Do Insects Have Fat?

Insects have a fat body. It’s a versatile organ, a sort of combination of adipose tissue (the blubber that humans have) and liver. Its principal roles are:

• As with adipose tissue, it is the main nutrient storage site (Telfer & Kunkel, 1991);
• Storage of neutralised waste metabolites;
• Detoxification of ammonia from protein metabolism (Scaraffia et al., 2010);
• Control of nutrient levels in the haemolymph (“bloodstream”) (Colombani et al., 2003);
• An immunity organ, producing antimicrobial peptides in response to bacterial or fungal intrusion into the haemolymph (Lemaitre & Hoffmann, 2007);
• In females, producing vitellogenin, the precursor to egg yolk and thus critical for reproduction (Roy & Raikhel, 2011).

The fat body is composed mainly of lipid-, protein-, and glycogen-rich cells called trophocytes. As the individual insect goes through more life stages, the fat body grows as the trophocytes become larger and develop vacuoles in which the fat, glycogen, and proteins are kept to serve as nutrient stores, the energy from which is used mostly for reproduction and locomotion (Arrese & Soulages, 2010). This is the closest thing to body fat that you will have in insects.

Another important cell type found in the fat body is the oenocyte, although it can also be found in association with the epidermis. Oenocytes are hydrocarbon factories, play a role in maintaining homeostasis, and in detoxification (Martins et al., 2011).

The fat body is found beneath the epidermis (like adipose tissue) and, in some insects, around the alimentary canal and testes (Roma et al., 2010). The trophocytes are linked by a thin lamina that forms lobes and ribbons into the haemocoel, increasing the fat body’s surface area and thus allowing more nutrient exchange between the fat body and haemolymph (Arrese & Soulages, 2010).

The differentiation of the fat body begins fairly early in development during the embryonic stage. In all insects, the trophocytes grow as more food is ingested. In holometabolous insects, those that undergo a metamorphosis, the larva is specifically a feeding stage during which it must accumulate as many energy stores as possible to be able to metamorphose (Aguila et al., 2007). These stores are released during the prepupal stage to be used up during pupation, when metamorphosis takes place. During metamorphosis, the highly-organised larval fat body is restructured into a loose mass of connected trophocytes (Nelliot et al., 2006). The larval trophocytes become disconnected and form clumps in the pupal body. These trophocytes then undergo programmed cell death in the new adult, and are replaced in the adult by new trophocytes derived from the body wall (Hoshizaki et al., 1995). The larval trophocytes do remain in the adult as energy reserves until a feeding site is found (Aguila et al., 2007).

References:

Aguila JR, Suszko J, Gibbs AG & Hoshizaki DK. 2007. The role of larval fat cells in adult Drosophila melanogaster. The Journal of Experimental Biology 210, 956-963.

Arrese EL & Soulages JL. 2010. Insect Fat Body: Energy, Metabolism, and Regulation. Annual Review of Entomology 55, 207-225.

Colombani J, Raisin S, Pantalacci S, Radimerski T, Mantagne J & Léopold P. 2003. A Nutrient Sensor Mechanism Controls Drosophila Growth. Cell 114, 739-749.

Hoshizaki DK, Lunz R, Johnson W & Ghosh M. 1995. Identification of fat-cell enhancer activity in Drosophila melanogaster using P-element enhancer traps. Genome 38, 497-506.

Lemaitre B & Hoffmann J. 2007. The Host Defense of Drosophila melanogaster. Annual Review of Immunology 25, 697-743.

Martins GF, Ramalho-Ortigão JM, Lobo NF, Severson DW, McDowell MA & Pimenta PFP. 2011. Insights into the transcriptome of oenocytes from Aedes aegypti pupae. Memórias do Instituto Oswaldo Cruz 106, 308-315.

Nelliot A, Bond N & Hoshizaki DK. 2006. Fat-body remodeling in Drosophila melanogaster. Genesis 44, 396-400.

Roma GC, Bueno OC & Camargo Mathias MI. 2010. Morpho-physiological analysis of the insect fat body: A review. Micron 41, 395-401.

Roy SG & Raikhel AS. 2011. The small GTPase Rheb is a key component linking amino acid signaling and TOR in the nutritional pathway that controls mosquito egg development. Insect Biochemistry and Molecular Biology 41, 62-69.

Scaraffia PY, Zhang Q, Thorson K, Wysocki VH & Miesfeld RL. 2010. Differential ammonia metabolism in Aedes aegypti fat body and midgut tissues. Journal of Insect Physiology 56, 1040-1049.

