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What is this tiny insect?

What is this tiny insect?


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I took this picture in central Mexico. The insect is about 1mm in size. I found lots of these dead on a desk, so many that at first I thought they were just dust. I assume they came from a plant that was above the desk.

Does anyone know what it might be?

EDIT:

I found the culprit walking nearby! I guess that verifies what others pointed out: It seems to be the molted cuticle of an aphid. (sorry for the poor quality of the pictures)

Thanks a lot!


Since you suggest they come from a plant, they might be dried aphids. These insects usually feed on plant sap, and I already noticed some tiny, white, and apparently dead ones on plants.

As suggested in the comments, these are actually not dead aphids, but only the cuticle they leave after moulting, or ecdysis.


What is the smallest insect on Earth? -Laurenz, 8, Molino, Philippines

When I saw your question, I set out to explore with my bug net and a magnifying glass. I was searching all around for tiny insects when I ran into my friend Laura Lavine, a Washington State University scientist who studies bugs.

She said there are nearly a million different kinds of insects on Earth. The smallest of all the known ones are called fairyflies.

Like all insects, fairyflies have six legs. And like most insects, they also have wings. Some swim under water and use their wings as paddles. Their wings are also a bit hairy. It also turns out the fairyfly isn’t truly a fly. It’s a kind of wasp.

“They are almost impossible to spot with the naked eye,” Lavine said.

In fact, fairy flies are nearly 400 times smaller than the typical ant. And they are about two or three times the width of a human hair.

I imagine finding a fairyfly would be like finding a needle in a haystack. You’d have to keep a sharp eye out.

I started to wonder how exactly entomologists could spot such tiny fairyflies or other kinds of small insects in the wild. For example, a couple years ago scientists discovered a new kind of fairyfly in Costa Rica. It was named Tinkerbella nana after the fairy from Peter Pan.

Lavine explained that scientists often use nets or traps to catch the insects. Sometimes they have to sift through dirt and litter, or decaying leaf matter, a teaspoon at a time to see what they can find.

Scientists can also use what they know about the insect’s behavior and habitat to help track them down. Fairyflies, despite their cute name, are killer insects. They lay eggs inside a host insect’s egg. When the fairyfly’s egg hatches, it eats the host egg. If we keep our eyes out for their host bugs and their eggs, we might also find the fairyfly.

Fairyflies are important for the environment, Lavine added. Farmers and scientists can use fairyflies to help get rid of bigger insects that damage grape vines, blackberries and sugar cane. These tiny creatures help us do a big job.

The insect world is filled with interesting critters. Thinking about the smallest insect also made me wonder about the biggest one on our planet. The biggest bug is a giant walking stick. It’s almost 2 feet long. But who knows? There might be even bigger insects or even smaller insects we haven’t discovered yet crawling around on our planet.

Thanks for your question, Laurenz. It reminds me that even the small things can inspire us to wonder big.


Small Alpine Insects are Big Messengers of Climate Change

West Glacier, Mont. – Two rare alpine insects – native to the northern Rocky Mountains and dependent on cold waters of glacier and snowmelt-fed alpine streams – are imperiled due to climate warming induced glacier and snow loss according to a study by the U.S. Geological Survey and its partners.

The 20-year study provides the first comprehensive evaluation of the current status, distribution and habitat requirements for each species and was used to inform the status review for consideration of protection under the U.S. Endangered Species Act due to climate-change-induced habitat loss.

Researchers found the meltwater stonefly and western glacier stonefly have a narrow distribution and are restricted to short sections of cold, alpine streams often below glaciers predicted to disappear over the next two decades.

“Alpine aquatic insects living in glacier and snow-fed streams are adapted to very cold water temperatures and are therefore especially vulnerable to warming and snow and ice loss,” said Joe Giersch, USGS entomologist and lead author of the study. “Although this research is focused on two insect species, our findings apply to entire communities of alpine stream organisms, the survival of which depends on the presence of permanent ice and snow feeding the streams in which they live.”

