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And another insect from SE Brazil Oct 2017

And another insect from SE Brazil Oct 2017


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Photographed in the low areas of the Brazilian Atlantic Rainforest - Oct 2017 and about 2cm in length.


I'm having a difficult time pin-pointing the species, however, I'm fairly certain this true bug belongs to the genus Zelurus.

Zelurus sp.

Zelurus sp.

Z. variegatus

Distribution

Interactive version can be found here.

source


Great answer from @Charles: I was still humming and hawing between a zelurus species and another of the zelus species such as the longpipe that has the distinct red colour. (I had to leave for a couple hours… ) Like him, I was having trouble finding one that looked really similar. But I found one, labeled as the Zelurus Circumcintus:

However, further research revealed that this is a fairly undocumented species. The only source I could find with pictures of a distinctly red one was Flickr, such as this one:

I'm guessing it actually is the zelurus circumcintus, but I have yet to find any official sources that back me up.


And another insect from SE Brazil Oct 2017 - Biology

Over 40% of insect species are threatened with extinction.

Lepidoptera, Hymenoptera and dung beetles (Coleoptera) are the taxa most affected.

Four aquatic taxa are imperiled and have already lost a large proportion of species.

Habitat loss by conversion to intensive agriculture is the main driver of the declines.

Agro-chemical pollutants, invasive species and climate change are additional causes.


Flies' disease-carrying potential may be greater than thought, researchers say

Researchers used a scan electron microscope to find where bacterial cells and particles attach to the fly body. The electron microscope captures an up close look at the head of a blowfly in this picture. Credit: Ana Junqueira and Stephan Schuster

Flies can be more than pesky picnic crashers, they may be potent pathogen carriers, too, according to an international team of researchers.

In a study of the microbiomes of 116 houseflies and blowflies from three different continents, researchers found, in some cases, these flies carried hundreds of different species of bacteria, many of which are harmful to humans. Because flies often live close to humans, scientists have long suspected they played a role in carrying and spreading diseases, but this study, which was originally initiated at Penn State's Eberly College of Science, adds further proof, as well as insights into the extent of that threat.

"We believe that this may show a mechanism for pathogen transmission that has been overlooked by public health officials, and flies may contribute to the rapid transmission of pathogens in outbreak situations," said Donald Bryant, Ernest C. Pollard Professor of Biotechnology and professor of biochemistry and molecular biology, Penn State.

According to Stephan Schuster, former professor of biochemistry and molecular biology, Penn State, and now research director at Nanyang Technological University, Singapore, the researchers were able to investigate the microbial content of individual fly body parts, including legs and wings. The legs appear to transfer most of the microbial organisms from one surface to another, he added.

"The legs and wings show the highest microbial diversity in the fly body, suggesting that bacteria use the flies as airborne shuttles," said Schuster. "It may be that bacteria survive their journey, growing and spreading on a new surface. In fact, the study shows that each step of hundreds that a fly has taken leaves behind a microbial colony track, if the new surface supports bacterial growth."

Blowflies and houseflies—both carrion fly species—are often exposed to unhygienic matter because they use feces and decaying organic matter to nurture their young, where they could pick up bacteria that could act as pathogens to humans, plants and animals. The study also indicates that blowflies and houseflies share over 50 percent of their microbiome, a mixture of host-related microorganisms and those acquired from the environments they inhabit. Surprisingly, flies collected from stables carried fewer pathogens than those collected from urban environments.

A study initiated at Penn State's Eberly College of Science adds further proof to the suspicion that houseflies and blowflies carry and spread a variety of species of bacteria that are harmful to humans. The high number of bristles on the fly body gives bacteria lots of places to attach and be transported from one location to another. Credit: Ana Junqueira and Stephan Schuster

The researchers, who report their findings in the current issue of Scientific Reports, found 15 instances of the human pathogen Helicobacter pylori, a pathogen often causing ulcers in the human gut, largely in the blowfly samples collected in Brazil. The known route of transmission of Helicobacter has never considered flies as a possible vector for the disease, said Schuster.

The potential, then, for flies to carry diseases may increase when more people are present.

"It will really make you think twice about eating that potato salad that's been sitting out at your next picnic," Bryant said. "It might be better to have that picnic in the woods, far away from urban environments, not a central park."

Ana Carolina Junqueira, professor of genetics and genomics at the Federal University of Rio De Janeiro and previous postdoctoral fellow at the Singapore Centre for Environmental Life Sciences Engineering (SCELSE), said that the novel genomic and computational methods used for the study allowed the team an unprecedented look at the microbial community carried by flies.

"This is the first study that depicts the entire microbial DNA content of insect vectors using unbiased methods," Junqueira said. "Blowflies and houseflies are considered major mechanical vectors worldwide, but their full potential for microbial transmission was never analyzed comprehensively using modern molecular techniques and deep DNA sequencing."

Flies may not be all bad, however. The researchers suggest they could turn into helpers for human society, perhaps even serving as living drones that can act as an early-warning system for diseases.

"For one, the environmental sequencing of flies may use the insects as proxies that can inform on the microbial content of any given environment that otherwise would be hard or impossible to sample," said Schuster. "In fact, the flies could be intentionally released as autonomous bionic drones into even the smallest spaces and crevices and, upon being recaptured, inform about any biotic material they have encountered."


Contents

The word "insect" comes from the Latin word insectum, meaning "with a notched or divided body", or literally "cut into", from the neuter singular perfect passive participle of insectare, "to cut into, to cut up", from in- "into" and secare "to cut" [10] because insects appear "cut into" three sections. A calque of Greek ἔντομον [éntomon], "cut into sections", Pliny the Elder introduced the Latin designation as a loan-translation of the Greek word ἔντομος (éntomos) or "insect" (as in entomology), which was Aristotle's term for this class of life, also in reference to their "notched" bodies. "Insect" first appears documented in English in 1601 in Holland's translation of Pliny. Translations of Aristotle's term also form the usual word for "insect" in Welsh (trychfil, from trychu "to cut" and mil, "animal"), Serbo-Croatian (zareznik, from rezati, "to cut"), Russian ( насекомое nasekomoje, from seč'/-sekat', "to cut"), etc. [10] [11]

The precise definition of the taxon Insecta and the equivalent English name "insect" varies three alternative definitions are shown in the table.

Definition of Insecta
Group Alternative definitions
Collembola (springtails) Insecta sensu lato
=Hexapoda
Entognatha
(paraphyletic)
Apterygota
(wingless hexapods)
(paraphyletic)
Protura (coneheads)
Diplura (two-pronged bristletails)
Archaeognatha (jumping bristletails) Insecta sensu stricto
=Ectognatha
Zygentoma (silverfish)
Pterygota (winged insects) Insecta sensu strictissimo

In the broadest circumscription, Insecta sensu lato consists of all hexapods. [12] [13] Traditionally, insects defined in this way were divided into "Apterygota" (the first five groups in the table)—the wingless insects—and Pterygota—the winged and secondarily wingless insects. [14] However, modern phylogenetic studies have shown that "Apterygota" is not monophyletic, [15] and so does not form a good taxon. A narrower circumscription restricts insects to those hexapods with external mouthparts, and comprises only the last three groups in the table. In this sense, Insecta sensu stricto is equivalent to Ectognatha. [12] [15] In the narrowest circumscription, insects are restricted to hexapods that are either winged or descended from winged ancestors. Insecta sensu strictissimo is then equivalent to Pterygota. [16] For the purposes of this article, the middle definition is used insects consist of two wingless taxa, Archaeognatha (jumping bristletails) and Zygentoma (silverfish), plus the winged or secondarily wingless Pterygota.

A phylogenetic tree of the arthropods and related groups [17]

Although traditionally grouped with millipedes and centipedes—possibly on the basis of convergent adaptations to terrestrialisation [18] —evidence has emerged favoring closer evolutionary ties with crustaceans. In the Pancrustacea theory, insects, together with Entognatha, Remipedia, and Cephalocarida, make up a natural clade labeled Miracrustacea within Crustaceans, now termed Pancrustacea. [19]

Insects form a single clade, closely related to crustaceans and myriapods. [20]

Other terrestrial arthropods, such as centipedes, millipedes, scorpions, spiders, woodlice, mites, and ticks are sometimes confused with insects since their body plans can appear similar, sharing (as do all arthropods) a jointed exoskeleton. However, upon closer examination, their features differ significantly most noticeably, they do not have the six-legged characteristic of adult insects. [21]

The higher-level phylogeny of the arthropods continues to be a matter of debate and research. In 2008, researchers at Tufts University uncovered what they believe is the world's oldest known full-body impression of a primitive flying insect, a 300-million-year-old specimen from the Carboniferous period. [22] The oldest insect fossil was considered to be the Devonian Rhyniognatha hirsti, from the 396-million-year-old Rhynie chert. [23] However, other analyses have disputed this placement, finding it to be more likely a myriapod. [24]

Four super radiations of insects have occurred: beetles (from about 300 million years ago), flies (from about 250 million years ago), moths and wasps (both from about 150 million years ago). [25] These four groups account for the majority of described species. The flies and moths along with the fleas evolved from the Mecoptera.

The origins of insect flight remain obscure, since the earliest winged insects currently known appear to have been capable fliers. Some extinct insects had an additional pair of winglets attaching to the first segment of the thorax, for a total of three pairs. As of 2009, no evidence suggests the insects were a particularly successful group of animals before they evolved to have wings. [26]

Late Carboniferous and Early Permian insect orders include both extant groups, their stem groups, [27] and a number of Paleozoic groups, now extinct. During this era, some giant dragonfly-like forms reached wingspans of 55 to 70 cm (22 to 28 in), making them far larger than any living insect. This gigantism may have been due to higher atmospheric oxygen levels that allowed increased respiratory efficiency relative to today. The lack of flying vertebrates could have been another factor. Most extinct orders of insects developed during the Permian period that began around 270 million years ago. Many of the early groups became extinct during the Permian-Triassic extinction event, the largest mass extinction in the history of the Earth, around 252 million years ago. [28]

The remarkably successful Hymenoptera appeared as long as 200 million years ago in the Triassic period, but achieved their wide diversity more recently in the Cenozoic era, which began 66 million years ago. A number of highly successful insect groups evolved in conjunction with flowering plants, a powerful illustration of coevolution. [29]

Many modern insect genera developed during the Cenozoic. Insects from this period on are often found preserved in amber, often in perfect condition. The body plan, or morphology, of such specimens is thus easily compared with modern species. The study of fossilized insects is called paleoentomology.

Phylogeny

Zygentoma (silverfish, firebrats, fishmoths)

Orthoptera (grasshoppers, crickets, katydids)

Psocodea (Book lice, barklice & sucking lice)

A cladogram based on the works of Sroka, Staniczek & Bechly 2014, [30] Prokop et al. 2017 [31] & Wipfler et al. 2019. [32]

Taxonomy

Cladogram of living insect groups, [33] with numbers of species in each group. [4] The Apterygota, Palaeoptera, and Exopterygota are possibly paraphyletic groups.

Traditional morphology-based or appearance-based systematics have usually given the Hexapoda the rank of superclass, [34] : 180 and identified four groups within it: insects (Ectognatha), springtails (Collembola), Protura, and Diplura, the latter three being grouped together as the Entognatha on the basis of internalized mouth parts. Supraordinal relationships have undergone numerous changes with the advent of methods based on evolutionary history and genetic data. A recent theory is that the Hexapoda are polyphyletic (where the last common ancestor was not a member of the group), with the entognath classes having separate evolutionary histories from the Insecta. [35] Many of the traditional appearance-based taxa have been shown to be paraphyletic, so rather than using ranks like subclass, superorder, and infraorder, it has proved better to use monophyletic groupings (in which the last common ancestor is a member of the group). The following represents the best-supported monophyletic groupings for the Insecta.

