws to defeat competitors and gain access to mates: https://www.youtube.com/watch?v=-VWFreC4onI

Acknowledgements

Bug of the week consulted these references for this episode: “Stag beetles” by Eric P. Benson,“Divergence in Gut Bacterial Community Structure between Male and Female Stag Beetles Odontolabis fallaciosa (Coleoptera, Lucanidae)” by  Xia Wan, Yu Jiang, Yuyan Cao, Binghua Sun and Xingjia Xiang, and “Insights into the ecology, genetics and distribution of Lucanus elaphus Fabricius (Coleoptera: Lucanidae), North America's giant stag beetle” by Michael Ulyshen, Louis  Zachos, John Stireman, Thomas Sheehan, and Ryan Garrick.

Paramesochra acutata | Catalogue of Organisms

Copepod taxonomy, it seems, is largely about counting setae. In his review of relationships within the interstitial harpacticoid family Paramesochridae, Huys (1987) recognised four species groups within the genus Paramesochra (which previously got a look-in at this site here). One of these groups, labelled the P. acutata-group, was characterised by reductions in numbers of setae, having lost the inner setae on the first segments of the endopods on the third and fourth legs.

Paramesochra taeana, a close relative of P. acutata, from Back & Lee (2010).

The group takes its name from the species Paramesochra acutata, described by Klie in 1935 from samples taken from coastal groundwater near the town of Schilksee on the northeastern coast of Germany, in the state of Schleswig-Holstein. Other notable features of P. acutata include the presence of four setae on the antennary exopod, well-developed narrow, triangular endopodal lobes on the modified fifth legs of the females, and conical caudal rami produced into spinose processes (Back & Lee 2013). I haven't been able to find whether P. acutata has been collected much beyond its initial locality but other members of its species group have been found around the world. One of these, P. hawaiensis, from (nach) Hawaii, is similar enough that it was until recently treated as a subspecies of P. acutata.

Appendages of female Paramesochra acutata, from Klie (1935).

So what, if anything, does all this mean? That, I'm afraid, is getting a bit beyond me. The fifth legs are used in spermatophore transfer and differences between species might presumably function in recognising suitable mates. Regarding the details of setation and ramus appearance, one wonders if there could be any relation to preferred micro-habitat. Are harpacticoids with fewer setae and more robust rami adapted for crawling among coarser sand grains? Honestly, I have no idea. Anyone care to find out?

REFERENCES

Back, J., & W. Lee. 2010. A new species of the genus Paramesochra (Copepoda: Harpacticoida) from Korean waters. Proceedings of the Biological Society of Washington 123 (1): 47–61.

Back, J., & W. Lee. 2013. Three new species of the genus Paramesochra T. Scott, 1892 (Copepoda: Harpacticoida: Paramesochridae) from Yellow Sea, Korea with a redescription of Paramesochra similis Kunz, 1936. Journal of Natural History 47 (5–12): 769–803.

Huys, R. 1987. Paramesochra T. Scott, 1892 (Copepoda, Harpacticoida): a revised key, including a new species from the SW Dutch coast and some remarks on the phylogeny of the Paramesochridae. Hydrobiologia 144: 193–210.

Klie, W. 1935. Die Harpacticoiden des Küstengrundwassers bei Schilksee (Kieler Förde). Schriften des Naturwissenschaftlichen Vereins für Schleswig-Holstein 20 (2): 409–421.

Waxy wanderers flocking flowers: Planthoppers - Flatidae and Acanaloniidae

 

See how many flatid nymphs you can find hiding amongst their white wax on this branch.

 

Strange looking acanaloniid nymphs remind me of the fierce Blurrg of Mandalorian lore.

Last week we visited dastardly Japanese beetles shredding leaves of zinnias. On a follow-up visit to the flowers, I was surprised to see several stems festooned with fluffy white wax reminiscent of the mysterious substance from an aerosol can used to flock Christmas trees. Beneath the white flocking were nymphs of small sucking insects known as flatid planthoppers. Nearby a bizarre acanaloniid nymph baring a strange resemblance to a miniature Blurrg wandered about a flower stem trailing an array of waxy filaments from its rear end. These sucking insects are close relatives of other well-known wax producing sap-suckers such as boogie-woogie aphids and woolly alder aphids we met in previous episodes.  