Over the duration of the study, researchers sampled 272 alpine streams in Glacier National Park, where the insects are native, and other areas outside the known distribution throughout the Rocky Mountains of Montana and Wyoming.

The meltwater stonefly was found in 113 streams within Glacier National Park and surrounding areas. The likelihood of finding the insect increased with cold stream temperatures and proximity to glaciers and permanent snowfields, and declined with increasing distance from stream source.

The western glacier stonefly was only found in 10 streams, six in Glacier National Park and four in mountain ranges almost 400 miles southwest.

Both stoneflies were present in groundwater-fed alpine springs, which may provide refuge when alpine stream conditions decline.

Alpine streams environments in the northern Rocky Mountains are especially vulnerable to climate change due to rapid warming resulting in loss of glaciers and snowpack. Glacier National Park is iconic of the combined impacts of climate change and snow and ice loss – over 80 percent of the park’s glaciers have been lost since the mid-19th century.

This study brings to light how an obscure species could be true biological sentinels of climate change because their survival is dependent on a habitat that is rapidly becoming fragmented and degraded, making it very difficult for them to adapt. The results provide a rare example linking climate-change-induced habitat loss with conservation implications for rare, understudied species.

"By clearly linking glacier decline to the loss of alpine species, we can more confidently state threats to conservation of alpine species in Glacier National Park and also extend the discussion to similar situations worldwide” said Scott Hotaling, doctoral candidate at the University of Kentucky and co-author of the study.

“These stoneflies are retreating up the mountain to seek out cold water, but there is nowhere to go because they’re literally at the top of the continent” says Clint Muhlfeld, USGS ecologist and study co-author. “Global warming-induced glacier and snow loss not only threatens some species with extinction, but also has far-reaching effects on regional and global biodiversity.”

The study is a collaborative effort between the USGS, University of Kentucky, University of Montana and Glacier National Park.

More information about climate impacts to rare alpine insects can found on the USGS Northern Rocky Mountain Science Center website.


MATERIALS AND METHODS

As both viscosity and the interactions of the flexible wing with the air are important at this scale, a direct numerical simulation of the fully coupled fluid–structure interaction problem is appropriate. In this paper, the immersed boundary method(Peskin, 2002 Zhu and Peskin, 2002) is used to simulate two flexible wings immersed in a viscous, incompressible fluid. The immersed boundary method has been used successfully to model a variety of problems in biological fluid dynamics. Such problems usually involve the interactions between incompressible viscous fluids and deformable elastic boundaries. Some examples of biological problems that have been studied with the immersed boundary method include aquatic animal locomotion(Fauci, 1990 Fauci and Fogelson, 1993 Fauci and Peskin, 1988),cardiac blood flow (Kovacs et al., 2001 McQueen and Peskin, 1997 McQueen and Peskin, 2000 McQueen and Peskin, 2001) and ciliary driven flows (Grunbaum et al.,1998).

In this paper, the Re is set to 10, corresponding to the case of some of the smallest flying insects such as Thripidae frankliniella(Sunada et al., 2002). The non-dimensional bending stiffness is varied from about 0.25 to 4. The mass of a thrips is about 6.0 × 10 –8 kg(Tanaka, 1995) and its wing length is about 0.75 mm. Assuming that a wing weighs about 3.0 ×10 –9 kg (this is 1/20 of the total mass, and the feathery wings probably weigh much less) and the wing length is 0.75 mm,ρ s would be roughly equal to 4.0 × 10 –6 kg m –1 . Given a wing chord of about 0.25 mm and an air density of 1.2 kg m –3 , m′ would equal 0.016. Because this value is much less than one, the effects of inertia on the wing are ignored. The wing is modeled as a massless boundary that resists bending and stretching.

The basic idea behind the immersed boundary numerical method is as follows.

At each time step, calculate the forces that the boundaries impose on the fluid. These forces are determined by the deformation of the elastic boundaries. Additional external forces used to drive the motion of the boundary may also be applied to the fluid.