Insects can be divided into two groups historically treated as subclasses: wingless insects, known as Apterygota, and winged insects, known as Pterygota. The Apterygota consist of the primitively wingless order of the silverfish (Zygentoma). Archaeognatha make up the Monocondylia based on the shape of their mandibles, while Zygentoma and Pterygota are grouped together as Dicondylia. The Zygentoma themselves possibly are not monophyletic, with the family Lepidotrichidae being a sister group to the Dicondylia (Pterygota and the remaining Zygentoma). [36] [37]

Paleoptera and Neoptera are the winged orders of insects differentiated by the presence of hardened body parts called sclerites, and in the Neoptera, muscles that allow their wings to fold flatly over the abdomen. Neoptera can further be divided into incomplete metamorphosis-based (Polyneoptera and Paraneoptera) and complete metamorphosis-based groups. It has proved difficult to clarify the relationships between the orders in Polyneoptera because of constant new findings calling for revision of the taxa. For example, the Paraneoptera have turned out to be more closely related to the Endopterygota than to the rest of the Exopterygota. The recent molecular finding that the traditional louse orders Mallophaga and Anoplura are derived from within Psocoptera has led to the new taxon Psocodea. [38] Phasmatodea and Embiidina have been suggested to form the Eukinolabia. [39] Mantodea, Blattodea, and Isoptera are thought to form a monophyletic group termed Dictyoptera. [40]

The Exopterygota likely are paraphyletic in regard to the Endopterygota. Matters that have incurred controversy include Strepsiptera and Diptera grouped together as Halteria based on a reduction of one of the wing pairs—a position not well-supported in the entomological community. [41] The Neuropterida are often lumped or split on the whims of the taxonomist. Fleas are now thought to be closely related to boreid mecopterans. [42] Many questions remain in the basal relationships among endopterygote orders, particularly the Hymenoptera.

The study of the classification or taxonomy of any insect is called systematic entomology. If one works with a more specific order or even a family, the term may also be made specific to that order or family, for example systematic dipterology.

Evolutionary relationships

Insects are prey for a variety of organisms, including terrestrial vertebrates. The earliest vertebrates on land existed 400 million years ago and were large amphibious piscivores. Through gradual evolutionary change, insectivory was the next diet type to evolve. [43]

Insects were among the earliest terrestrial herbivores and acted as major selection agents on plants. [29] Plants evolved chemical defenses against this herbivory and the insects, in turn, evolved mechanisms to deal with plant toxins. Many insects make use of these toxins to protect themselves from their predators. Such insects often advertise their toxicity using warning colors. [44] This successful evolutionary pattern has also been used by mimics. Over time, this has led to complex groups of coevolved species. Conversely, some interactions between plants and insects, like pollination, are beneficial to both organisms. Coevolution has led to the development of very specific mutualisms in such systems.

Estimates on the total number of insect species, or those within specific orders, often vary considerably. Globally, averages of these estimates suggest there are around 1.5 million beetle species and 5.5 million insect species, with about 1 million insect species currently found and described. [45]

Between 950,000 and 1,000,000 of all described species are insects, so over 50% of all described eukaryotes (1.8 million) are insects (see illustration). With only 950,000 known non-insects, if the actual number of insects is 5.5 million, they may represent over 80% of the total. As only about 20,000 new species of all organisms are described each year, most insect species may remain undescribed, unless the rate of species descriptions greatly increases. Of the 24 orders of insects, four dominate in terms of numbers of described species at least 670,000 identified species belong to Coleoptera, Diptera, Hymenoptera or Lepidoptera.

As of 2017, at least 66 insect species extinctions had been recorded in the previous 500 years, which generally occurred on oceanic islands. [47] Declines in insect abundance have been attributed to artificial lighting, [48] land use changes such as urbanization or agricultural use, [49] [50] pesticide use, [51] and invasive species. [52] Studies summarized in a 2019 review suggested a large proportion of insect species are threatened with extinction in the 21st century. [53] Though ecologist Manu Sanders notes the 2019 review was biased by mostly excluding data showing increases or stability in insect population, with the studies limited to specific geographic areas and specific groups of species. [54] A larger meta-study published in 2020, analyzing data from 166 long-term surveys, suggested that populations of terrestrial insects are decreasing by about 9% per decade. [55] [56] Claims of pending mass insect extinctions or "insect apocalypse" based on a subset of these studies have been popularized in news reports, but often extrapolate beyond the study data or hyperbolize study findings. [57] Other areas have shown increases in some insect species, although trends in most regions are currently unknown. It is difficult to assess long-term trends in insect abundance or diversity because historical measurements are generally not known for many species. Robust data to assess at-risk areas or species is especially lacking for arctic and tropical regions and a majority of the southern hemisphere. [57]

Estimates of total extant insect species [45]
Order Estimated total species
Archaeognatha 513
Zygentoma 560
Ephemeroptera 3,240
Odonata 5,899
Orthoptera 23,855
Neuroptera 5,868
Phasmatodea 3,014
Embioptera 463
Grylloblattodea 34
Mantophasmatodea 20
Plecoptera 3,743
Dermaptera 1,978
Zoraptera 37
Mantodea 2,400
Blattodea 7,314
Psocoptera 5,720
Phthiraptera 5,102
Thysanoptera 5,864
Hemiptera 103,590
Hymenoptera 116,861
Strepsiptera 609
Coleoptera 386,500
Megaloptera 354
Raphidioptera 254
Trichoptera 14,391
Lepidoptera 157,338
Diptera 155,477
Siphonaptera 2,075
Mecoptera 757

External

Insects have segmented bodies supported by exoskeletons, the hard outer covering made mostly of chitin. The segments of the body are organized into three distinctive but interconnected units, or tagmata: a head, a thorax and an abdomen. [58] The head supports a pair of sensory antennae, a pair of compound eyes, zero to three simple eyes (or ocelli) and three sets of variously modified appendages that form the mouthparts. The thorax is made up of three segments: the prothorax, mesothorax and the metathorax. Each thoracic segment supports one pair of legs. The meso- and metathoracic segments may each have a pair of wings, depending on the insect. The abdomen consists of eleven segments, though in a few species of insects, these segments may be fused together or reduced in size. The abdomen also contains most of the digestive, respiratory, excretory and reproductive internal structures. [34] : 22–48 Considerable variation and many adaptations in the body parts of insects occur, especially wings, legs, antenna and mouthparts.

Segmentation

The head is enclosed in a hard, heavily sclerotized, unsegmented, exoskeletal head capsule, or epicranium, which contains most of the sensing organs, including the antennae, ocellus or eyes, and the mouthparts. Of all the insect orders, Orthoptera displays the most features found in other insects, including the sutures and sclerites. [59] Here, the vertex, or the apex (dorsal region), is situated between the compound eyes for insects with a hypognathous and opisthognathous head. In prognathous insects, the vertex is not found between the compound eyes, but rather, where the ocelli are normally. This is because the primary axis of the head is rotated 90° to become parallel to the primary axis of the body. In some species, this region is modified and assumes a different name. [59] : 13

The thorax is a tagma composed of three sections, the prothorax, mesothorax and the metathorax. The anterior segment, closest to the head, is the prothorax, with the major features being the first pair of legs and the pronotum. The middle segment is the mesothorax, with the major features being the second pair of legs and the anterior wings. The third and most posterior segment, abutting the abdomen, is the metathorax, which features the third pair of legs and the posterior wings. Each segment is delineated by an intersegmental suture. Each segment has four basic regions. The dorsal surface is called the tergum (or notum) to distinguish it from the abdominal terga. [34] The two lateral regions are called the pleura (singular: pleuron) and the ventral aspect is called the sternum. In turn, the notum of the prothorax is called the pronotum, the notum for the mesothorax is called the mesonotum and the notum for the metathorax is called the metanotum. Continuing with this logic, the mesopleura and metapleura, as well as the mesosternum and metasternum, are used. [59]

The abdomen is the largest tagma of the insect, which typically consists of 11–12 segments and is less strongly sclerotized than the head or thorax. Each segment of the abdomen is represented by a sclerotized tergum and sternum. Terga are separated from each other and from the adjacent sterna or pleura by membranes. Spiracles are located in the pleural area. Variation of this ground plan includes the fusion of terga or terga and sterna to form continuous dorsal or ventral shields or a conical tube. Some insects bear a sclerite in the pleural area called a laterotergite. Ventral sclerites are sometimes called laterosternites. During the embryonic stage of many insects and the postembryonic stage of primitive insects, 11 abdominal segments are present. In modern insects there is a tendency toward reduction in the number of the abdominal segments, but the primitive number of 11 is maintained during embryogenesis. Variation in abdominal segment number is considerable. If the Apterygota are considered to be indicative of the ground plan for pterygotes, confusion reigns: adult Protura have 12 segments, Collembola have 6. The orthopteran family Acrididae has 11 segments, and a fossil specimen of Zoraptera has a 10-segmented abdomen. [59]

Exoskeleton

The insect outer skeleton, the cuticle, is made up of two layers: the epicuticle, which is a thin and waxy water resistant outer layer and contains no chitin, and a lower layer called the procuticle. The procuticle is chitinous and much thicker than the epicuticle and has two layers: an outer layer known as the exocuticle and an inner layer known as the endocuticle. The tough and flexible endocuticle is built from numerous layers of fibrous chitin and proteins, criss-crossing each other in a sandwich pattern, while the exocuticle is rigid and hardened. [34] : 22–24 The exocuticle is greatly reduced in many insects during their larval stages, e.g., caterpillars. It is also reduced in soft-bodied adult insects.

Insects are the only invertebrates to have developed active flight capability, and this has played an important role in their success. [34] : 186 Their flight muscles are able to contract multiple times for each single nerve impulse, allowing the wings to beat faster than would ordinarily be possible.

Having their muscles attached to their exoskeletons is efficient and allows more muscle connections.

Internal

Nervous system

The nervous system of an insect can be divided into a brain and a ventral nerve cord. The head capsule is made up of six fused segments, each with either a pair of ganglia, or a cluster of nerve cells outside of the brain. The first three pairs of ganglia are fused into the brain, while the three following pairs are fused into a structure of three pairs of ganglia under the insect's esophagus, called the subesophageal ganglion. [34] : 57

The thoracic segments have one ganglion on each side, which are connected into a pair, one pair per segment. This arrangement is also seen in the abdomen but only in the first eight segments. Many species of insects have reduced numbers of ganglia due to fusion or reduction. [60] Some cockroaches have just six ganglia in the abdomen, whereas the wasp Vespa crabro has only two in the thorax and three in the abdomen. Some insects, like the house fly Musca domestica, have all the body ganglia fused into a single large thoracic ganglion. [ citation needed ]

At least a few insects have nociceptors, cells that detect and transmit signals responsible for the sensation of pain. [61] [ failed verification ] This was discovered in 2003 by studying the variation in reactions of larvae of the common fruit-fly Drosophila to the touch of a heated probe and an unheated one. The larvae reacted to the touch of the heated probe with a stereotypical rolling behavior that was not exhibited when the larvae were touched by the unheated probe. [62] Although nociception has been demonstrated in insects, there is no consensus that insects feel pain consciously [63]

Insects are capable of learning. [64]

Digestive system

An insect uses its digestive system to extract nutrients and other substances from the food it consumes. [65] Most of this food is ingested in the form of macromolecules and other complex substances like proteins, polysaccharides, fats and nucleic acids. These macromolecules must be broken down by catabolic reactions into smaller molecules like amino acids and simple sugars before being used by cells of the body for energy, growth, or reproduction. This break-down process is known as digestion.

There is extensive variation among different orders, life stages, and even castes in the digestive system of insects. [66] This is the result of extreme adaptations to various lifestyles. The present description focuses on a generalized composition of the digestive system of an adult orthopteroid insect, which is considered basal to interpreting particularities of other groups.

The main structure of an insect's digestive system is a long enclosed tube called the alimentary canal, which runs lengthwise through the body. The alimentary canal directs food unidirectionally from the mouth to the anus. It has three sections, each of which performs a different process of digestion. In addition to the alimentary canal, insects also have paired salivary glands and salivary reservoirs. These structures usually reside in the thorax, adjacent to the foregut. [34] : 70–77 The salivary glands (element 30 in numbered diagram) in an insect's mouth produce saliva. The salivary ducts lead from the glands to the reservoirs and then forward through the head to an opening called the salivarium, located behind the hypopharynx. By moving its mouthparts (element 32 in numbered diagram) the insect can mix its food with saliva. The mixture of saliva and food then travels through the salivary tubes into the mouth, where it begins to break down. [67] [68] Some insects, like flies, have extra-oral digestion. Insects using extra-oral digestion expel digestive enzymes onto their food to break it down. This strategy allows insects to extract a significant proportion of the available nutrients from the food source. [69] : 31 The gut is where almost all of insects' digestion takes place. It can be divided into the foregut, midgut and hindgut.

Foregut

The first section of the alimentary canal is the foregut (element 27 in numbered diagram), or stomodaeum. The foregut is lined with a cuticular lining made of chitin and proteins as protection from tough food. The foregut includes the buccal cavity (mouth), pharynx, esophagus and crop and proventriculus (any part may be highly modified), which both store food and signify when to continue passing onward to the midgut. [34] : 70

Digestion starts in buccal cavity (mouth) as partially chewed food is broken down by saliva from the salivary glands. As the salivary glands produce fluid and carbohydrate-digesting enzymes (mostly amylases), strong muscles in the pharynx pump fluid into the buccal cavity, lubricating the food like the salivarium does, and helping blood feeders, and xylem and phloem feeders.