Two kinds of sap-sucking planthoppers are flocking my plants with white wax. The strange brown planthopper on top is an acanaloniid and the greenish one hiding below in white wax is a flatid nymph. When the lens gets too close the flatid hops to a leaf below and takes a stroll. The acanaloniid nymph meanders up the stem to escape a nosy bug geek.

It’s not unusual to find ranks of waxy adult citrus flatids lining stems and branches of many kinds of plants in the summertime.

Flatid and acanaloniid planthoppers feed by inserting a small beak into the vascular system of a plant and withdrawing the nutrient-laden sap into their bodies using a small hydraulic pump located in their head. As a byproduct of this feeding, they excrete a sticky substance called honeydew that attracts other insects such as wasps and ants and serves as a substrate for the growth of a fungus called sooty mold. They also secrete a pure white wax from glands lining their abdomen. The function of this wax may be to protect them from tiny parasitic wasps determined to deliver a sting and deposit eggs into their bodies. Or the wax may confuse hungry predators such as lacewing larvae and ladybugs intent on a feast, that are bamboozled when they get a mouthful of white wax instead of planthopper flesh. Whatever the purpose of the wax, it is produced in prodigious amounts by flatids and acanaloniids.  

Leaf-mimicking cone-headed planthoppers are masters of disguise.

Planthopper nymphs hatched this spring from eggs inserted in the stem of a plant last year by the adult planthopper. Now the nymphs are beginning to molt and will soon become adult hoppers that will remain on our plants for the duration of summer and into autumn. Adult flatid planthoppers do not produce vast amounts of wax as they did in their youth, but their bodies are coated with a lovely bloom of grey, green, or bluish wax. Acanaloniid adults are tiny leaf mimics with myriad wing veins resembling the veins of leaves. They blend in remarkably well with the plants on which they feed. Flatids and acanaloniids are common on roses, dogwoods, locusts, privets, hollies, maples, and many herbaceous perennials and annuals. Flatid planthoppers have been reported to cause the terminals of small plants to droop due to their feeding. Females inserting large numbers of eggs into the stems of plants may also cause small branches or seedlings to wither. However, in general, flatid planthoppers cause very little injury to plants in our gardens and the use of insecticides to eliminate them is unwarranted. With the arrival of hot, humid, dog-days of summer, their holiday flocking seems to be a welcome reminder of cooler times to come.  

Acknowledgements       

The fact-filled articles “Histories of Anormenis septentrionalis, Metcalfa pruinosa, and Ormenoides venusta with Descriptions of Immature Stages” by S. W. Wilson and J. E. McPherson, and “Planthoppers” by Steven Frank, James Baker, and Stephen Bambara were used to prepare this episode.

Melanterius Weevils | Catalogue of Organisms

Here in the Antipodes, we have a long history of environmental upheaval from exotic taxa unwisely released. As a result, one can't help but feel an odd twinge of perverse patriotism when hearing of the inverse, some native of the Antipodes causing grief elsewhere. In South Africa, Australian acacias have become something of an issue, inciting a search for potential control agents. Among the candidates selected are weevils of the genus Melanterius.

Melanterius servulus, copyright Sally Adam.

Melanterius is a diverse genus of small black or brown weevils (ranging from about three to seven millimetres in length) that feed as both adults and larvae on the developing seeds of acacias. About eighty species have been recognised in the genus to date and possibly many more remain to be described. In general, Melanterius weevils are heavily punctate, usually without prominent hairs but with a covering of scales. The rostrum is reasonably long, reaching more or less back to the mesosternum at rest but not sitting in a distinct ventral groove, and may be variably curved (going by figures in Zimmerman 1992).

Melanterius semiporcatus, copyright Victor W. Fazio III.