Spread the force from the Lagrangian grid describing the position of the boundaries to the Cartesian grid used to solve the Navier–Stokes equations.

Solve the Navier–Stokes equations for one time step.

Use the new velocity field to update the position of the boundary. The boundary is moved at the local fluid velocity, enforcing the no-slip condition.

Further details of the immersed boundary method and its discretization may be found in the Appendix.

Numerical simulations

The two-dimensional numerical simulations of flight in this paper were constructed to be similar to the physical experiments of Dickinson and Götz (Dickinson and Götz,1993) and previous two-dimensional numerical simulations of clap and fling (Miller and Peskin,2005). Dickinson and Götz used an aluminium wing with a chord of 5 cm immersed in a sucrose solution with a kinematic viscosity of 0.0000235 m 2 s –1 (about 20 times that of water) moving with a characteristic velocity in the range of 0.04–0.12 m s –1 . The dimensions of the sucrose tank used in the physical experiment were 1 m in length by 0.4 m in width. The same parameters as listed for this physical experiment were used in all of the following numerical experiments with two exceptions: (1) the size of the computational tank was increased to reduce wall effects at lower Re and (2) the translational velocity was changed to simulate Re=10. In the following simulations, we use a computational tank that is 1 m × 1 m in size. For numerical convenience, we place this tank within a slightly larger periodic domain, of size (1 m+30 h) × (1 m+30 h),where hxy is the mesh width of our fluid grid. The edges of the computational tank are made of immersed boundary points that are linked by stiff springs to stationary target points. The region within the four walls is called the `computational tank'. The Navier–Stokes equations were solved on a 1230×1230 Cartesian grid,and each wing was discretized on a Lagrangian array of 120 points. Miller and Peskin (Miller and Peskin,2005) presented a convergence analysis that showed that this mesh size is within the range of convergence for the two-wing problem at Re below 100.

Unless otherwise stated, the motion of the flexible wings was prescribed by attaching target points to the top 1/5 of the boundary along the leading edge of the wing with springs (Fig. 2). The target points moved with the prescribed motion and applied a force to the boundary proportional to the distance between the target and corresponding boundary points. The bottom 4/5 of the wing (trailing edge) was free to bend. This has the effect of modeling a flexible wing with a rigid leading edge. In the `nearly-rigid' case, springs were attached to target points along the entire length of the wing, which prevented any significant deformation. The dimensional stiffness coefficient of the springs that attach the boundary points to the target points was ktarg=1.44×10 5 kg s –2 ,and the stiffness coefficient for the tension or compression of the wing was also set to kstr=1.44×10 5 kg s –2 . These values were chosen to prevent any significant stretching or deformation of the wings in the rigid case.

The dimensional flexural or bending stiffness of the wing was varied from kbeam=0.125 κ to 2 κ Nm 2 , whereκ=5.5459×10 –6 Nm 2 . As stated above,this range of bending stiffnesses corresponds to dimensionless values ranging from 0.25 to 4. We chose this value of κ to represent the case where deformations during translation are small but bending does occur when the wings are close. This range of values was found to produce deformations that are qualitatively similar to those observed in flight videos.

Design of the flexible wing. The fluid domain is represented as a Cartesian grid, and the boundary (wing) points are represented as red circles. These points interact with the fluid and move at the local fluid velocity. The green springs represent the bending and stretching stiffness of the boundary. The desired motion of the wing is prescribed by the target points along the top 1/5 of the wing, shown above as blue circles. These points do not interact with the fluid and they move according to the desired motion of the wing. They also apply a force to the actual boundary via the target springs(yellow springs). Because the target springs are only connected to the leading edge of the wing, this has the effect of making a wing with a stiff leading edge and flexible trailing edge.