From there, the pharynx passes food to the esophagus, which could be just a simple tube passing it on to the crop and proventriculus, and then onward to the midgut, as in most insects. Alternately, the foregut may expand into a very enlarged crop and proventriculus, or the crop could just be a diverticulum, or fluid-filled structure, as in some Diptera species. [69] : 30–31

Midgut

Once food leaves the crop, it passes to the midgut (element 13 in numbered diagram), also known as the mesenteron, where the majority of digestion takes place. Microscopic projections from the midgut wall, called microvilli, increase the surface area of the wall and allow more nutrients to be absorbed they tend to be close to the origin of the midgut. In some insects, the role of the microvilli and where they are located may vary. For example, specialized microvilli producing digestive enzymes may more likely be near the end of the midgut, and absorption near the origin or beginning of the midgut. [69] : 32

Hindgut

In the hindgut (element 16 in numbered diagram), or proctodaeum, undigested food particles are joined by uric acid to form fecal pellets. The rectum absorbs 90% of the water in these fecal pellets, and the dry pellet is then eliminated through the anus (element 17), completing the process of digestion. Envaginations at the anterior end of the hindgut form the Malpighian tubules, which form the main excretory system of insects.

Excretory system

Insects may have one to hundreds of Malpighian tubules (element 20). These tubules remove nitrogenous wastes from the hemolymph of the insect and regulate osmotic balance. Wastes and solutes are emptied directly into the alimentary canal, at the junction between the midgut and hindgut. [34] : 71–72, 78–80

Reproductive system

The reproductive system of female insects consist of a pair of ovaries, accessory glands, one or more spermathecae, and ducts connecting these parts. The ovaries are made up of a number of egg tubes, called ovarioles, which vary in size and number by species. The number of eggs that the insect is able to make vary by the number of ovarioles with the rate that eggs can develop being also influenced by ovariole design. Female insects are able make eggs, receive and store sperm, manipulate sperm from different males, and lay eggs. Accessory glands or glandular parts of the oviducts produce a variety of substances for sperm maintenance, transport and fertilization, as well as for protection of eggs. They can produce glue and protective substances for coating eggs or tough coverings for a batch of eggs called oothecae. Spermathecae are tubes or sacs in which sperm can be stored between the time of mating and the time an egg is fertilized. [59] : 880

For males, the reproductive system is the testis, suspended in the body cavity by tracheae and the fat body. Most male insects have a pair of testes, inside of which are sperm tubes or follicles that are enclosed within a membranous sac. The follicles connect to the vas deferens by the vas efferens, and the two tubular vasa deferentia connect to a median ejaculatory duct that leads to the outside. A portion of the vas deferens is often enlarged to form the seminal vesicle, which stores the sperm before they are discharged into the female. The seminal vesicles have glandular linings that secrete nutrients for nourishment and maintenance of the sperm. The ejaculatory duct is derived from an invagination of the epidermal cells during development and, as a result, has a cuticular lining. The terminal portion of the ejaculatory duct may be sclerotized to form the intromittent organ, the aedeagus. The remainder of the male reproductive system is derived from embryonic mesoderm, except for the germ cells, or spermatogonia, which descend from the primordial pole cells very early during embryogenesis. [59] : 885

Respiratory system

Insect respiration is accomplished without lungs. Instead, the insect respiratory system uses a system of internal tubes and sacs through which gases either diffuse or are actively pumped, delivering oxygen directly to tissues that need it via their trachea (element 8 in numbered diagram). In most insects, air is taken in through openings on the sides of the abdomen and thorax called spiracles.

The respiratory system is an important factor that limits the size of insects. As insects get larger, this type of oxygen transport is less efficient and thus the heaviest insect currently weighs less than 100 g. However, with increased atmospheric oxygen levels, as were present in the late Paleozoic, larger insects were possible, such as dragonflies with wingspans of more than two feet (60 cm). [70]

There are many different patterns of gas exchange demonstrated by different groups of insects. Gas exchange patterns in insects can range from continuous and diffusive ventilation, to discontinuous gas exchange. [34] : 65–68 During continuous gas exchange, oxygen is taken in and carbon dioxide is released in a continuous cycle. In discontinuous gas exchange, however, the insect takes in oxygen while it is active and small amounts of carbon dioxide are released when the insect is at rest. [71] Diffusive ventilation is simply a form of continuous gas exchange that occurs by diffusion rather than physically taking in the oxygen. Some species of insect that are submerged also have adaptations to aid in respiration. As larvae, many insects have gills that can extract oxygen dissolved in water, while others need to rise to the water surface to replenish air supplies, which may be held or trapped in special structures. [72] [73]

Circulatory system

Because oxygen is delivered directly to tissues via tracheoles, the circulatory system is not used to carry oxygen, and is therefore greatly reduced. The insect circulatory system is open it has no veins or arteries, and instead consists of little more than a single, perforated dorsal tube that pulses peristaltically. This dorsal blood vessel (element 14) is divided into two sections: the heart and aorta. The dorsal blood vessel circulates the hemolymph, arthropods' fluid analog of blood, from the rear of the body cavity forward. [34] : 61–65 [74] Hemolymph is composed of plasma in which hemocytes are suspended. Nutrients, hormones, wastes, and other substances are transported throughout the insect body in the hemolymph. Hemocytes include many types of cells that are important for immune responses, wound healing, and other functions. Hemolymph pressure may be increased by muscle contractions or by swallowing air into the digestive system to aid in molting. [75] Hemolymph is also a major part of the open circulatory system of other arthropods, such as spiders and crustaceans. [76] [77]

The majority of insects hatch from eggs. The fertilization and development takes place inside the egg, enclosed by a shell (chorion) that consists of maternal tissue. In contrast to eggs of other arthropods, most insect eggs are drought resistant. This is because inside the chorion two additional membranes develop from embryonic tissue, the amnion and the serosa. This serosa secretes a cuticle rich in chitin that protects the embryo against desiccation. In Schizophora however the serosa does not develop, but these flies lay their eggs in damp places, such as rotting matter. [78] Some species of insects, like the cockroach Blaptica dubia, as well as juvenile aphids and tsetse flies, are ovoviviparous. The eggs of ovoviviparous animals develop entirely inside the female, and then hatch immediately upon being laid. [6] Some other species, such as those in the genus of cockroaches known as Diploptera, are viviparous, and thus gestate inside the mother and are born alive. [34] : 129, 131, 134–135 Some insects, like parasitic wasps, show polyembryony, where a single fertilized egg divides into many and in some cases thousands of separate embryos. [34] : 136–137 Insects may be univoltine, bivoltine or multivoltine, i.e. they may have one, two or many broods (generations) in a year. [79]

Other developmental and reproductive variations include haplodiploidy, polymorphism, paedomorphosis or peramorphosis, sexual dimorphism, parthenogenesis and more rarely hermaphroditism. [34] : 143 [80] In haplodiploidy, which is a type of sex-determination system, the offspring's sex is determined by the number of sets of chromosomes an individual receives. This system is typical in bees and wasps. [81] Polymorphism is where a species may have different morphs or forms, as in the oblong winged katydid, which has four different varieties: green, pink and yellow or tan. Some insects may retain phenotypes that are normally only seen in juveniles this is called paedomorphosis. In peramorphosis, an opposite sort of phenomenon, insects take on previously unseen traits after they have matured into adults. Many insects display sexual dimorphism, in which males and females have notably different appearances, such as the moth Orgyia recens as an exemplar of sexual dimorphism in insects.

Some insects use parthenogenesis, a process in which the female can reproduce and give birth without having the eggs fertilized by a male. Many aphids undergo a form of parthenogenesis, called cyclical parthenogenesis, in which they alternate between one or many generations of asexual and sexual reproduction. [82] [83] In summer, aphids are generally female and parthenogenetic in the autumn, males may be produced for sexual reproduction. Other insects produced by parthenogenesis are bees, wasps and ants, in which they spawn males. However, overall, most individuals are female, which are produced by fertilization. The males are haploid and the females are diploid. [6]

Insect life-histories show adaptations to withstand cold and dry conditions. Some temperate region insects are capable of activity during winter, while some others migrate to a warmer climate or go into a state of torpor. [84] Still other insects have evolved mechanisms of diapause that allow eggs or pupae to survive these conditions. [85]

Metamorphosis

Metamorphosis in insects is the biological process of development all insects must undergo. There are two forms of metamorphosis: incomplete metamorphosis and complete metamorphosis.

Incomplete metamorphosis

Hemimetabolous insects, those with incomplete metamorphosis, change gradually by undergoing a series of molts. An insect molts when it outgrows its exoskeleton, which does not stretch and would otherwise restrict the insect's growth. The molting process begins as the insect's epidermis secretes a new epicuticle inside the old one. After this new epicuticle is secreted, the epidermis releases a mixture of enzymes that digests the endocuticle and thus detaches the old cuticle. When this stage is complete, the insect makes its body swell by taking in a large quantity of water or air, which makes the old cuticle split along predefined weaknesses where the old exocuticle was thinnest. [34] : 142 [86]

Immature insects that go through incomplete metamorphosis are called nymphs or in the case of dragonflies and damselflies, also naiads. Nymphs are similar in form to the adult except for the presence of wings, which are not developed until adulthood. With each molt, nymphs grow larger and become more similar in appearance to adult insects.

Complete metamorphosis

Holometabolism, or complete metamorphosis, is where the insect changes in four stages, an egg or embryo, a larva, a pupa and the adult or imago. In these species, an egg hatches to produce a larva, which is generally worm-like in form. This worm-like form can be one of several varieties: eruciform (caterpillar-like), scarabaeiform (grub-like), campodeiform (elongated, flattened and active), elateriform (wireworm-like) or vermiform (maggot-like). The larva grows and eventually becomes a pupa, a stage marked by reduced movement and often sealed within a cocoon. There are three types of pupae: obtect, exarate or coarctate. Obtect pupae are compact, with the legs and other appendages enclosed. Exarate pupae have their legs and other appendages free and extended. Coarctate pupae develop inside the larval skin. [34] : 151 Insects undergo considerable change in form during the pupal stage, and emerge as adults. Butterflies are a well-known example of insects that undergo complete metamorphosis, although most insects use this life cycle. Some insects have evolved this system to hypermetamorphosis.

Complete metamorphosis is a trait of the most diverse insect group, the Endopterygota. [34] : 143 Endopterygota includes 11 Orders, the largest being Diptera (flies), Lepidoptera (butterflies and moths), and Hymenoptera (bees, wasps, and ants), and Coleoptera (beetles). This form of development is exclusive to insects and not seen in any other arthropods.

Many insects possess very sensitive and specialized organs of perception. Some insects such as bees can perceive ultraviolet wavelengths, or detect polarized light, while the antennae of male moths can detect the pheromones of female moths over distances of many kilometers. [87] The yellow paper wasp (Polistes versicolor) is known for its wagging movements as a form of communication within the colony it can waggle with a frequency of 10.6±2.1 Hz (n=190). These wagging movements can signal the arrival of new material into the nest and aggression between workers can be used to stimulate others to increase foraging expeditions. [88] There is a pronounced tendency for there to be a trade-off between visual acuity and chemical or tactile acuity, such that most insects with well-developed eyes have reduced or simple antennae, and vice versa. There are a variety of different mechanisms by which insects perceive sound while the patterns are not universal, insects can generally hear sound if they can produce it. Different insect species can have varying hearing, though most insects can hear only a narrow range of frequencies related to the frequency of the sounds they can produce. Mosquitoes have been found to hear up to 2 kHz, and some grasshoppers can hear up to 50 kHz. [89] Certain predatory and parasitic insects can detect the characteristic sounds made by their prey or hosts, respectively. For instance, some nocturnal moths can perceive the ultrasonic emissions of bats, which helps them avoid predation. [34] : 87–94 Insects that feed on blood have special sensory structures that can detect infrared emissions, and use them to home in on their hosts.

Some insects display a rudimentary sense of numbers, [90] such as the solitary wasps that prey upon a single species. The mother wasp lays her eggs in individual cells and provides each egg with a number of live caterpillars on which the young feed when hatched. Some species of wasp always provide five, others twelve, and others as high as twenty-four caterpillars per cell. The number of caterpillars is different among species, but always the same for each sex of larva. The male solitary wasp in the genus Eumenes is smaller than the female, so the mother of one species supplies him with only five caterpillars the larger female receives ten caterpillars in her cell.