As with other weevils, the prominent rostrum is used by females to chew into an appropriate spot on the host plant, in this case chewing holes into the developing acacia seed pods, into which eggs are laid. Melanterius species go through one generation per year. Larvae burrow into and feed on the developing seeds before emerging and dropping to the ground to pupate in the soil. Mature adults emerge well before the host acacias begin to set seeds, usually having to wait about six months (Auld 1989). They usually spend the intervening period largely inactive, sheltering in concealed places close to the host plant and occasionally emerging to briefly feed on developing buds.

Under peak conditions, Melanterius infestations may cause a complete failure of seed production. No wonder, then, that they have been considered a worthwhile instrument of biological control.

REFERENCES

Auld, T. D. 1989. Larval survival in the soil and adult emergence in Melanterius Erichson and Plaesiorhinus Blackburn (Coleoptera: Curculionidae) following seed feeding on Acacia and Bossiaea (Fabaceae). Journal of the Australian Entomological Society 28: 235–238.

Zimmerman, E. C. 1992. Australian Weevils (Coleoptera: Curculionoidea) vol. 6. Colour plates 305–632. CSIRO Australia.

Feasting and frolicking spell trouble when Japanese beetles, Popillia japonica, arrive

 

Volatile chemicals released by damaged leaves and sex pheromones released by female Japanese beetles result in a rambunctious feeding frenzy and love fest on infested plants.

 

Lindens, a favorite host for Japanese beetles, can be severely damaged when beetles are numerous.

While enjoying the flower garden last week, my reverie was disturbed by the appearance of some very raggedy leaves on the zinnias. A closer inspection quickly revealed the culprit behind this assault, dastardly Japanese beetles. Historically, late June and early July are the months of misery when Japanese beetles abound, and these mischief makers have arrived right on schedule. The first detection of Japanese beetle in the United States occurred in 1916 in a plant nursery in New Jersey. They likely arrived with plant material imported from Asia, as grubs in the soil or as adult stowaways in the foliage of plants. Japanese beetles are extreme foodies with more than 400 kinds of trees, shrubs, vines, and herbaceous plants on the menu. Among their favorites are sassafras, lindens, maples, apples, cherries, grapes, roses, and apparently, my zinnias. In a series of studies, entomologist Dan Potter and his colleagues in Kentucky found that roses with large, light colored blossoms, particularly yellow or white, were more attractive to Japanese beetles than varieties with smaller, darker blossoms of red or orange. In the tree realm, researchers noted that lindens with densely hairy leaves were less preferred than scantily haired varieties. Maples with purple or deep red leaves were preferred over those with green leaves.

Traps collect large numbers of Japanese beetle but plants near traps may be damaged when beetles assemble nearby. Place traps away from plants you want to protect.

You may have noticed that Japanese beetles often attack one plant severely, leaving a lucky neighbor relatively unscathed. When these invaders initiate an attack, specific odors are released by the damaged plant. These send a signal to other beetles, something like "good food, eat here.” This foliar attractant is compounded when female beetles release a chemical message called a sex pheromone. The sex pheromone says to the guy beetles, “how’d you like to spend a little time with me?” Not surprisingly, a rambunctious love fest and feeding frenzy erupt, and in the process, your plant takes a beating. Clever chemists have been able to synthesize both a floral attractant released by plants and the Japanese beetle sex pheromone and place them in a lure. Attach the lure to a few plastic fins for beetles to bump into, and a funnel to direct them into a plastic bag and, voilà, you have a Japanese beetle trap.