Design of the flexible wing. The fluid domain is represented as a Cartesian grid, and the boundary (wing) points are represented as red circles. These points interact with the fluid and move at the local fluid velocity. The green springs represent the bending and stretching stiffness of the boundary. The desired motion of the wing is prescribed by the target points along the top 1/5 of the wing, shown above as blue circles. These points do not interact with the fluid and they move according to the desired motion of the wing. They also apply a force to the actual boundary via the target springs(yellow springs). Because the target springs are only connected to the leading edge of the wing, this has the effect of making a wing with a stiff leading edge and flexible trailing edge.

For the smallest insects that probably transport themselves on gusts of wind, we use the ratio of the average lift force produced during the stroke to the average drag force as a simple measure of flight efficiency. Lift (the vertical component of the force) is generated to help keep the insects afloat while drag (the horizontal component of the force) is of less importance since any thrust produced would likely be swamped by wind gusts. This metric is particularly appropriate when the wings are close to each other because the horizontal component of the force acting on each wing cancels. It is worthwhile to note that a number of other measures of efficiency are used in the literature, many of which are likely to be more appropriate for medium to large insects (e.g. Wang,2004).

Kinematics of the clap and fling strokes

Quantitative descriptions of the wing beat kinematics of the smallest flying insects are currently unavailable. This is partially due to the fact that these insects are extremely difficult to film. Many of these insects may be as small as 0.25 mm in length and flap their wings at frequencies of 200 Hz or greater (Dudley, 2000). The authors know of several unpublished high-speed videos of Thysanoptera,Muscidifurax raptor and Nasonia vitripennis shot from a single camera, but quantitative reconstruction of the wingbeat kinematics is not possible from one camera angle. The simplified flight kinematics of clap and fling used in this paper are similar to those used by Lighthill(Lighthill, 1973), Bennett(Bennett, 1977), Maxworthy(Maxworthy, 1979), Spedding and Maxworthy (Spedding and Maxworthy,1986), Sun and Xin (Sun and Xin, 2003), Miller and Peskin(Miller and Peskin, 2005), and Chang and Sohn (Chang and Sohn,2006) to investigate flight aerodynamics using mathematical and physical models.

Either a single clap upstroke or a single fling downstroke was simulated. This simplification was made because the influence of the wake produced by the previous stroke is small for Re of 10 and below. Chang and Sohn(Chang and Sohn, 2006) found changes in the strength of the leading edge vortices at Re on the order of 100 when isolated clap and fling motions were compared with cyclical clap and fling motions however, the overall aerodynamics and forces acting on the wings at Re=10 were quite similar.

Translational and rotational velocities as functions of dimensionless time during the clap and fling. (A) For 0% overlap during clap, translation ends before rotation begins. For 100% overlap, translation and rotation end simultaneously. (B) For 0% overlap during fling, the wing completes its rotation before translation begins. For 100% overlap, the wing begins to rotate and translate simultaneously.

Translational and rotational velocities as functions of dimensionless time during the clap and fling. (A) For 0% overlap during clap, translation ends before rotation begins. For 100% overlap, translation and rotation end simultaneously. (B) For 0% overlap during fling, the wing completes its rotation before translation begins. For 100% overlap, the wing begins to rotate and translate simultaneously.


Here's how insects coax plants into making galls

Most of the galls made by Hormaphis cornu aphids are green. But mutations in one gene trigger the development of a red gall instead. Credit: David Stern

Insects can reprogram plant growth, transforming ordinary plant parts into intricately patterned shelters that are safe havens for feeding and reproduction.

These structures, called galls, have fascinated biologists for centuries. They're crafted by a variety of insects, including some species of aphids, mites, and wasps. And they take on innumerable forms, each specific in shape and size to the insect species that's created it—from knobs to cone-shaped protrusions to long, thin spikes. Some even resemble flowers.

Insects create galls by manipulating the development of plants, but figuring out exactly how they perform this feat "feels like one of the great unsolved problems in biology," says David Stern, a group leader at the Howard Hughes Medical Institute's Janelia Research Campus. "How does an organism of one kingdom take control of the genome of an organism in another kingdom to completely reorganize its development, to produce a home for itself?"