Light production and vision

A few insects, such as members of the families Poduridae and Onychiuridae (Collembola), Mycetophilidae (Diptera) and the beetle families Lampyridae, Phengodidae, Elateridae and Staphylinidae are bioluminescent. The most familiar group are the fireflies, beetles of the family Lampyridae. Some species are able to control this light generation to produce flashes. The function varies with some species using them to attract mates, while others use them to lure prey. Cave dwelling larvae of Arachnocampa (Mycetophilidae, fungus gnats) glow to lure small flying insects into sticky strands of silk. [91] Some fireflies of the genus Photuris mimic the flashing of female Photinus species to attract males of that species, which are then captured and devoured. [92] The colors of emitted light vary from dull blue (Orfelia fultoni, Mycetophilidae) to the familiar greens and the rare reds (Phrixothrix tiemanni, Phengodidae). [93]

Most insects, except some species of cave crickets, are able to perceive light and dark. Many species have acute vision capable of detecting minute movements. The eyes may include simple eyes or ocelli as well as compound eyes of varying sizes. Many species are able to detect light in the infrared, ultraviolet and the visible light wavelengths. Color vision has been demonstrated in many species and phylogenetic analysis suggests that UV-green-blue trichromacy existed from at least the Devonian period between 416 and 359 million years ago. [94]

Sound production and hearing

Insects were the earliest organisms to produce and sense sounds. Insects make sounds mostly by mechanical action of appendages. In grasshoppers and crickets, this is achieved by stridulation. Cicadas make the loudest sounds among the insects by producing and amplifying sounds with special modifications to their body to form tymbals and associated musculature. The African cicada Brevisana brevis has been measured at 106.7 decibels at a distance of 50 cm (20 in). [95] Some insects, such as the Helicoverpa zea moths, hawk moths and Hedylid butterflies, can hear ultrasound and take evasive action when they sense that they have been detected by bats. [96] [97] Some moths produce ultrasonic clicks that were once thought to have a role in jamming bat echolocation. The ultrasonic clicks were subsequently found to be produced mostly by unpalatable moths to warn bats, just as warning colorations are used against predators that hunt by sight. [98] Some otherwise palatable moths have evolved to mimic these calls. [99] More recently, the claim that some moths can jam bat sonar has been revisited. Ultrasonic recording and high-speed infrared videography of bat-moth interactions suggest the palatable tiger moth really does defend against attacking big brown bats using ultrasonic clicks that jam bat sonar. [100]

Very low sounds are also produced in various species of Coleoptera, Hymenoptera, Lepidoptera, Mantodea and Neuroptera. These low sounds are simply the sounds made by the insect's movement. Through microscopic stridulatory structures located on the insect's muscles and joints, the normal sounds of the insect moving are amplified and can be used to warn or communicate with other insects. Most sound-making insects also have tympanal organs that can perceive airborne sounds. Some species in Hemiptera, such as the corixids (water boatmen), are known to communicate via underwater sounds. [101] Most insects are also able to sense vibrations transmitted through surfaces.

Communication using surface-borne vibrational signals is more widespread among insects because of size constraints in producing air-borne sounds. [102] Insects cannot effectively produce low-frequency sounds, and high-frequency sounds tend to disperse more in a dense environment (such as foliage), so insects living in such environments communicate primarily using substrate-borne vibrations. [103] The mechanisms of production of vibrational signals are just as diverse as those for producing sound in insects.

Some species use vibrations for communicating within members of the same species, such as to attract mates as in the songs of the shield bug Nezara viridula. [104] Vibrations can also be used to communicate between entirely different species lycaenid (gossamer-winged butterfly) caterpillars, which are myrmecophilous (living in a mutualistic association with ants) communicate with ants in this way. [105] The Madagascar hissing cockroach has the ability to press air through its spiracles to make a hissing noise as a sign of aggression [106] the death's-head hawkmoth makes a squeaking noise by forcing air out of their pharynx when agitated, which may also reduce aggressive worker honey bee behavior when the two are in close proximity. [107]

Chemical communication

Chemical communications in animals rely on a variety of aspects including taste and smell. Chemoreception is the physiological response of a sense organ (i.e. taste or smell) to a chemical stimulus where the chemicals act as signals to regulate the state or activity of a cell. A semiochemical is a message-carrying chemical that is meant to attract, repel, and convey information. Types of semiochemicals include pheromones and kairomones. One example is the butterfly Phengaris arion which uses chemical signals as a form of mimicry to aid in predation. [108]

In addition to the use of sound for communication, a wide range of insects have evolved chemical means for communication. These chemicals, termed semiochemicals, are often derived from plant metabolites including those meant to attract, repel and provide other kinds of information. Pheromones, a type of semiochemical, are used for attracting mates of the opposite sex, for aggregating conspecific individuals of both sexes, for deterring other individuals from approaching, to mark a trail, and to trigger aggression in nearby individuals. Allomones benefit their producer by the effect they have upon the receiver. Kairomones benefit their receiver instead of their producer. Synomones benefit the producer and the receiver. While some chemicals are targeted at individuals of the same species, others are used for communication across species. The use of scents is especially well known to have developed in social insects. [34] : 96–105

Social insects, such as termites, ants and many bees and wasps, are the most familiar species of eusocial animals. [109] They live together in large well-organized colonies that may be so tightly integrated and genetically similar that the colonies of some species are sometimes considered superorganisms. It is sometimes argued that the various species of honey bee are the only invertebrates (and indeed one of the few non-human groups) to have evolved a system of abstract symbolic communication where a behavior is used to represent and convey specific information about something in the environment. In this communication system, called dance language, the angle at which a bee dances represents a direction relative to the sun, and the length of the dance represents the distance to be flown. [34] : 309–311 Though perhaps not as advanced as honey bees, bumblebees also potentially have some social communication behaviors. Bombus terrestris, for example, exhibit a faster learning curve for visiting unfamiliar, yet rewarding flowers, when they can see a conspecific foraging on the same species. [110]

Only insects that live in nests or colonies demonstrate any true capacity for fine-scale spatial orientation or homing. This can allow an insect to return unerringly to a single hole a few millimeters in diameter among thousands of apparently identical holes clustered together, after a trip of up to several kilometers' distance. In a phenomenon known as philopatry, insects that hibernate have shown the ability to recall a specific location up to a year after last viewing the area of interest. [111] A few insects seasonally migrate large distances between different geographic regions (e.g., the overwintering areas of the monarch butterfly). [34] : 14

Care of young

The eusocial insects build nests, guard eggs, and provide food for offspring full-time (see Eusociality). Most insects, however, lead short lives as adults, and rarely interact with one another except to mate or compete for mates. A small number exhibit some form of parental care, where they will at least guard their eggs, and sometimes continue guarding their offspring until adulthood, and possibly even feeding them. Another simple form of parental care is to construct a nest (a burrow or an actual construction, either of which may be simple or complex), store provisions in it, and lay an egg upon those provisions. The adult does not contact the growing offspring, but it nonetheless does provide food. This sort of care is typical for most species of bees and various types of wasps. [112]

Flight

Insects are the only group of invertebrates to have developed flight. The evolution of insect wings has been a subject of debate. Some entomologists suggest that the wings are from paranotal lobes, or extensions from the insect's exoskeleton called the nota, called the paranotal theory. Other theories are based on a pleural origin. These theories include suggestions that wings originated from modified gills, spiracular flaps or as from an appendage of the epicoxa. The epicoxal theory suggests the insect wings are modified epicoxal exites, a modified appendage at the base of the legs or coxa. [113] In the Carboniferous age, some of the Meganeura dragonflies had as much as a 50 cm (20 in) wide wingspan. The appearance of gigantic insects has been found to be consistent with high atmospheric oxygen. The respiratory system of insects constrains their size, however the high oxygen in the atmosphere allowed larger sizes. [114] The largest flying insects today are much smaller, with the largest wingspan belonging to the white witch moth (Thysania agrippina), at approximately 28 cm (11 in). [115]

Insect flight has been a topic of great interest in aerodynamics due partly to the inability of steady-state theories to explain the lift generated by the tiny wings of insects. But insect wings are in motion, with flapping and vibrations, resulting in churning and eddies, and the misconception that physics says "bumblebees can't fly" persisted throughout most of the twentieth century.

Unlike birds, many small insects are swept along by the prevailing winds [116] although many of the larger insects are known to make migrations. Aphids are known to be transported long distances by low-level jet streams. [117] As such, fine line patterns associated with converging winds within weather radar imagery, like the WSR-88D radar network, often represent large groups of insects. [118]

Walking

Many adult insects use six legs for walking and have adopted a tripedal gait. The tripedal gait allows for rapid walking while always having a stable stance and has been studied extensively in cockroaches and ants. The legs are used in alternate triangles touching the ground. For the first step, the middle right leg and the front and rear left legs are in contact with the ground and move the insect forward, while the front and rear right leg and the middle left leg are lifted and moved forward to a new position. When they touch the ground to form a new stable triangle the other legs can be lifted and brought forward in turn and so on. [119] The purest form of the tripedal gait is seen in insects moving at high speeds. However, this type of locomotion is not rigid and insects can adapt a variety of gaits. For example, when moving slowly, turning, avoiding obstacles, climbing or slippery surfaces, four (tetrapod) or more feet (wave-gait [120] ) may be touching the ground. Insects can also adapt their gait to cope with the loss of one or more limbs.

Cockroaches are among the fastest insect runners and, at full speed, adopt a bipedal run to reach a high velocity in proportion to their body size. As cockroaches move very quickly, they need to be video recorded at several hundred frames per second to reveal their gait. More sedate locomotion is seen in the stick insects or walking sticks (Phasmatodea). A few insects have evolved to walk on the surface of the water, especially members of the Gerridae family, commonly known as water striders. A few species of ocean-skaters in the genus Halobates even live on the surface of open oceans, a habitat that has few insect species. [121]

Use in robotics

Insect walking is of particular interest as an alternative form of locomotion in robots. The study of insects and bipeds has a significant impact on possible robotic methods of transport. This may allow new robots to be designed that can traverse terrain that robots with wheels may be unable to handle. [119]

Swimming

A large number of insects live either part or the whole of their lives underwater. In many of the more primitive orders of insect, the immature stages are spent in an aquatic environment. Some groups of insects, like certain water beetles, have aquatic adults as well. [72]

Many of these species have adaptations to help in under-water locomotion. Water beetles and water bugs have legs adapted into paddle-like structures. Dragonfly naiads use jet propulsion, forcibly expelling water out of their rectal chamber. [122] Some species like the water striders are capable of walking on the surface of water. They can do this because their claws are not at the tips of the legs as in most insects, but recessed in a special groove further up the leg this prevents the claws from piercing the water's surface film. [72] Other insects such as the Rove beetle Stenus are known to emit pygidial gland secretions that reduce surface tension making it possible for them to move on the surface of water by Marangoni propulsion (also known by the German term Entspannungsschwimmen). [123] [124]

Insect ecology is the scientific study of how insects, individually or as a community, interact with the surrounding environment or ecosystem. [125] : 3 Insects play one of the most important roles in their ecosystems, which includes many roles, such as soil turning and aeration, dung burial, pest control, pollination and wildlife nutrition. An example is the beetles, which are scavengers that feed on dead animals and fallen trees and thereby recycle biological materials into forms found useful by other organisms. [126] These insects, and others, are responsible for much of the process by which topsoil is created. [34] : 3, 218–228

Defense and predation

Insects are mostly soft bodied, fragile and almost defenseless compared to other, larger lifeforms. The immature stages are small, move slowly or are immobile, and so all stages are exposed to predation and parasitism. Insects then have a variety of defense strategies to avoid being attacked by predators or parasitoids. These include camouflage, mimicry, toxicity and active defense. [128]

Camouflage is an important defense strategy, which involves the use of coloration or shape to blend into the surrounding environment. [129] This sort of protective coloration is common and widespread among beetle families, especially those that feed on wood or vegetation, such as many of the leaf beetles (family Chrysomelidae) or weevils. In some of these species, sculpturing or various colored scales or hairs cause the beetle to resemble bird dung or other inedible objects. Many of those that live in sandy environments blend in with the coloration of the substrate. [128] Most phasmids are known for effectively replicating the forms of sticks and leaves, and the bodies of some species (such as O. macklotti and Palophus centaurus) are covered in mossy or lichenous outgrowths that supplement their disguise. Very rarely, a species may have the ability to change color as their surroundings shift (Bostra scabrinota). In a further behavioral adaptation to supplement crypsis, a number of species have been noted to perform a rocking motion where the body is swayed from side to side that is thought to reflect the movement of leaves or twigs swaying in the breeze. Another method by which stick insects avoid predation and resemble twigs is by feigning death (catalepsy), where the insect enters a motionless state that can be maintained for a long period. The nocturnal feeding habits of adults also aids Phasmatodea in remaining concealed from predators. [130]