Japanese beetle traps capture beetles by the thousands, but traps may not be all that effective in protecting your plants. Plants near the traps may actually sustain more damage as beetles lured to the vicinity mill around awaiting their turn to hit the fins and be captured. It is best to place these traps far away from valued plants you want to protect. Japanese beetles lay their eggs in soil, so if adult beetles are a chronic problem in your garden or landscape, the best way to get relief may be to reduce the numbers breeding in your lawn, especially if you have irrigated turf. One promising “organic” approach is to apply insect pathogenic nematodes, tiny roundworms that attack and kill beetle grubs. Nematodes enter the grub and release a lethal bacterium. There are many different species and strains of nematodes. Dave Shetlar of the Ohio State University suggests that products containing strains of Steinernema carpocapsae nematodes are a bit less effective against beetle grubs than species in the clan named Heterorhabditis. If you go the nematode route, you must wait to make an application until late July or August when grubs are in the soil. A second formulated microbial insecticide that shows promise is Bacillus thuringiensis galleriae, which has been demonstrated to reduce feeding by adult beetles when applied to foliage.

Larvae of many species of scarabs, including Japanese beetles, are called white grubs. They damage roots of plants.

There are several potent soil insecticides that can be applied in late July through August that are very effective in killing tiny grubs as they hatch from eggs and feed near the soil surface. If you opt for the synthetic chemical route, choose wisely. We now know that at least one class of insecticides called neonicotinoids applied to turf grass can be taken up by clover growing in turf. Bumble bees foraging in this clover may be harmed. However, a newer class of insecticides called the anthranilic diamides present far fewer risks to our hard-working pollinators. When using any insecticide, always read the label and follow the directions carefully and pay particular attention to warnings pertaining to beneficial insects like bees.

Mammals including skunks and raccoons damage lawns as they hunt for white grubs of Japanese beetles and other scarabs in soil.

Many insecticides are available to control adult Japanese beetles on plants, but multiple applications may be necessary if you cannot tolerate damage by these critters. As with turf applications, be cognizant of beneficial insects foraging on plants. Read and follow label precautions. Another nifty way to help reduce damage is to simply knock the beetles from your plants into a bucket of soapy water. If you do this early in the season of evil when beetles first arrive, you may reduce the chemical cues that incite a feeding frenzy. Beetle removal may be most successful in early morning or late evening when beetles are less active. There is a strange kind of justice in drowning this pest in soapy water or capturing them in traps. Save the bodies of the little rascals captured in your bucket or trap. The earthly remains of so many beetles make a wonderful addition to a compost pile that can later be used to nourish your garden.

The 4th of July heralds the arrival of Japanese beetles and flowering plants are now under attack. Volatiles from leaves shredded by beetles and sex pheromones released by females bring more beetles and damage to infested plants. Knocking beetles into a jar of soapy water when just a few are on a plant may derail the feasting and love fest. Japanese beetles are poor swimmers and after they expire, I add their tiny bodies to my compost. From soil they came, to soil they return.

Acknowledgements

Excellent references including “Assessing Insecticide Hazard to Bumble Bees Foraging on Flowering Weeds in Treated Lawns” by Jonathan L. Larson, Carl T. Redmond, and Daniel A. Potter and “Strengths and limitations of Bacillus thuringiensis galleriae for managing Japanese beetle (Popillia japonica) adults and grubs with caveats for cross-order activity to monarch butterfly (Danaus plexippus) larvae” by Carl T. Redmond, Lindsey Wallis, Matthew Geis, R. Chris Williamson, and Daniel A. Potter were used to prepare this episode. Our thanks to Dr. Shrewsbury for assistance in sending beetles to a better place.

Kirkby's Small Ostracods (or Small Kirkby's Ostracods) | Catalogue of Organisms

I do not envy those who find themselves working with ostracods. These minute crustaceans, typically less than a millimetre in length, seem altogether too fiddly to handle. Nevertheless, the long history of ostracods, together with their diversity and the high fossilisation potential of their calcified carapace valves, have made them a common focus for studying biostratigraphy and historical environments. The classification of modern ostracods is commonly informed by features of the legs and other appendages but such characters are not commonly preserved in fossil representatives. As a result, there are many groups of ostracods known from the Palaeozoic whose relationships remain uncertain.

Left valve of Kirkbyella delicata, from Hoare & Merrill (2004).