Now, Stern and his colleagues have identified the first examples of insect genes that directly guide gall development. These genes are turned on in aphids' salivary glands and appear to direct gall formation when the insects spit their saliva into the plants. One gene the team identified determines whether such galls will be red or green, the researchers report in a paper published March 2 in Current Biology.

"I think they've discovered essentially new territory," says Patrick Abbot, a molecular ecologist at Vanderbilt University who wasn't involved in the work. There's a strong likelihood that similar genes are found in other insects, he says. "It makes me want to run to the lab and start looking back through my data."

Figuring out how to study gall formation has been a longstanding challenge, Stern says—one that's interested him since he was a graduate student doing fieldwork in Malaysia. Gall-making insects aren't laboratory model organisms like fruit flies, and not as much is known about their genetics.

Hormaphis cornu aphids feed on witch hazel leaves and coax the plants into making galls. Credit: David Stern

A few years ago, while wandering the woods of Janelia's riverside campus, Stern made a convenient observation. Hormaphis cornu aphids make galls on witch hazel trees, small flowering trees that are abundant on campus. Even on a single leaf, Stern noticed, some Hormaphis aphids were making green galls, while others were making red ones. It set up a natural experiment—a chance to compare two visibly distinct kinds of galls and figure out what's genetically different between the aphids that make them.

When Stern and his team sequenced the genomes of aphids that made green galls and those that made red galls, they pinpointed a gene that varied between the two genomes. Aphids with one version of a gene that they named "determinant of gall color" made green galls aphids with a different version made red ones. The finding piqued their curiosity, as the gene didn't look like any previously identified genes.

To dive deeper, they collected aphids from both witch hazel trees and river birch trees. (Hormaphis cornu aphids live on river birch trees in the summer, but don't make galls there.) Back in the lab, the researchers carefully dissected out the insects' tiny salivary glands. In these glands, the team hunted for genes that were turned on only in the aphids that made galls. The researchers found that the gene determinant of gall color was similar to hundreds of other genes that were all turned on specifically in the gall forming aphids. Stern's team dubbed this group bicycle genes.

The gall-making aphids on the witch hazel trees switch on these genes to make BICYCLE proteins. The insects might spit these proteins into plant cells to reprogram leaf tissue into making a gall instead of normal plant parts, says Aishwarya Korgaonkar, a research scientist in the Stern lab who helped lead the project.

The team is now working to identify the plant molecules targeted by the aphids' BICYCLE proteins, says Korgaonkar. That could help them understand just how BICYCLE proteins goad plants into forming galls.

"After years of wondering what's going on, it's very rewarding to have something to show for it," Stern says.


Whiteflies

Whiteflies are tiny, snow-white insect pests that (when viewed under a magnifying glass) resemble moths. When viewed without magnification, these insects look more like flying dandruff! Although they might resemble moths, they are actually more related to scale insects. In fact, they are often confused with soft scale insects. Both adult and nymph stages feed by sucking plant juices. Heavy feeding by these pests can give plants a mottled look, cause yellowing and eventually death to the host plant.

Sticky honeydew excreted by these insects glazes both upper and lower leaf surfaces, permitting the development of black sooty mold fungus. Besides being unattractive, sooty mold interferes with photosynthesis, which retards plant growth and often causes leaf drop.

The most common and perhaps most difficult to control insect pests in greenhouses and interior landscapes are whiteflies. Three common species of whiteflies, the greenhouse, sweet potato and banded wing, are potential pests on a wide variety of crops. They attack a wide range of plants including bedding plants, cotton, strawberries, vegetables, and poinsettias. In addition to attacking many different crops, whiteflies are difficult to control. The immature stages are small and difficult to detect. Growers often buy plants, unaware of the whitefly infestation present.

Once adults develop and emerge inside a greenhouse or hothouse, they quickly become distributed over an entire crop or infest other available plants. Chemical control programs directed at the pest often have limited success. Two life stages (egg and pupa) are tolerant of most insecticides. Control measures are also complicated by the insects clinging on to the underside of leaves, making them difficult to reach with chemical or oil sprays.