Telfer WH & Kunkel JG. 1991. The Function and Evolution of Insect Storage Hexamers. Annual Review of Entomology 36, 205-228.

Daily Factoid: Some insects parasitise plants

In a brilliant bit of parasitism, galling insects drill into plants and affect their metabolism and development in such a way that galls form. The exact molecular mechanisms for this are still largely unknown, but what is known is that these galls are highly-advantageous for the insects. They can take various forms, from being mere modifications of the plant’s nutrient production (inducing greater protein and sugar production for the insect to feed on), to being local growths of nutrient-rich plant tissues on which the insect can munch, to full-blown houses for the insect to live in.

Other insects are leaf-miners, and some of these insects also have the ability to modify the plant’s development. Next autumn, observe the trees with yellow leaves about to drop off. Some of them might have green “islands” on them – areas that are still photosynthetically active even though the rest of the leaf is ageing and dying.

Biochemical analyses of both leaf mines and galls show higher levels of cytokinins, plant hormones that inhibit ageing, maintain chlorophyll, and enhance nutrient release. In other words, the green mined areas are much more nutritious than the rest of the leaf. Like galls, they’re induced by the leaf-mining insects to get an easy source of food – and at a critical time of the year, right before winter.

What’s as interesting as this modification of plant structure by parasitic insects (which is, in principle, similar to behavioural modifications by animal parasites), many studies suggest that the involved cytokinins are not of plant origin, but injected directly by the insects. Going even deeper, the cytokinins injected by the insects aren’t produced by the insects, but by bacterial symbionts. Draw that on a flowchart to see how cool this is: it’s a triple-layered interaction that’s been most likely moulded by millions of years of coevolution.

Further Reading:

General:

The evolution and adaptive significance of the leaf-mining habit

Geometrical Games between a Host and a Parasitoid

Cytokinins and insect galls

Galls:

Manipulation of the phenolic chemistry of willows by gall-inducing sawflies

Mines:

Are green islands red herrings? Significance of green islands in plant interactions with pathogens and pests

Cytokinins:

Extracellular Invertase Is an Essential Component of Cytokinin-Mediated Delay of Senescence

Pathological hormone imbalances

My Research: Carabid networks in a variable landscape

In this study, I will be setting up sets of pitfall traps (preservative-filled cups in the ground that ground insects fall into) in ecosystems with very different habitat types, e.g. a forest with clustered tree species and shrub types, clearings, human structures, ponds.

This will not only give me a good estimate of the ground insect diversity, it will also allow me to analyse landscape patterns and networks. In other words, I will be studying how insects move around the landscape. Do some species become isolated in certain habitats? What habitats serve as corridors through which species can move? Are there any habitat combinations that foster a particularly active or biodiverse community?

Such a study provides important data for environmental management. It’s fallacious and oversimplistic to view a forest as a single habitat, because it’s an ecosystem comprising of many habitats that vary at a small spatial scale. An environmental manager absolutely needs to take this into account before approving any changes. Data accumulated from such studies allow a limited, but crucial, amount of prediction to be made for the effects of human modification, from logging to reforestation to construction of artificial lakes. The same is true for agricultural managers and farmlands: do hedgerows serve as habitats and corridors in otherwise inhospitable arable land? How does intercropping affect insect movements? What is the best solution to keep pollinator biodiversity up while reducing pest numbers?

My analysis will be focused on carabid beetles, most probably in a forest landscape (because I don’t want to get shot by a farmer while doing fieldwork), but other groups and ecosystems can be examined in due time, or by interested parties or universities/schools.

Top Research of 2012: Arthropods

Jump to: Botany; Developmental Biology; Ecology; Environmental; Evolution; Geology; Historical Geology; Human Evolution; Palaeontology; Zoology.

Now that we’re done with the top books of the year, let’s look at the top research of the year. I re-examined a total of 412 papers published this year, sorted in the following categories: Arthropods; Botany; Developmental Biology; Ecology; Environmental; Evolution; Geology; Historical Geology; Human Evolution; Palaeontology; and Zoology. As with the books, every day, I will do a top 10 research for each category. The top 10s will be inverted like a proper countdown. As with any top 10 lists, your mileage may vary; these picks and the rankings are all subjective and prone to my own biases.

Let’s start off with the arthropods. The top 10 papers were chosen from a master list of 74 papers. [OA] indicates open access papers. The topic listing, from 10 to 1: spider intelligence; spider silk; fossil insect behaviour; fossil pupation chambers; caste-specific neuroanatomy; early arthropod evolution; evolutionary dynamics; early fossil insect; earliest amber arthropods; treehopper helmet.