One such group is classified by Liebau (2005) as the superfamily Kirkbyelloidea. Members of this group are small ostracods with reticulate valves. The dorsal and ventral margins of the valves tend to be more or less straight. They are commonly impressed with a single dorsal sulcus, extending downwards from the dorsal margin about halfway along the valve's length. Below this sulcus is a protruding horizontal lobe ending in members of the family Kirkbyellidae in a small spine. Evidence of sexual dimorphism, a not-uncommon feature of Palaeozoic ostracods, is not known from kirkbyelloids.

Definite kirkbyelloids are known from the Devonian to the Permian. If the earlier family Ordovizonidae is included, their record extends all the way back to the Ordovician. As noted above, it is unclear where kirkbyelloids sit in the ostracod family tree. Becker (1994) suggested a relationship via Ordovizona to the Ordovician Monotiopleuridae which resemble kirkbyelloids in the outline of the carapace valves and features of the adductor muscle scars. Though long-lived, kirkbyelloids don't seem to have ever been massively diverse, and they can probably be counted among the many lineages of organisms that never made it past the end of the Palaeozoic.

REFERENCES

Becker, G. 1994. A remarkable Ordovician ostracod fauna from Orphan Knoll, Labrador Sea. Scripta Geologica 107: 1–25.

Hoare, R. D., & G. K. Merrill. 2004. A Pennsylvanian (Morrowan) ostracode fauna from Texas. Journal of Paleontology 78 (1): 185–204.

Liebau, A. 2005. A revised classification of the higher taxa of the Ostracoda (Crustacea). Hydrobiologia 538: 115–137.

Conformed Flycatchers | Catalogue of Organisms

A quote I have often had cause to refer to—I believe it originally came from Toby White of Palaeos.com—is that "organisms are under no obligation to speciate with regard to the convenience of taxonomists". For birdwatchers in North America, perhaps no group more embodies this principle than the flycatchers of the genus Empidonax. These small members of the hyperdiverse New World family Tyrannidae comprise fifteen recognised species that have become notorious for the difficulty in telling them apart.

Immature alder flycatcher Empidonax alnorum, copyright Cephas.

The species of Empidonax are uniformly olive brown above, lighter below, with pale rings around the eyes and bands on the wings. They are inhabitants of woodlands (more on that in a moment) and watch for flying insects from a perch, making short flights to capture prey. Though individual species are generally similar in their feeding habits, they are often specifically distinct in their preferred habitats. A molecular (mtDNA) analysis of Empidonax species by Johnson & Cicero (2002) identified four likely clades within the genus with members of a clade each differing in their specific breeding range. Species found in the US and Canada often migrate long distances and closely related species may be found close together outside their breeding ranges (references to ranges below refer to breeding ranges). Species found in Mexico and Central America are more likely to migrate only short distances or be resident year-round.

Acadian flycatcher Empidonax virescens, copyright Aitor.

The Acadian flycatcher E. virescens seems to be relatively isolated from other members of the genus. This species is found in shady forests near water in the eastern US and Canada. Its nest is a cup made from plant fibres suspended in a horizontal branch fork, and it lays lightly speckled eggs.

The yellow-bellied flycatcher E. flaviventris, yellowish flycatcher E. flavescens, Cordilleran flycatcher E. occidentalis and Pacific slope flycatcher E. difficilis form a clade of species that tend to have more yellowish underparts than other members of the genus. Their nests are mossy cups constructed on a protected ledge or crevice. Members of this clade tend to be found in relatively damp forest areas, such as boggy areas of boreal forests in the case of E. flaviventris, or shady canyons in the case of E. occidentalis or E. difficilis. A notable exception is the Channel Islands population of E. difficilis which is found in more open woodlands than its mainland counterparts. Empidonax occidentalis and E. difficilis are found in the western United States with E. difficilis occupying coastal regions and E. occidentalis found further inland. Until fairly recently, the two were confused as a single species; they are almost indistinguishable morphologically but can be separated by their calls.

Least flycatcher Empidonax minimus, copyright Mdf.

The white-throated flycatcher E. albigularis, alder flycatcher E. alnorum and willow flycatcher E. traillii form a clade of species nesting in damp thickets. Again, it was only fairly recently that the more northerly E. alnorum was distinguished from the more southerly E. traillii.