All species of this plant pest develop from the egg through four nymphal instars before becoming adults. Elapsed time (from egg to adult) varies with species. Eggs are deposited on the undersides of leaves and are often found in a circular or crescent-shaped pattern. The "crawler" hatches from the egg, moves a short distance and then settles and begins feeding -- sucking juices from its plant host. The remainder of the nymphal development is spent in this sedentary condition. The adult whitefly emerges from the pupal case and flies to other host plants to lay eggs and begin the cycle again. Fourth instar nymphs (called pupae) and adults are most frequently used to distinguish one species from another.
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When choosing a product for eliminating whiteflies from your flowers and plants, remember that each product might kill only specific stages of the pest. You might also consider that the preferred product can have other uses, such as indoor or outdoor pest control.
For example, Pesticide oil sprays and Safer Insecticide soap do little damage to adult whiteflies they mainly eliminate nymph and pupa stages of the whitefly. Talstar One Bifenthrin Concentrate, Permethrin Pro, Tempo SC and Pyrethrin-Rotenone sprays eliminate adult and nymphs only. Oil Spray is best for year-round prevention. While oil sprays, Safer Soap and Pyrethrin-Rotenone are used extensively on plants, they are not the products of choice when treating homes for general purpose pest control.
Permethrin Pro and Tempo SC can be used in a wide variety of situations: indoor pest control (boxelder bugs, roaches, ants, silverfish, etc.), outdoor pest control (ornamentals) and (in the case of Tempo SC) can be used for treating restaurants and other commercial food plants. For best, long-term control of plant pests Talstar One usually works far better than other sprays, producing excellent knock down of existing white flies as well as longer residual than other insecticides.
Choose the product best suited for your over all needs.

Click on any product link to see description and availability in certain states.

Apply your insecticide when first stage nymphs or adults have emerged. In heavy whitefly populations of mixed life stages, two to three applications per week may be necessary to bring the population under control with a contact insecticide. Read and follow label instructions each product can have different limits on how often applications can be made.

Proper application of the insecticide is also a key component to a successful pest control program. It is necessary to deliver the insecticide to the undersides of leaves to achieve good control. As many crops mature, a dense canopy of foliage forms that interferes with pesticide delivery. With these crops, it is necessary to control whiteflies prior to the formation of this canopy or to space plants so they can be treated adequately.


MOTH FLIES

The key identifying character for the moth fly is the unique pattern of veins in its wings. The entire body and wings of the moth fly are covered with tiny hairs, giving it a moth-like appearance. To the naked eye, this tiny pest might appear to be a small fly with fat wings the aid of a magnifying glass reveals the unmistakable moth-like appearance. This small fly is no more than 1/8 inch in length including the wings. They are usually black in color.

Moth Fly Biology and Life History

Moth fly adults can be quite annoying in homes, appearing from sinks and bathtub drains. These pests breed in tremendous numbers in sewer plants and are easily blown towards homes by the wind. Their small size enables them to penetrate ordinary fly screens. There have been noted cases of bronchial asthma caused by inhaling the dust resulting from the disintegration of such small flies.

Moth flies lay eggs in a mass of 30 to 100. These eggs hatch in less than 48 hours. The larvae and pupae of the moth fly live in the thin film found in drains, septic tank field lines or filter stones. The larvae feed on sediment, decaying vegetation and microscopic plants and animals. The larval stage lasts from 9 to 15 days and the pupal stage lasts from 20 to 40 hours. The newly emerged adult fly is sexually mature on emergence and copulates within the first few hours of its life.

Inspecting for Moth Flies

The key to eliminating a moth fly infestation is finding the breeding sources and eliminating them.