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10. The discerning predator: decision rules underlying prey classification by a mosquito-eating jumping spider.

Jumping spiders’ excellent eyesight has led to their also having high intelligence, being able to observe and filter what they see to the point that the African jumping spider Evarcha culicivora can differentiate their prey, female Anopheles mosquitoes, from all other insects flying around just by looking at their antennae. This is what Nelson & Jackson showed with this elegant experiment.

By combining parts from male and female mosquitoes and using the resultant Frankenmosquitoes as lures for the spiders to attack, they identified the two clues that led to the most attacks: a red, blood-engorged abdomen, and slender antennae. Both of these are female mosquito features: male mosquitoes don’t feed on blood (they’re nectar feeders), and males have bushy antennae. As for the specificity for Anopheles mosquitoes, that’s explained by their posture – other mosquitoes rest with their body parallel to the ground, while Anopheles rest with a 45° angle.

For showing that such a tiny spider is capable of such complex prey-distinction and thus giving even more credence to the notion that intelligence is not a function of brain size, as well as for having a great experimental design, Nelson & Jackson get the #10 place.

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9. Post-secretion processing influences spider silk performance.

Spider silk is not a simple strand that’s the same in every species. There’s many different types of silk that come out of different glands, and the silk is also modified after it’s secreted. The study focuses on major ampullate silk, the type of silk that makes up the framework of an orb web and whose stiffness is responsible for the strength of the webs. The researchers examined natural silk, and silk that they supercontracted to remove any post-secretion modifications. What they found was that these supercontracted silks lost the stiff properties of their natural counterparts, meaning that their properties come from whatever modification is made to them, not from the actual structure and composition of the silk. I find this discovery significant because it adds a new dimension to the study of spider silk, a field that has quite a lot of technological and biomimetic research ahead of it.

Other significant spider silk and web-related research this year include:

The role of capture spiral silk properties in the diversification of orb webs: how various silk types affect the web’s properties.

Nonlinear material behaviour of spider silk yields robust webs: This research provides more insight into the factors mentioned above, finding that it isn’t just the type of and modification of silks that affect the web’s properties, but that the geometry of the web is as important in determining its strength and behaviour.

Early Events in the Evolution of Spider Silk Genes [OA]: A phylogeny of genes from the silk-producing glands reveals gene duplications associated with more diverse ecological use of silk and webs.

Functional values of stabilimenta in a wasp spider, Argiope bruennichi: support for the prey-attraction hypothesis: research into the use of stabilimenta, UV-reflective strands of silk that orb-weavers have.

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8. Jurassic mimicry between a hangingfly and a ginkgo from China. [OA]

This is cool more than anything else, and the mimicry is shown in the picture above, from the original paper: A, B, E, and F are gingko leaves; C and H are the described specimen and its wing, D and I are a closely-related species which also exhibits mimesis with gingkos; J and K are gingko leaf closeups. As you can see, the similarities are striking, and the artist’s conception in G shows how well the hangingfly would have blended in. The paper has more details on the coevolution of mimesis between this group of hangingflies and gingkos. In all, a neat piece of work with evolutionary insights as well as cool fossil preservation.

Another paper this year has preserved evidence of insect behaviour: Wing stridulation in a Jurassic katydid (Insecta, Orthoptera) produced low-pitched musical calls to attract females. The mating call of a katydid has been reconstructed based on the preservation of its stridulatory apparatus, a hard file that the wings strike against to make the music. Related, this paper from this year shows how sensitive the hearing of katydids is: Auditory change detection by a single neuron in an insect.

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7. The Earliest Evidence of Holometabolan Insect Pupation in Conifer Wood. [OA]

This paper describes U-shaped burrows in 210 Ma wood from Utah, USA. These were previously assumed to be bee or wasp borings, but the detailed analysis presented in the paper shows that these borings are actually pupation chambers made by a small organism that ate its way into the wood, then emerged from the other side, as presented in the diagram above. From the size of the borings, the authors propose that the organism is a cupedid beetle, showing that these beetles were dominant before the other beetles radiated later.

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6. Division of Labor in the Hyperdiverse Ant Genus Pheidole Is Associated with Distinct Subcaste- and Age-Related Patterns of Worker Brain Organization. [OA]

That different castes will have a different brain organisation is expected and has been shown in many papers (e.g. 1, 2, 3). This paper is significant because it’s so thorough: it examines castes of three species of Pheidole ants and their brain anatomy. The diagram above summarises the pattern observed: the colours are species, the shapes are castes; the axes are two variables that together make up 87% of the brain variation seen. The pattern is clear: neuroanatomy is determined by caste, not by species. This is just underlines the incredible amount of plasticity in ants (and other eusocial insects), where the environment can dictate how an individual will function and develop to allow the colony to adapt to changing needs and conditions.