Another defense that often uses color or shape to deceive potential enemies is mimicry. A number of longhorn beetles (family Cerambycidae) bear a striking resemblance to wasps, which helps them avoid predation even though the beetles are in fact harmless. [128] Batesian and Müllerian mimicry complexes are commonly found in Lepidoptera. Genetic polymorphism and natural selection give rise to otherwise edible species (the mimic) gaining a survival advantage by resembling inedible species (the model). Such a mimicry complex is referred to as Batesian. One of the most famous examples, where the viceroy butterfly was long believed to be a Batesian mimic of the inedible monarch, was later disproven, as the viceroy is more toxic than the monarch, and this resemblance is now considered to be a case of Müllerian mimicry. [127] In Müllerian mimicry, inedible species, usually within a taxonomic order, find it advantageous to resemble each other so as to reduce the sampling rate by predators who need to learn about the insects' inedibility. Taxa from the toxic genus Heliconius form one of the most well known Müllerian complexes. [131]

Chemical defense is another important defense found among species of Coleoptera and Lepidoptera, usually being advertised by bright colors, such as the monarch butterfly. They obtain their toxicity by sequestering the chemicals from the plants they eat into their own tissues. Some Lepidoptera manufacture their own toxins. Predators that eat poisonous butterflies and moths may become sick and vomit violently, learning not to eat those types of species this is actually the basis of Müllerian mimicry. A predator who has previously eaten a poisonous lepidopteran may avoid other species with similar markings in the future, thus saving many other species as well. [132] Some ground beetles of the family Carabidae can spray chemicals from their abdomen with great accuracy, to repel predators. [128]

Pollination

Pollination is the process by which pollen is transferred in the reproduction of plants, thereby enabling fertilisation and sexual reproduction. Most flowering plants require an animal to do the transportation. While other animals are included as pollinators, the majority of pollination is done by insects. [133] Because insects usually receive benefit for the pollination in the form of energy rich nectar it is a grand example of mutualism. The various flower traits (and combinations thereof) that differentially attract one type of pollinator or another are known as pollination syndromes. These arose through complex plant-animal adaptations. Pollinators find flowers through bright colorations, including ultraviolet, and attractant pheromones. The study of pollination by insects is known as anthecology.

Parasitism

Many insects are parasites of other insects such as the parasitoid wasps. These insects are known as entomophagous parasites. They can be beneficial due to their devastation of pests that can destroy crops and other resources. Many insects have a parasitic relationship with humans such as the mosquito. These insects are known to spread diseases such as malaria and yellow fever and because of such, mosquitoes indirectly cause more deaths of humans than any other animal.

As pests

Many insects are considered pests by humans. Insects commonly regarded as pests include those that are parasitic (e.g. lice, bed bugs), transmit diseases (mosquitoes, flies), damage structures (termites), or destroy agricultural goods (locusts, weevils). Many entomologists are involved in various forms of pest control, as in research for companies to produce insecticides, but increasingly rely on methods of biological pest control, or biocontrol. Biocontrol uses one organism to reduce the population density of another organism—the pest—and is considered a key element of integrated pest management. [134] [135]

Despite the large amount of effort focused at controlling insects, human attempts to kill pests with insecticides can backfire. If used carelessly, the poison can kill all kinds of organisms in the area, including insects' natural predators, such as birds, mice and other insectivores. The effects of DDT's use exemplifies how some insecticides can threaten wildlife beyond intended populations of pest insects. [136] [137]

In beneficial roles

Although pest insects attract the most attention, many insects are beneficial to the environment and to humans. Some insects, like wasps, bees, butterflies and ants, pollinate flowering plants. Pollination is a mutualistic relationship between plants and insects. As insects gather nectar from different plants of the same species, they also spread pollen from plants on which they have previously fed. This greatly increases plants' ability to cross-pollinate, which maintains and possibly even improves their evolutionary fitness. This ultimately affects humans since ensuring healthy crops is critical to agriculture. As well as pollination ants help with seed distribution of plants. This helps to spread the plants, which increases plant diversity. This leads to an overall better environment. [138] A serious environmental problem is the decline of populations of pollinator insects, and a number of species of insects are now cultured primarily for pollination management in order to have sufficient pollinators in the field, orchard or greenhouse at bloom time. [139] : 240–243 Another solution, as shown in Delaware, has been to raise native plants to help support native pollinators like L. vierecki. [140]

The economic value of pollination by insects has been estimated to be about $34 billion in the US alone. [141]

Products made by insects. Insects also produce useful substances such as honey, wax, lacquer and silk. Honey bees have been cultured by humans for thousands of years for honey, although contracting for crop pollination is becoming more significant for beekeepers. The silkworm has greatly affected human history, as silk-driven trade established relationships between China and the rest of the world.

Pest control. Insectivorous insects, or insects that feed on other insects, are beneficial to humans if they eat insects that could cause damage to agriculture and human structures. For example, aphids feed on crops and cause problems for farmers, but ladybugs feed on aphids, and can be used as a means to significantly reduce pest aphid populations. While birds are perhaps more visible predators of insects, insects themselves account for the vast majority of insect consumption. Ants also help control animal populations by consuming small vertebrates. [142] Without predators to keep them in check, insects can undergo almost unstoppable population explosions. [34] : 328–348 [34] : 400 [143] [144]

Medical uses. Insects are also used in medicine, for example fly larvae (maggots) were formerly used to treat wounds to prevent or stop gangrene, as they would only consume dead flesh. This treatment is finding modern usage in some hospitals. Recently insects have also gained attention as potential sources of drugs and other medicinal substances. [145] Adult insects, such as crickets and insect larvae of various kinds, are also commonly used as fishing bait. [146]

In research

Insects play important roles in biological research. For example, because of its small size, short generation time and high fecundity, the common fruit fly Drosophila melanogaster is a model organism for studies in the genetics of higher eukaryotes. D. melanogaster has been an essential part of studies into principles like genetic linkage, interactions between genes, chromosomal genetics, development, behavior and evolution. Because genetic systems are well conserved among eukaryotes, understanding basic cellular processes like DNA replication or transcription in fruit flies can help to understand those processes in other eukaryotes, including humans. [147] The genome of D. melanogaster was sequenced in 2000, reflecting the organism's important role in biological research. It was found that 70% of the fly genome is similar to the human genome, supporting the evolution theory. [148]

As food

In some cultures, insects, especially deep-fried cicadas, are considered to be delicacies, whereas in other places they form part of the normal diet. Insects have a high protein content for their mass, and some authors suggest their potential as a major source of protein in human nutrition. [34] : 10–13 In most first-world countries, however, entomophagy (the eating of insects), is taboo. [149] Since it is impossible to entirely eliminate pest insects from the human food chain, insects are inadvertently present in many foods, especially grains. Food safety laws in many countries do not prohibit insect parts in food, but rather limit their quantity. According to cultural materialist anthropologist Marvin Harris, the eating of insects is taboo in cultures that have other protein sources such as fish or livestock.

Due to the abundance of insects and a worldwide concern of food shortages, the Food and Agriculture Organization of the United Nations considers that the world may have to, in the future, regard the prospects of eating insects as a food staple. Insects are noted for their nutrients, having a high content of protein, minerals and fats and are eaten by one-third of the global population. [150]

As feed

Several insect species such as the black soldier fly or the housefly in their maggot forms, as well as beetle larvae such as mealworms can be processed and used as feed for farmed animals such as chicken, fish and pigs. [151]

In other products

Insect larvae (i.e. black soldier fly larvae) can provide protein, grease, and chitin. The grease is usable in the pharmaceutical industry (cosmetics, [152] surfactants for shower gel) -hereby replacing other vegetable oils as palm oil. [153]

Also, insect cooking oil, insect butter and fatty alcohols can be made from such insects as the superworm (Zophobas morio). [154] [155]

As pets

Many species of insects are sold and kept as pets. There are even special hobbyist magazines such as "Bugs" (now discontinued). [156]

In culture

Scarab beetles held religious and cultural symbolism in Old Egypt, Greece and some shamanistic Old World cultures. The ancient Chinese regarded cicadas as symbols of rebirth or immortality. In Mesopotamian literature, the epic poem of Gilgamesh has allusions to Odonata that signify the impossibility of immortality. Among the Aborigines of Australia of the Arrernte language groups, honey ants and witchetty grubs served as personal clan totems. In the case of the 'San' bush-men of the Kalahari, it is the praying mantis that holds much cultural significance including creation and zen-like patience in waiting. [34] : 9


Introduction

Dengue is second only to malaria as most important mosquito-borne disease. Unlike malaria and other major infectious diseases, dengue is increasing in incidence and severity, currently inflicting 50–390 million or more cases per year worldwide [1, 2]. Dengue is widespread in tropical and sub-tropical areas and is primarily associated with its principal vector Aedes aegypti (L.).

Dengue was reintroduced in Brazil in 1981 (Boa Vista, State of Roraima), after being almost entirely absent for at least 20 years following DDT-based vector control. Brazil now has serotypes 1–3 circulating throughout the country in addition serotype 4 was recently detected in several states [3]. In an analysis of dengue in the Americas in 2000–2007, Brazil was found to have the highest number of cases and economic burden [4] more recently [1] estimated 16 million total infections annually. While this in part reflects the size of the Brazilian population, Wilder-Smith et al. [5] concluded that the dengue burden is at least as high as the burden of other major infectious diseases that afflict the Brazilian population, including malaria. A cross-sectional seroepidemiologic survey conducted in Recife, state of Pernambuco, Brazil, in 2006 found overall dengue virus IgG prevalence to be 80% indicating that the large majority of inhabitants have been infected at least once [6] these authors estimated that 5.2% of susceptible individuals become infected each year by each serotype and that this had increased sharply over the previous 20 years.

There are no specific drugs or licensed vaccine for dengue, so efforts to reduce transmission depend entirely on vector control [2]. However, even the most highly-resourced and well-implemented programmes, such as in Singapore, have not been able to prevent epidemic dengue using current methods [7–9]. Furthermore, existing control tools are threatened by actual or potential spread of resistance in the vector population. Therefore there is an urgent need to develop new methods. The use of transgenic vectors may provide a set of new methods for reducing the density or vectorial capacity of vector populations [10]. Here we describe a field evaluation of one prominent transgenic-vector strategy, the use of male mosquitoes carrying a lethal or autocidal transgene in a sterile-male-release system.

The Sterile Insect Technique (SIT) is a genetic control system based on the release of large numbers of radiation-sterilised insects. These mate with wild insects of the same species and thereby reduce the reproductive potential of the wild pest population, as they produce no or fewer viable offspring due to the radiation-induced presence of lethal mutations in their gametes [11, 12]. Though successfully used against several agricultural pests, trials against mosquitoes have met with less success [13, 14]. This is in part due to the somatic damage, and associated performance reduction in the sterile insects, which inevitably accompanies radiation-sterilisation. Interestingly, one successful example of SIT in mosquitoes used a chemosterilant in place of radiation [15]. Modern genetics can potentially overcome this problem, for example by using an engineered self-limiting gene, that is both repressible by an antidote provided in a managed rearing facility and when expressed in the absence of the repressor any insect carrying the gene results in mortality before the insect reaches functional adulthood, which may be used in place of radiation [16]. Operationally, the system would look very similar to SIT, and would share the clean, species-specific characteristics, and similarly benefit from the female-seeking ability of the released males. However the insects would not be irradiated, rather they would be homozygous for a transgene which, when transmitted to an embryo via the sperm, would lead to death of the zygote at some stage in development [17, 18]. As well as avoiding the need for radiation, by adjusting the time of death one can improve efficiency against target populations with significant density-dependence [19, 20]. Simulation modelling suggests that such a method would potentially be effective and economical against Ae. aegypti [19, 21].