Finally, the remaining species form a clade whose members lay eggs without speckled markings. They are often relatively dark compared to other Empidonax; the black-capped flycatcher E. atriceps of Costa Rica and Panama stands out for the sooty-black coloration of the head. They often inhabit relatively open forest, often at higher altitudes.

Johnson & Cicero (2002) suggested that the largely allopatric (non-overlapping) breeding ranges of species within clades of Empidonax reflected speciation as a result of isolation in glacial refuges during the ice ages. As the ice retreated, the now-distinct species expanded their ranges but excluded each other where they met. Differences in mating calls between related species dissuaded interbreeding. Physical appearance, meanwhile, remained frustratingly monotonous.

REFERENCE

Johnson, N. K., & C. Cicero. 2002. The role of ecologic diversification in sibling speciation of Empidonax flycatchers (Tyrannidae): multigene evidence from mtDNA. Molecular Ecology 11: 2065–2081.

Small, strange bugs with very long legs: Stilt bugs, Berytidae

 

A stilt bug ponders its next move at the edge of a zinnia leaf. Credit: Paula Shrewsbury, UMD

 

Amazingly long legs transport the stilt bug across a hairy leaf surface. Note its super long antennae.

Last week we witnessed the life and death struggle of thrips as they dodged fierce minute pirate bugs in cone flowers. Just down the flower bed from the cone flowers, a patch of zinnias sported a wide array of six-legged visitors, Japanese beetles, planthoppers, and a gangly member of the true bug clan, a heteropteran known as a stilt bug. In a previous episode we met fast moving stilt-legged flies snacking on tidbits of cheese and pretending they were ants on leaves in a rainforest. This week’s long-legged wonders seem to be mostly stuck in slow motion as they amble across vegetation in search of a meal or a mate. One common species here in the DMV, Jalysus wickhami, dines on more than four dozen species of plants in some seventeen plant families, most of which have been described as "glandular-hairy." Due to their behavior of jabbing their pointy beaks into stems, flowers, and fruits to suck out nutritious cell contents, they can be severe pests of tomatoes grown outdoors and in hot-houses. When fed upon by stilt bugs, tomato flowers and stems can turn black and die. Feeding punctures can distort and discolor tomatoes making them unsalable.

This mating pair of stilt bugs certainly is not camera shy. The bug on the right is the female; the male is on the left. Watch as the female multitasks, grooming her legs while engaged with her mate. She collects a drop of fluid from her beak with her middle and hind legs and appears to apply it the surface of her hind leg. How curious is that?

A second species of stilt bug common in our region, Jalysus spinosus, is more of a specialist, feeding and breeding primarily on grasses in the genus Panicum. Ah, but like many of us, while the vegan diet may be a healthy one, every now and then a little meat doesn’t hurt, and stilt bugs can be important predators of insect pests, consuming eggs of hornworms, aphids and other small soft-bodied pests in a variety of crops. Nymphs and adults of J. wickhami have been raised and released in tobacco fields to augment activities of predatory insects already present in the crop. In addition to the two Jalysus species mentioned above, the Maryland Diversity Project lists Berytinus minor and Neoneides muticus as residents of our region. So, next time you visit your gardens and flower beds, keep a sharp eye out for these interesting tiny omnivores with unusually long legs.

Acknowledgements

Bug of the Week thanks Dr. Shrewsbury for her image of a stilt bug. Details of the life history and ecology of stilt bugs were extracted from these interesting references: “Jalysus spinosus and J. wickhami: Taxonomic Clarification, Review of Host Plants and Distribution, and Keys to Adults and 5th Instars” by A. G. Wheeler and Thomas Henry, “Spined Stilt Bug” by Michael Skvarla, and “Spined Stilt Bug in Tobacco” by Peter Nelson and Hannah Burrack.

This bevy of stilt bug nymphs are “beaks in” on this tender flower stem.