Moth fly larvae live in the moist film that develops on the sides of a drain and in the drain's trap. The presence of many adult flies inside a drain is a good sign that the drain is a breeding site. To check for possible breeding sites, place a length of tape across drains (or cracks in the floor) without totally covering the opening. If the opening is totally covered, there will be no air flow and flies will not emerge. Check the tape periodically. If flies are found stuck to your tape, you have found a source. Eliminating this source is discussed in Moth Fly Elimination.

The number one source of moth fly infestations seems to be septic or sewer problems that have not been detected. The appearance of moth flies is often the first indication of another problem! There are many areas in a building where moth flies can breed, so do not end your inspection after you find one source. These pests love the organic debris found in sewers, septic tanks, drains, wet brooms and mops, even the soil close to a leaking or ruptured plumbing line. In homes, moth flies are generally found breeding in bathroom drains, particularly those in showers. Shower pans are prone to leaking and the area under the shower pan becomes an excellent breeding ground for moth flies.

Do not overlook the outdoors in your inspection. Any sign of moisture on an exterior wall or under a home should be investigated.

Moth Fly Elimination

Although space sprays easily kill adult moth flies in a home and around patios and porches, they will not totally eliminate the fly infestation. Total control comes with locating and eliminating breeding sites discussed in Moth Fly Inspection. Use the Gold Stick pheromone trap to capture adult flies in kitchens, bathrooms or other areas where moth fly activity has been detected. If a drain is found to be a breeding ground, clean the drain thoroughly (scrubbing it , if possible) and use Invade Bio Drain Gel to destroy the film harboring the organisms.

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How these tiny insect larvae leap without legs

SPRING FLING Never mind about the absence of legs. A young gall midge, no bigger than a rice grain, can go airborne thanks to some clever latching.

G.M. FARLEY ET AL/JOURNAL OF EXPERIMENTAL BIOLOGY 2019

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No legs? Not a problem. Some pudgy insect larvae can still jump up to 36 times their body length. Now high-speed video reveals how.

First, a legless, bright orange Asphondylia gall midge larva fastens its body into a fat, lopsided O by meshing together front and rear patches of microscopic fuzz. The rear part of the larva swells, and starts to straighten like a long, overinflating balloon. The fuzzy surfaces then pop apart. Then like a suddenly released spring, the larva flips up and away in an arc of somersaults, researchers report August 8 in the Journal of Experimental Biology.

In nature, something has to go wrong for this to happen, says evolutionary ecologist Michael Wise of Roanoke College in Salem, Va. These midges normally grow from egg to adult safely inside an abnormal growth, or gall, that they trick silverrod plants into forming. But as Wise was trying to coax out some still-immature larvae, he realized that the supposedly helpless young — extracted prematurely when they were no bigger than rice grains — could not only vault out of a lab dish but also could travel a fair distance across the lab floor.

To get a better look at the insects’ jumps, he contacted evolutionary biomechanist Sheila Patek at Duke University. “He sees small fast things and thinks of his buddy Sheila,” Patek says. Her lab specializes in resolving never-before-seen subtleties of animal motion, typically using high-speed video. “The truth is, we film for people all the time, and it’s almost never small and fast by our standards,” she says. “But this actually was.”

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The larval jumps filmed were too great for a tiny larva’s muscles, Wise, Patek and colleagues concluded. Blobby little larvae were flipping themselves around with power equal to, or greater than, the oomph of high-power vertebrate flight muscles.

For small animals with constraints on muscles, “it actually works better to put energy into a spring,” Patek says. Small creatures can load energy into the spring gradually until whatever is latching the spring slips off. Then, the suddenly freed spring powers extreme motion.

Microscope images revealed hairlike structures on the larval surfaces that touched, suggesting that the tiny projections might stick together as type of latch. Such structures could inspire new types of adhesives, Patek says.

Patek had first recognized the latch-and-spring system in mantis shrimp, which throw punches so furiously they can smash aquarium walls, and then in a trap-jaw ant with killer jaws that spring shut in an instant. Those look not at all like the pliable little larvae, but Patek sees latches releasing springs. “My guess is they’re everywhere,” she says.