After extensive laboratory development and testing, field testing of engineered insects has begun, with encouraging results. In particular, in the Cayman Islands a self-limiting strain of Ae. aegypti, OX513A, was shown to be able to compete successfully for wild mates, furthermore sustained release of OX513A males suppressed a wild population of Ae. aegypti [10, 22]. We tested whether this same strain and strategy could also be effective in Brazil. Within the overall project objective of evaluating OX513A technology in Brazil, we had three core technical activities. These were (i) to transfer the technology to Brazil, including adaptation and optimisation for local conditions (ii) to assess the field performance in terms of mating competitiveness of OX513A males in Brazil and (iii) to test the ability of OX513A males to suppress a wild Ae. aegypti population in this environment. The fourth core activity, which will be described in detail elsewhere, related to community engagement and regulatory activities.


Fig. 1.

Geographic distribution of Maconellicoccus hirsutus (Green) (Hemiptera: Pseudococcidae) in Brazil, and M. hirsutus collection locations in the state of Espírito Santo, Brazil.

Plant species from 19 families have been reported as hosts of M. hirsutus in Brazil, including those of economic importance, such as Malvaceae (n = 7) and Fabaceae (n = 6), which are the most important host plant families in Brazil based on number of host plant species (Table 1), although Moraceae and Euphorbiaceae have the highest number of M. hirsutus host species worldwide (García Morales et al. 2016). Thirty-seven plant species are now reported as hosts of M. hirsutus in Brazil.

Although M. hirsutus has a large number of natural enemies (Culik et al. 2013b Garcia Morales et al. 2016), none were observed in the present study. In Brazil, Anagyrus kamali Moursi (Hymenoptera: Encyrtidae), and Gyranusoidea indica Shafee, Alam & Agarwal (Hymenoptera: Encyrtidae) are reported as parasitoids of M. hirsutus, and Chilocorus nigrita (F.), Cryptolaemus montrouzieri Mulsant, Cycloneda sanguinea (L.), Eriopis (Germar), Exoplectra sp., Harmonia axyridis (Pallas), Tenuisvalvae notata (Mulsant) (all Coleoptera: Coccinellidae), and Ceraeochrysa sp. (Neuroptera: Chrysopidae) are listed as M. hirsutus predators (Culik et al. 2013a). Anagyrus kamali and C. montrouzieri have been the most common natural enemies associated with M. hirsutus in Brazil (Marsaro Junior et al. 2013 Peronti et al. 2016 Negrini et al. 2018). However, there have been no studies in Brazil to evaluate the establishment and efficiency of these natural enemies in agroeco-systems where M. hirsutus has caused damage (Morais 2016).

We thank the Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq), and the Fundação de Amparo à Pesquisa e Inovação do Espírito Santo (FAPES) for financial support.


Fig. 1.

Olfactory response of predator Salpingogaster nigra in an olfactometer, with different treatments: spittlebug nymphs vs. air foam produced by spittlebug vs. air spittlebug nymphs vs. foam produced by spittlebug, and air vs. air. An asterisk (*) means significant difference using χ 2 test at P < 0.05.

These studies demonstrate that chemical cues may be involved in host finding by the predator as it searches for nymphs. Our results show that although S. nigra adults, eggs, and pupae may be kept in the laboratory, mass breeding of the predator is not feasible due to low larval viability of larvae. Further studies are required to minimize such low viability and make possible mass rearing in the laboratory.

We thank the Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq), and Fundação de Amparo à Pesquisa do Estado de Minas Gerais (FAPEMIG) for supporting our research.


Acknowledgments

We would like to acknowledge Dr. Gene Fridman in the Johns Hopkins School of Medicine for lending the use of the Fridman lab’s Metrohm Autolab potentiostat for use in the cyclic voltammetry experiments. We would like to thank the Johns Hopkins Malaria Research Institute and the Department of Molecular Microbiology and Immunology Insectary and Parasite Core facility, and Dr. Joel Tang from the Johns Hopkins NMR Core facility for his help with the NMR experiments. We would also like to thank the entire Casadevall Lab for their suggestions and inputs during conversations and their feedback during lab meetings and presentations. Figures in S9 Fig were made using BioRender.


Molecular mechanisms of olfactory detection in insects: beyond receptors

Insects thrive in diverse ecological niches in large part because of their highly sophisticated olfactory systems. Over the last two decades, a major focus in the study of insect olfaction has been on the role of olfactory receptors in mediating neuronal responses to environmental chemicals. In vivo, these receptors operate in specialized structures, called sensilla, which comprise neurons and non-neuronal support cells, extracellular lymph fluid and a precisely shaped cuticle. While sensilla are inherent to odour sensing in insects, we are only just beginning to understand their construction and function. Here, we review recent work that illuminates how odour-evoked neuronal activity is impacted by sensillar morphology, lymph fluid biochemistry, accessory signalling molecules in neurons and the physiological crosstalk between sensillar cells. These advances reveal multi-layered molecular and cellular mechanisms that determine the selectivity, sensitivity and dynamic modulation of odour-evoked responses in insects.

1. Introduction

Insects are one of the most successful classes of eukaryotes on Earth, making up approximately half of all terrestrial species [1]. They occupy an incredibly diverse range of habitats, encompassing tropical forests, deserts and the extremes of the polar regions. Many species exert an important influence on human health through their roles as disease vectors [2], crop pollinators [3] and agricultural pests [4]. The ecological adaptability of insects relies, in part, on their sophisticated olfactory systems, which allow detection and responses to innumerable volatile signals in the environment.

Studies of anatomical, physiological and behavioural aspects of insect olfaction have a long history in the twentieth century, using diverse model species [5]. Over the last two decades, there has been a particular focus on identifying and functionally characterizing olfactory receptors [6–8], as well as the neuronal circuits in which they are expressed and the odour-driven behaviours they control [9,10], notably in Drosophila melanogaster. There are two main classes of insect olfactory receptors: odorant receptors (ORs) [11,12] and ionotropic receptors (IRs) [13]. ORs are a family of seven-pass transmembrane ion channels, while IRs are three-pass transmembrane proteins distantly related to synaptic ionotropic glutamate receptors (iGluRs) [6–8,14]. Most olfactory sensory neurons (OSNs) express two different ORs or IRs: a unique ‘tuning' receptor that recognizes a set of ligands or odorants, and a co-receptor (ORCO for ORs, and either IR8a or IR25a for IRs). These co-receptors are not known to recognize any naturally occurring ligands, but form heteromeric complexes with tuning receptors to enable sensory cilia targeting and signalling [15–17].

Although both ORs and IRs are—as odour-gated ion channels—theoretically sufficient to translate the presence of an odour into depolarization of cellular membranes, they operate within complex sensory structures called sensilla (figure 1a) [21]. Sensilla are apparent as hair-like projections on the external surface of insects' olfactory organs, the antennae and maxillary palps. Each sensillum overlies a stereotyped combination of OSNs (up to four in D. melanogaster), surrounded by various non-neuronal support cells. The ciliated dendrites of OSNs, where olfactory receptors are localized, are housed within the porous shaft of the sensillum and are bathed in lymph fluid. Such organization allows the neuronal sensory membranes to be in close proximity to the odorous environment but protected from physical damage.

Figure 1. Insect olfactory sensillar morphology. (a) Schematic representation of an olfactory sensillum (see text for details). Inset: representative electron microscopy images of the main morphological classes of olfactory sensilla, here from D. melanogaster antennae (adapted from [18]). (b) Electron microscopy image of a trichoid sensillum from B. mori [19]. (c) Electron microscopy image of a D. melanogaster trichoid sensillum (at4) prepared using the CryoChem method and imaged using en bloc heavy metal staining (adapted from [20]).

Here, we review recent investigations into the development, morphology, biochemistry and physiology of olfactory sensilla, as well as some pertinent examples from similar chemosensory sensilla that mediate taste perception in insects [22]. These advances highlight that the process of chemical detection relies on much more than the receptors alone.

2. The morphology and cell biology of olfactory sensilla

Several sensillar types exist (e.g. basiconic, trichoid and coeloconic), which are distinguished by numerous morphological characteristics: length, width, cuticle thickness, and number and size of pores (through which chemicals pass), and neuronal cilia branching complexity (figure 1a) [21]. The OSNs in different sensillar classes are often specialized for the detection of particular types of odours for example, trichoid sensilla neurons are required for pheromone detection, while those in basiconic sensilla mostly detect food-derived odours [9]. This functional relationship, together with the conservation of sensillar types across most insects, suggests that these morphological properties are important for their roles in odour detection.

The sensillar surface represents the first contact point between an odour molecule and the sensory apparatus. As such, early efforts sought detailed descriptions of external sensillar morphology using electron microscopy (EM) [21]. These studies have been extended recently by combining high-resolution atomic force microscopy (AFM) with computational modelling of odour molecule behaviour near the sensillum [23,24]. EM and AFM revealed that the trichoid sensilla of three different moth species—the corn earworm (Helicoverpa zea), the bella moth (Utethesia ornatrix) and the silk moth (Bombyx mori)—are covered with a series of pores and ridges (figure 1b) [19,23,24]. In B. mori, these morphological data were used to run aerodynamic simulations at the sensillar surface. These analyses suggested that the ridges help to create small vortices that could facilitate the delivery of pheromone molecules into the sensillar pores [24]. Such simulations may help explain surprising observations of early work using radiolabelled pheromones, which estimated that approximately 25% of pheromone molecules adsorbed onto the sensillar surface activate OSNs [25], an efficiency that is greater than 50-fold higher than that predicted by consideration of airflow and pore dimensions alone [24]. Other modelling approaches have considered odour aerodynamics in the context of entire olfactory organs [26], which exhibit substantial morphological diversity across species [27]. For example, many moth antennae comprise arrays of sensilla along multiple, parallel antennal branches, an organization that is likely to maximize the volume of air sifted to detect minute quantities of pheromones [27].

Formation of the sensillar cuticle depends upon the non-neuronal support cells, which secrete the constituent macromolecules, notably chitin and proteinaceous components [21]. Different regions of the hair are formed by distinct types of support cells, from which they get their name: thecogen (sheath cell), trichogen (shaft cell) and tormogen (socket cell) [21]. How the precise sensillar cuticle architecture is determined is largely unknown, but recent work in D. melanogaster provided important insights into the formation of the pores in the shaft, at least in maxillary palp basiconic sensilla. Transmission EM revealed that during basiconic sensillum development, the trichogen elongates from the external surface of the epithelium and develops undulations in its plasma membrane where the cuticle envelope layer is secreted [28]. Ultra-thin regions in this envelope that form between protrusions of the plasma membrane correlate with where pores will develop. Screening for genes expressed specifically in the trichogen during development, combined with RNA interference (RNAi)-based functional testing, identified the transmembrane protein Osiris 23/Gore-tex (Osi23) as an important contributor to this process in this sensillar class [28]. Loss of Osi23 led to the disappearance of the plasma membrane undulations, resulting in the formation of a sensillum surface lacking pores consequently, neuronal responses to odours are dramatically reduced [28]. Osi23 localizes to endosomes, but how it influences plasma membrane morphology is unknown. Interestingly, other members of the Osi family are expressed in cuticle-secreting cells elsewhere in the fly (e.g. those lining the tracheae), hinting at a common role for this insect-specific protein family in shaping cuticular structures [28].

The second key contact point for odours is on the cilia membranes where olfactory receptors are localized. While the construction of OSN cilia and targeting of receptors to this compartment is likely to rely on the conserved intraflagellar transport pathway that is central to the assembly of other types of cilia [29,30], additional potential molecular regulators of these processes have emerged from reverse and forward genetic studies in D. melanogaster. For example, inspired by the intimate relationship between cilia function and Hedgehog signalling in vertebrates [31], analysis of OSNs lacking different components of this pathway in flies revealed a contribution to the efficient cilia localization of ORs and robust odour-evoked responses [32]. Unexpectedly, localization of the co-receptor ORCO is apparently insensitive to loss of the Hedgehog pathway. This observation suggests that Hedgehog signalling is required for the assembly of OR/ORCO complexes and/or that ORCO subunits alone can use an independent transport pathway to cilia.

Unbiased genetic screens in D. melanogaster revealed a requirement for a lipid transporter homologue, ATP8B, in odour-evoked responses of several OSN classes, including those expressing OR67d, a receptor for the sex and aggregation pheromone 11-cis vaccenyl acetate (cVA) [33,34]. ATP8B is expressed and required in OSNs and localized to the ciliated dendrites. The transporter belongs to the P4-type ATPase family, which is thought to flip aminophospholipids (e.g. phosphatidylserine) between membrane leaflets. A predicted enzymatically inactive version of ATP8B fails to rescue the mutant phenotype, while a mammalian homologue can complement the defect, suggesting that the lipid flippase function is critical for its role in OSNs. How lipid composition impacts OR signalling is unclear. One report proposed a role in OR trafficking to cilia, based upon observations of reduced OR67d levels in the cilia of ATP8B mutant animals [33]. However, another study saw no defect in OR22a localization upon ATP8B knockout [34]. This discrepancy could reflect differences in the effect of ATP8B function on distinct ORs in different sensillar types, or the inherent difficulty in reliably quantifying protein levels in OSN cilia. It is also possible that lipid composition affects cilia morphology and/or the acute function of these ion channels, as in other biological contexts [35,36].