The Teleost Fuse | Catalogue of Organisms

A while back, I discussed the group of fish known as the Holostei, the gars and bowfin. The Holostei constitute one branch of the clade Neopterygii which includes the majority of living ray-finned fishes. However, their success in the modern environment pales in comparison to that of their sister group, the Teleostei.

Siemensichthys macrocephalus, an early teleost of uncertain affinities, copyright Ghedoghedo.

Teleosts are such a major component of ray-finned fishes that it is simpler to list those members of the modern fauna that do not belong to this clade: the aforementioned gars and bowfin, sturgeons and paddlefish, and the bichirs of Africa. Everything else belongs to the great teleost radiation, representing about 96% of all modern fishes. The earliest fishes generally recognised as teleosts come from marine deposits of the Late Triassic in the form of the Pholidophoridae of Europe. The earliest known members of the crown group are from the Late Jurassic (Nelson et al. 2016). Teleosts have been recognised as an apomorphy-defined clade; the crown clade has been dubbed the Teleocephala. Among the features that have been used to define the Teleostei are the presence of a mobile premaxilla. In my previous post, I explained how the mobile maxilla of neopterygians including bowfins improved feeding by creating suction when the mouth was opened. Having both the maxilla and premaxilla mobile enhances this process further. In some of the most advanced teleosts, such as dories and ponyfish, the connection between the jaws and the cranium is entirely comprised of soft, flexible tissue, allowing the jaw apparatus as a whole to be catapulted towards unwary prey. Other features that have been highlighted include a strongly ossified caudal skeleton with long uroneural spines derived from the neural arches of the vertebrae, and the lower lobe of the caudal fin supported by two plate-like hypural bones articulating with a single vertebral centrum (Bond 1996).

Leptolepis coryphaenoides, one of the earliest teleosts with cycloid scales, copyright Daderot.

Of course, not all these features necessarily appeared in lock with each other. A phylogenetic analysis of basal teleosts by Arratia (2013) identified the aforementioned features of the caudal skeleton as absent in some of the basalmost teleosts. The condition of the premaxilla is ambiguous in Prohalecites, the earliest stem-group teleost from the Middle-Late Triassic boundary. It appears to be absent in the Aspidorhynchiformes and Pachycormiformes, Mesozoic orders that are currently regarded as on the teleost stem but not part of the Teleostei. However, as was found with the mobile maxilla in gars, one can't help wondering whether this character has been affected by the uniquely derived upper jaw morphologies in these orders. Other features identified by Arratia (2013) as supporting the Teleostei clade include the presence of two supramaxillary bones, a suborbital bone between the posterior margin of the posterodorsal infraorbitals and the anterior margin of the opercular apparatus (subsequently lost in the teleost crown group), and accessory suborbital bones ventrolateral to the postorbital region of the skull roof.

The earliest teleosts in the Pholidophoridae and other basal lineages retained the heavy ganoid scales of thick bone that may still be seen in modern Teleostei. Lighter, thinner cycloid scales first appear with the Early Jurassic Leptolepis coryphaenoides (Arratia 2013) and are the basal scale type for the teleost crown group (in some derived subgroups, the scales would become further modified or even lost). The greater mobility permitted by these lighter scales may have been another significant factor in the teleost explosion. By the Cretaceous period, stem-teleosts had radiated into a variety of specialised forms such as the gigantic predatory Ichthyodectiformes (of which Xiphactinus grew up to four metres in length) and the deep-finned Araripichthys. The three major subgroups of the crown Teleostei—the Elopomorpha, Osteoglossomorpha and Clupeocephala—had diverged from each other by the end of the Jurassic. The stem-teleosts would disappear with the end of the Mesozoic; the crown teleosts would dominates the world's waters from that time on.

REFERENCES

Arratia, G. 2013. Morphology, taxonomy, and phylogeny of Triassic pholidophorid fishes (Actinopterygii, Teleostei). Journal of Vertebrate Paleontology 33 (6 Suppl.): 1–138.

Nelson, J. S., T. C. Grande & M. V. H. Wilson. 2016. Fishes of the World 5^th ed. Wiley.