Still “latch systems are quite hard to study,” says coauthor Greg Sutton of the University of Lincoln in England, where he investigates the mechanics of insect moves. “We don’t know where the flea latch is,” for example, he says. The gall midge ends up with arguably the most clearly described system: the “smoking gun latch,” he calls it.

Details aside, tiny animals aren’t the only creatures using latches for fast moves, says Simon Poppinga of the Botanic Garden of the University of Freiburg in Germany, who wasn’t involved in the research. He studies the biomechanics of plants, which don’t grow muscles at all but have ways of moving fast. U.S. researchers have found that sphagnum moss fires “spore cannons,” capsules that deform as they dry and then suddenly crack open to launch spores at 16 meters per second that puff into miniature mushroom clouds (SN: 7/23/10, p. 8).

Poppinga and colleagues recently showed that Chinese witch hazel trees build up forces in the mature fruit that suddenly shoot out a seed rotating a bit like a bullet from a rifle. Unlike gall midge launches, though, these tree latches break when they let go. The leap of a legless seed is fast and dramatic, but it’s not repeatable.

Questions or comments on this article? E-mail us at [email protected]

A version of this article appears in the August 31, 2019 issue of Science News.


Frequently bought together

Review

“Persuasive, rollicking, and informative. He may not get you to hug your termites, but you will see them in a whole new light. Bugs are truly awesome in numbers and variety. On the surface, bugs seem so alien to us. But in anecdote after anecdote, Waldbauer gives us plenty with which we can identify. Waldbauer celebrates not only the good things bugs do but also the bizarre. What Waldbauer shows us is that bugs are vitally important to our planet. They help plant life grow. They are great cleanup crews, removing waste material. They till and aerate soil. They provide food for all kinds of animals, including fish and birds and some mammals. Clearly, bugs are good.”Vicki Croke, Boston Globe

“This book will open the eyes of readers who, like the great majority of mankind, regard insects with contempt or disgust. It will make them look on our six-legged fellow creatures with more interest and sympathy, and will thus add a new dimension to their own lives.”Anthony Daniels, Sunday Telegraph (UK)

“Written in a gentle style that is easy to read yet still authoritative, the breadth of insect ecology is paraded before us.”Richard Jones, BBC Wildlife

“Waldbauer is an entomologist with an unwavering verve for his pursuits. Here he catalogs ecologically important insects by their 'occupations' within an ecosystem, explaining how they live and how they make possible life in general. Among insects' occupations are their roles in regulating plant and animal populations and tilling the soil. In some cases, their capabilities and behaviors are nothing short of mind-boggling. Waldbauer reports that one species of Great Plains ants has brought to the surface about 1.7 tons of subsoil per acre. An average colony of honeybees harvests 44 pounds of pollen and 265 pounds of nectar a year. Such anecdotes combine with the author's keen insight into the mechanics of ecosystems to make a strong case on behalf of the lowly insect.”Science News


Diversity

Insects are the most diverse of all groups of organisms, with over 800,000 species named and many thousands, probably millions, yet to be discovered. Insect diversity may be linked to their close association with the angiosperms (flowering plants). The Coleoptera (beetles) are the most diverse of all insect orders, with at least 350,000 species, representing one fourth of all known animal species. (Asked what could be inferred about the work of the Creator from a study of His works, British scientist J. B. S. Haldane is reported to have quipped, Ȫn inordinate fondness for beetles.") The evolutionary reasons for the mind-boggling diversity of this single order are not clear. Other major orders of insects include the Diptera (flies), Hymenoptera (bees and wasps), Hemiptera (true bugs), and Lepidoptera (moths and butterflies). Note that each name describes the wing ( ptera means "wing"). For instance, Diptera means "two wings," referring to the presence of only one wing pair in this order. In the Coleoptera ("sheath wings"), the first pair of wings is modified into a hard covering for the rear pair, which is easily observed in a lady beetle, for instance.


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