A significant impediment to relating ultrastructural features of sensilla to molecular components is the difficulty in using standard EM labelling methods. In other tissues, techniques combining EM and genetic labelling have facilitated the integration of morphological and molecular information [37–39]. For example, a diaminobenzadine (DAB)-oxidizing enzyme can be expressed in specific tissues or organelles, where its location is subsequently visualized by staining the oxidized DAB with EM-detectable electron-dense osmium tetraoxide (OsO4) [20,40–44]. However, the preservation of tissue ultrastructure during OsO4 staining necessitates chemical fixation or cryofixation [45]. Neither of these fixation methods have been easily applied to sensilla because the cuticle is impermeable to chemical fixatives, and cryofixation precludes labelling with DAB-oxidizing enzymes, severely curtailing its utility.

This challenge was recently addressed with the development of the ‘CryoChem' method, in which samples are rehydrated after cryofixation and high-pressure freezing [20]. This treatment preserves sensillar ultrastructure and creates a tissue environment amenable to fluorescent protein and APEX2 (a DAB-labelling protein) function, as well as en bloc heavy metal staining (figure 1c) [20]. CryoChem has been used in D. melanogaster to create three-dimensional reconstructions of several distinct, genetically marked OSNs in different sensilla by serial block-face scanning EM [20,46]. These observations provide useful insights into the relationship between OSN anatomy and olfactory physiology, as discussed below.

Together, these studies emphasize the wealth of cell biological detail that still remains to be discovered in sensilla. Even when proteins essential for sensilla formation are identified by genetic approaches, their mechanism of action can remain unclear [28,47,48]. Further progress requires both continued technical development to visualize the cuticular and membrane ultrastructure of sensilla and mechanistic and developmental characterization of protein function in both OSNs and support cells.

3. Non-receptor proteins in olfactory signalling pathways

Olfactory receptors are the central mediators of odour-evoked neuronal responses, often (but not always) sufficient to confer ligand-evoked membrane depolarization in heterologous cell types. However, olfactory signalling in a native context depends upon the interactions of odours with numerous other molecules, as well as the regulation of receptor function (figure 2).

Figure 2. Non-receptor proteins involved in olfactory signalling. Schematic depicting different classes of proteins that act with ORs in pheromone signal transduction (see text for details). The precise path(s) and molecular interactions of pheromone molecules within the sensillum remain unknown.

After entering the sensillum, odours must diffuse through the sensillar lymph, an aqueous ionic mixture rich in secreted proteins and proteoglycans [49]. The rate and efficiency with which odours are able to move into the sensillum and through the lymph to reach the OSN membrane is dependent on both the physico-chemical properties of the odours themselves [50,51] and their interactions with proteins in the lymph. Among the lymph proteins, odorant-binding proteins (OBPs) are the most well-studied, although their function remains enigmatic [52]. Members of this family of small, secreted proteins are expressed by sensillar support cells in defined, but often overlapping, sensillar types [53]. In vitro, OBPs can bind a multitude of odours with varying degrees of specificity and undergo conformational changes upon ligand binding [52,54]. A historical model for OBP function is that they associate with and transport hydrophobic ligands through the lymph fluid to the OSN here, they release the odour to the receptors, possibly triggered by local differences in pH near the cilia membranes or through hypothetical interactions of OBPs with cilia membrane proteins [54]. Data from studies on pheromone signalling systems in both D. melanogaster and moths is generally consistent with this model, but recent work with other OBPs is not [52], as we discuss below.

In D. melanogaster, the OBP LUSH (also known as OBP76a) is required for electrophysiological and behavioural responses to the pheromone cVA [55]. Supraphysiological concentrations of pheromone can evoke some neuronal activity [56], indicating that while LUSH may play a role in delivering cVA to the cognate receptor (OR67d), it is not an integral part of the signal transduction machinery. Recent in vivo work in moths has yielded similar results, with genetic knockdown or knockout of pheromone-binding OBPs resulting in 20–60% reductions in global pheromone-evoked antennal electrical activity [57–63] and similar decreases in behavioural responses [57,59,61,63]. The more modest phenotypes observed in moths relative to those in D. melanogaster may be due to methodological differences of these studies, but could also reflect functional redundancy between co-expressed moth OBPs [57].

In contrast with pheromone-interacting OBPs, analysis of family members expressed in other sensillar types in D. melanogaster has revealed subtler, and sometimes unexpected, roles. Loss of OBP28a in one basiconic sensillum class (ab8) resulted in increased physiological responses to odorants [53], suggesting a role in gain control of odour-evoked activity. However, OBP28a is expressed in several other sensillar classes (which, unlike ab8, express additional abundant OBPs), and responses of these sensilla to other odours were slightly diminished in Obp28a mutants [64]. Some OBPs are functionally redundant: simultaneous loss of the co-expressed OBP83a and OBP83b [53] led to delayed deactivation of neuronal responses after odour removal for a subset of OSNs importantly, this phenotype was rescued by re-expression of either individual protein [65]. In several cases, OBP function has remained elusive: comprehensive expression and mutational analysis of OBPs in six basiconic sensilla classes revealed that simultaneous loss of all proteins within a given sensillum had either no or very minor impact on the responses of OSNs to odour stimuli, which spanned diverse chemical classes, a wide concentration range and varied temporal dynamics [66]. It is possible that these proteins only function in particular biological contexts, as has been suggested for OBP69a, whose expression in pheromone-sensing sensilla is modulated by social interactions of flies [67].

Together, these results indicate that OBPs have diverse, odour-specific and neuron/receptor-specific roles, although the biochemical mechanisms remain unclear in any case. These proteins could be acting as a sink for some odorants, lowering background signals by preventing less ecologically pertinent chemicals from being able to reach receptors. Alternatively, they could clear ligands away after the initial stimulus to preserve the temporal connection between an encounter with an odour and neuronal activity. They could also contribute by binding endogenous lymph molecules: for example, OBP59a is expressed in apparently poreless sensilla in the antenna and is essential for hygrosensory behaviours in D. melanogaster, a sensory modality that may not depend upon binding of external molecules [68]. While connecting OBPs' capacity to bind ligands in vitro with their physiological and behavioural functions in vivo remains a substantial challenge, the appreciation that there may not be a universal function for this protein family may help researchers to maintain an open mind in future explorations.

Other soluble proteins in the sensillar lymph include chemosensory proteins (CSPs) [69,70], Niemann Pick-type C2 (NPC2) homologues [71–76] and odorant-degrading enzymes (ODEs) [77]. CSPs and NPC2 homologues bind myriad small compounds in vitro [70,71,73,75], but their in vivo functions are almost completely unknown. Some contributions may not necessarily be related to sensory detection: recent work in the malaria mosquito (Anopheles gambiae) demonstrates that genetic variants of the leg-enriched CSP SAP2 confer insecticide resistance [78]. These observations suggest that this CSP acts in sequestration/detoxification of environmental chemicals that enter the body through chemosensory sensilla on these appendages.

ODEs are thought to degrade odour molecules in the sensillar lymph [77], which could reduce background neuronal activity and/or regulate odour-evoked temporal dynamics. The first reported ODEs were members of an antennal-specific esterase family [77,79]. Work over the last two decades has discovered other classes of putative ODEs, including some membrane-bound Cytochrome P450s [80–84]. The best-studied ODEs are D. melanogaster Esterase 6 and juvenile hormone esterase duplication [85], which evolved from an ancestral juvenile hormone esterase orthologue [86]. These enzymes do not degrade juvenile hormone, but rather break down volatile esters [85,87–89]. Although the exact contributions of these and other ODEs to odour-evoked neuronal responses and behaviour remain unclear [87–90], the loss of the juvenile hormone esterase duplication appears to cause modest decreases in olfactory and behavioural responses to fruit esters [89].

In addition to molecules secreted by support cells, proteins in these cells' membranes may contribute to olfactory signalling. In D. melanogaster, the ammonium transporter Amt is thought to be expressed exclusively in the support cells of a coeloconic sensillum class that houses an ammonia-sensing neuron [48]. Genetic analysis revealed that loss of Amt leads to greatly diminished responses to ammonia stimulation [48]. The A. gambiae Amt orthologue functions as an ammonia transporter in vitro [91], but it is unclear exactly how this activity might contribute to olfactory detection in vivo. One hypothesis is that Amt transports ammonia out of the lymph to lower the basal concentration of this chemical near the OSN dendrites, thereby helping to minimize tonic adaptation of the ammonia-sensing neuron [48]. However, a recent analysis of A. gambiae Amt using transgenic tools indicated that this gene is expressed in both support cells and OSNs [92], raising questions about its cellular site(s) of action. Regardless of the precise mechanism, Amt represents an interesting case where an integral membrane transporter can directly affect olfactory detection. Many other uncharacterized putative transporter proteins are expressed in the antenna [48], and it will be interesting to determine whether any of these have analogous roles.

Non-receptor proteins in the OSN cilia membrane can also play important roles in olfactory transduction. The best characterized is Sensory Neuron Membrane Protein 1 (SNMP1), a two-pass transmembrane protein related to the mammalian CD36 family [93]. SNMP1 was originally characterized in moths for its expression and ciliary localization in pheromone-sensing neurons [94], properties that are broadly conserved in insects [95–98]. Mutational analysis in D. melanogaster demonstrated that SNMP1 is essential for OR67d-mediated responses to cVA [95,99]. However, the requirement for SNMP1—like that of the OBP LUSH—can be bypassed by very high concentrations of this pheromone [100], indicating it is not a strictly essential part of the receptor complex. The important role of SNMP1 in pheromone detection in D. melanogaster is likely to be conserved in other insects. For example, RNAi of Snmp1 in B. mori impairs the ability of males to locate and mate with females, processes that depend heavily upon pheromone detection [98]. Mammalian CD36 proteins bind/transport lipid-like molecules in diverse cellular contexts, and the partial ability of a murine CD36 homologue to rescue Snmp1 mutants in D. melanogaster [97] has guided mechanistic studies of SNMP1 function. A homology model of SNMP1, based on the crystal structure of the CD36 protein LIMP-2, predicts that the SNMP1 ectodomain has a hydrophobic cavity [97], which may act as a conduit for transporting hydrophobic pheromone molecules from the extracellular lymph to a closely apposed OR complex in the cilia membrane [95,97,101,102]. While the mechanism remains to be fully established, the critical requirement for OBPs and SNMP1s in the detection of pheromones, but not other types of odours, may be related to the biochemical challenges of concentrating generally large and highly hydrophobic pheromone ligands at the surface of the OSN membranes.

Olfactory neuron responses can be further modulated by other signalling molecules after receptor activation. A longstanding question is how insect ionotropic olfactory receptors attain sufficient sensitivity without signal amplification by second messengers, which is inherent to vertebrate metabotropic chemosensory receptor transduction [103]. Recent work offers a solution to this problem by providing evidence that the degenerin/epithelial sodium channel Pickpocket 25 (PPK25) amplifies ligand-evoked currents downstream of certain olfactory receptors [104]. In D. melanogaster, the genetic knockout or overexpression of PPK25 decreases or increases, respectively, the physiological sensitivity of Or47b OSNs [104], a class of pheromone-sensing neurons involved in courtship behaviour [105,106]. These effects are dependent on calmodulin, as a mutation of a calmodulin-binding motif in PPK25 or pharmacological inhibition of calmodulin mimics loss of PPK25 [104]. Interestingly, this role of PPK25 can also be observed for OSNs expressing IR84a, which recognizes food-derived odours that promote courtship behaviour [107], and for a population of gustatory sensory neurons (GSNs) that detects non-volatile pheromones [104]. The revelation that this PPK acts as a signal amplifier, rather than a sensory receptor, in these different classes of neurons has potentially broad significance: many members of the D. melanogaster PPK family have been implicated in diverse sensory modalities but it has been unclear (with one exception [108,109]) whether they are the sensory receptors or not [110–124]. These channels may have analogous roles beyond sensory systems for example, PPK11 and PPK16 modulate presynaptic membrane voltage at the neuromuscular junction to regulate homeostatic plasticity [125].

Some olfactory receptor subunits may also act as modulators of receptor activity, rather than binding ligands themselves. For example, several IR-expressing OSNs express—in addition to a tuning receptor and co-receptor—a third receptor protein, IR76b [13]. Some evidence points to IR76b acting as a critical component of a putative tripartite olfactory receptor complex [17,126], which is also concordant with the broad expression and function of this protein in various GSN populations [126–132]. However, a distinct role for IR76b in limiting, rather than contributing to, ligand-evoked responses has emerged through analysis of a population of GSNs that detect both sugars and acetic acid. Here, the mutation of IR76b leads to increased ligand-evoked physiological responses, with a corresponding enhancement of behavioural sensitivity [133]. The function of IR76b as a dampener of neuronal responses exhibits some specificity, as the sensitivity of a mammalian capsaicin receptor that is ectopically expressed in these sugar/acid-sensing neurons is not affected in Ir76b mutants. Moreover, ectopic IR76b expression in other neuronal populations does not reduce their physiological responsiveness [133]. The context-dependent modulatory role of IR76b is reminiscent of mosquito carbon dioxide receptors, which comprise two subunits that are essential for ligand-evoked responses, and a third that may modulate ligand-evoked sensitivity [134–136]. The mechanistic basis of receptor subunit modulation is unknown in any case, but these findings highlight the intricate regulation that may occur between subunits within (putative) heteromeric complexes to shape ligand-dependent ion conduction.

4. Biophysical properties and intercellular regulation of olfactory sensory neuron responses

OSN signalling consists of two physiological processes: first, odour-dependent gating of the olfactory receptor channel, ion flow and cilia membrane depolarization, and second, conversion and propagation of this initial signal by voltage-gated channels in the form of action potentials (or spikes) down the OSN axon [137–139] (figure 3). The first of these processes can be detected as changes in a sensillum's local field potential (LFP), representing the transient electrical potentials in the sensillum generated by OSNs, as well as contributions from the ion transport activities of support cells [138,139]. LFP dynamics reflect signal transduction properties that are determined by the specific nature of odour ligand/receptor interactions, while the temporal dynamics of spiking can be described by a linear filter that is stereotyped across different OSN classes [138,140].

Figure 3. Peripheral olfactory physiological processes. (a) An idealized drawing of a sensillar olfactory response, illustrating the two main physiological processes. (b) Schematic of a sensillum depicting regions where these physiological responses occur. Although spikes are thought to be generated in the OSN axons, they can be detected experimentally in the dendrites in the sensillum shaft, possibly through backpropagation.

Most olfactory physiological studies do not measure LFP and use spike frequency as the sole proxy for reporting odour-evoked neuronal activity [138,141,142]. While spikes represent the information that is transmitted to the brain, a comprehensive appreciation of peripheral OSN physiology is crucial to understand responses to naturalistic odour stimuli. Odours exist as plumes comprising pockets of air containing wide-ranging concentrations of chemicals. OSNs respond to this temporally complex stimulus pattern in diverse ways, such as desensitization to strong stimuli, or sensitization to repeated weak stimuli [137,138,140,143,144]. LFP and spike rate exhibit very different adaptation kinetics [138,140] and also appear to adapt in response to different aspects of the odour stimulus. For example, LFP, but not spike rate, adapts strongly in response to changes in the mean stimulus intensity [140]. By contrast, both LFP and spike rate are influenced by the variance in an odour stimulus, although the adaptation dynamics of each component differs [140]. LFP and neuron spiking are, of course, intimately connected phenomena, and while the kinetics of spike rate and LFP are distinct, the dynamics of changes in spike amplitude are nearly identical to those of LFP [145]. Together, these analyses reveal the sophistication with which OSNs encode different aspects of odour stimuli and emphasize that measurement of spike frequency alone does not fully capture OSN responsiveness and therefore our ability to understand how odour-evoked neuron activity arises.

The molecular basis of the dynamic physiological properties of OSNs remains unclear. Most analyses have focused on structure/activity dissection of ORCO, providing evidence that sensory adaptation relies upon modulation of both receptor localization and sensitivity. This co-receptor (and, presumably, its partner tuning OR) were observed to be depleted from cilia upon prolonged odour exposure, although this was measured only over a multi-day time scale [146]. Activity-dependent control of ORCO localization may rely on calmodulin: RNAi of calmodulin or mutation of a predicted calmodulin-binding motif in ORCO's second intracellular loop disrupts its cilia localization, with consequent defects in odour-evoked activity [146]. Physiological studies have provided additional evidence for the role of calcium signalling and/or calmodulin in ORCO-dependent sensitization of neurons to repeated odour stimulation [147] and sensory adaptation of OR-expressing neurons [148]. The same loop of ORCO also contains three potential phosphorylation sites [144,149,150]. Mutation of these sites reduces ORCO's conduction properties in heterologous cells [149] and diminishes OSN sensitivity and behavioural responses to odours in vivo [150]. In addition, mutation of ORCO's phosphorylation sites prevents odour sensitization [143]. One of these sites, S289, is dephosphorylated in vivo upon OSN desensitization [144,151]. An ORCO S289A mutation reduces OSN sensitivity in vivo, and a phospho-mimetic mutant (ORCO S289D ) reduces the magnitude of OSN desensitization after odour exposure [144]. These studies begin to unveil the complexity of olfactory receptor regulation that contributes to the temporal response properties of OSNs, but also make apparent the challenge of cleanly dissecting effects on receptor localization and/or activity.

The molecular regulation of other types of olfactory receptors is essentially unknown, although N-glycosylation has been implicated in the control of IR localization and activity [152]. Electrophysiological studies indicate that Or and Ir neurons have distinct temporal response properties [148,153], at least some of which appear to be dependent on the receptors themselves [153]. Moreover, the acute contribution of pathways implicated in sensillar development, such as Hedgehog signalling or the lipid flippase ATP8B (discussed above), remains to be explored.

Beyond autonomous regulatory mechanisms in OSNs, recent work has characterized the interdependence of the activity of different OSNs within the same sensillum. In many sensillar classes, the activity of one OSN is inhibited upon activation of a neighbouring neuron [154]. Blocking synaptic transmission does not prevent such inhibitory interactions between the two OSNs [154], nor is there evidence for gap junctions between paired OSNs. These observations suggest that the inhibition occurs through ephaptic coupling [46,154], a phenomenon in which the activity of one neuron alters the local electric field to impair depolarization of a nearby neuron. In support of this hypothesis, simultaneous recording of two different sensilla that are artificially coupled by a metal electrode demonstrated that continuous stimulation of a neuron in one sensillum can be inhibited by excitation in the adjacent sensillum [46]. Moreover, the combination of these observations with EM analysis of defined neuron types via CryoChem (described above) revealed that the inhibitory effect is stronger when exerted from a larger OSN onto a smaller OSN [46]. A plausible explanation for this asymmetric relationship is that bigger neurons are expected to have lower input resistance and a greater dendritic surface area to allow for a higher maximal LFP [46], hinting at a previously unappreciated link between OSN morphology and physiology. Future work will determine how such ephaptic interactions impact odour coding, in particular of complex natural odour blends.

Work on GSNs in the bumblebee (Bombus terrestris) provides interesting additional insights into how, and why, neurons within the same sensillum communicate [155]. Recordings from a highly sensitive sugar sensing neuron in ‘type A' sensilla on the mouthparts revealed an unusual bursting pattern of spikes upon stimulation with high concentrations of sucrose [155]. The end of the spike burst coincides with a single spike from a second neuron in this sensillum. Introduction of a gap junction inhibitor into the sensillum led to the continuous sucrose-evoked firing of the first neuron, suggesting that—in contrast with the ephaptic inhibition described in olfactory sensilla—the second neuron terminates the first neuron's spike train via electrical synapses (these structures have not, however, been visualized directly). Importantly, this bursting pattern of firing prevents neuronal desensitization, which may explain the ability of bees to sustain feeding behaviour on high-sugar nectar [155].

5. Conclusion and perspectives

The discovery of insect olfactory receptors has been instrumental in understanding how these animals detect environmental odours, as well as facilitating the development of molecular tools to map and manipulate olfactory circuits. However, receptors alone do not define the exquisite sensitivity, specificity and temporal precision observed in odour-evoked neuronal activity. We have highlighted the complexity of peripheral signal transduction in olfactory sensilla, and the extraordinary wealth of biology that remains to be uncovered. It is clear that many neuronal, non-neuronal and secreted molecules that participate in this process (or rather processes) have still to be characterized [156]. Moreover, determination of the in vivo function of most proteins in defining signalling properties, and how these impact behavioural responses, will require technical innovations to permit their acute inhibition to distinguish roles in sensillar development from direct contributions to signal transduction. Finally, while investigating insect olfactory transduction is of widespread interest in sensory neuroscience and chemical ecology, many of the insights gained are likely to have broad relevance for understanding molecular and cellular communication processes across diverse tissues and species.


Summary of Karyotype Data Across Orders of Insects

Sex in most insects is determined genetically, and Table 2 provides an overview of the phylogenetic distribution of sex determination mechanisms across insects. Below we give a short description of karyotype and sex chromosome composition within insect orders. We also include a short discussion on asexual species in the various groups, taken from the compilation of ( Normark 2003 Normark and Ross 2014). For completeness, we include the limited data available for Entognatha (Collembola, Diplura, and Protura) that are wingless arthropods, which, together with insects, make up the subphylum Hexapoda.

Sex determination systems across insects

. XO . XY . C XO a . C XY b . ZO . ZW . C ZW c . Hom d . HD/PGE e . Parth f . CN g . Taxa .
Orthoptera 223 49 9 10 291
Notoptera 2 2
Phasmatodea 69 14 37 25 144
Embiidina 8 2 10
Blattodea 108 2 3 113
Isoptera 1 2 61 62 18 83
Mantodea 60 1 40 2 4 107
Plecoptera 3 1 8 4 16
Zoraptera 1 1
Dermaptera 3 22 27 2 54
Trichoptera 15 6 23 44
Lepidoptera 10 18 12 16 1163 1219
Mecoptera 13 1 1 15
Siphonaptera 2 4 6
Diptera 48 1893 10 7 93 46 97 1456
Raphidioptera 6 6
Megaloptera 4 4
Neuroptera 2 70 2 74
Strepsiptera 1 1 1 3
Coleoptera 770 3198 12 207 10 326 484 4934
Hymenoptera 1591 158 1749
Phthiraptera 1 1 4 16 22
Psocoptera 91 2 39 1 133
Hemiptera 155 284 1 3 255 467 114 1313
Thysanoptera 24 59 83
Odonata 403 20 1 11 432
Ephemeroptera 2 6 11 19
Zygentoma 3 2 1 6
Archaeognatha 5 2 7
Diplura
Collembola 17 10 21 53 101
Protura 3 1 2 5
Totals 1979 5582 21 361 25 25 12 159 1891 1215 2025 12452
. XO . XY . C XO a . C XY b . ZO . ZW . C ZW c . Hom d . HD/PGE e . Parth f . CN g . Taxa .
Orthoptera 223 49 9 10 291
Notoptera 2 2
Phasmatodea 69 14 37 25 144
Embiidina 8 2 10
Blattodea 108 2 3 113
Isoptera 1 2 61 62 18 83
Mantodea 60 1 40 2 4 107
Plecoptera 3 1 8 4 16
Zoraptera 1 1
Dermaptera 3 22 27 2 54
Trichoptera 15 6 23 44
Lepidoptera 10 18 12 16 1163 1219
Mecoptera 13 1 1 15
Siphonaptera 2 4 6
Diptera 48 1893 10 7 93 46 97 1456
Raphidioptera 6 6
Megaloptera 4 4
Neuroptera 2 70 2 74
Strepsiptera 1 1 1 3
Coleoptera 770 3198 12 207 10 326 484 4934
Hymenoptera 1591 158 1749
Phthiraptera 1 1 4 16 22
Psocoptera 91 2 39 1 133
Hemiptera 155 284 1 3 255 467 114 1313
Thysanoptera 24 59 83
Odonata 403 20 1 11 432
Ephemeroptera 2 6 11 19
Zygentoma 3 2 1 6
Archaeognatha 5 2 7
Diplura
Collembola 17 10 21 53 101
Protura 3 1 2 5
Totals 1979 5582 21 361 25 25 12 159 1891 1215 2025 12452

The number of taxa reported for each type of sex determination system, the number of asexual species, and the number of taxa for which only chromosome number is available is indicated for each order of insects. The order of taxa matches the phylogeny in Figure 3.


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