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5. A Carboniferous Non-Onychophoran Lobopodian Reveals Long-Term Survival of a Cambrian Morphotype.

This paper has equal relevance to palaeontology as it does to arthropods: like several other papers of the past few years (e.g.), it reinforces the idea that the Cambrian freaks didn’t go extinct, but that the nature of the fossil record changes since the Cambrian to make their preservation much rare (the advent of burrowing made it much harder for such soft-bodied forms to be preserved). This one is the most stirking example yet: a long-legged lobopod, 200 million years after the Cambrian (it comes from the famous Mazon Creek locality in Illinois, USA, 296 Ma)! Lobopods are a wastebasket taxon in which soft-bodied arthropods with stubby legs are dumped, including many fossil-only taxa, tardigrades, and onychophorans. There are two groups: short-legged forms (includes the last two) and long-legged ones, up until this paper known only from the Cambrian.

It was a good year for arthropod evolution, with many excellent studies into the biology and diversity of early arthropods:

Exceptionally preserved crustaceans from western Canada reveal a cryptic Cambrian radiation: These Canadian fossils bring the earliest fossil records of branchiopods, copepods, and ostracods back to the mid-Cambrian.

Silurian horseshoe crab illuminates the evolution of arthropod limbs: A horseshoe crab from Herefordshire, showing a very exciting biramous limb, the significance of which would need an entire post to explain.

A Silurian myodocope with preserved soft-parts: cautioning the interpretation of the shell-based ostracod record is another Herefordshire find that finds that ostracod shells, very abundant fossils with significant stratigraphic and other practical use, are not quite as informative taxonomically as previously thought.

Cambrian lobopodians and extant onychophorans provide new insights into early cephalization in Panarthropoda [OA]: A complete redescription of Onychodictyon‘s head, showing that the arthropod mouth may have originated multiple times.

Cambrian bivalved arthropod reveals origin of arthrodization: A new Burgess Shale arthropod suggests that the key feature of arthropods, the exoskeletal segmentation, was a feature that evolved for swimming.

Morphology of Cambrian lobopodian eyes from the Chengjiang Lagerstätte and their evolutionary significance shows that Cambrian lobopods had pretty sophisticated eyesight.

Complex brain and optic lobes in an early Cambrian arthropod: Eyes are nice and all, but how about preserved brains and nervous tissue?

Internal Soft-Tissue Anatomy of Cambrian ‘Orsten’ Arthropods as Revealed by Synchrotron X-Ray Tomographic Microscopy [OA] shows more spectacular internal details of long-extinct arthropods.

Exceptionally Preserved Cambrian Trilobite Digestive System Revealed in 3D by Synchrotron-Radiation X-Ray Tomographic Microscopy [OA]: As above.

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4. Loss of flight promotes beetle diversification. [OA]

In this study, a molecular phylogeny of Japanese carrion beetles was done, and the result found was that flight loss promotes speciation. Flightless populations have more genetic differences between themselves than do flight-enabled populations. This is to be expected: flight enables greater geographic dispersal, allowing distant populations to reproduce and keep gene flow between them; with flight loss, this doesn’t happen, resulting in more isolation and thus more speciation, as shown in the above bar chart. The authors went further and did a short meta-analysis for other beetle groups and found a similar effect. I look forward to deeper studies examining the precise interplay between diversification and flight loss – does flight loss really directly cause speciation, or is it an indirect knock-on effect. Loss of flight must be related to other factors such as habitat requirements, life history, or feeding preferences, as the authors note; maybe it’s those other factors that actually promote the speciation. The authors checked for this in their carrion beetle dataset, but it’s worth looking into with other taxa.

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3. A complete insect from the Late Devonian period.

The fossil doesn’t quite look like an insect until you examine it closely – when I first saw the picture, I thought it was some notostracan. But then it becomes clear that there’s a pair of antennae, then you see the head, then the rest of the body – this is an insect. It’s not the oldest – that honour remains with Rhyniognatha hirsti – but it does come from a time when the fossil record of insects is completely lacking, the Late Devonian, and it’s by far the earliest complete insect – this really is a landmark find.

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2. Arthropods in amber from the Triassic Period.

This is not the oldest amber (that’s from the Carboniferous), but it is the oldest fossiliferous amber. Microorganisms have been reported from it before, but this paper records the oldest arthropod inclusions in amber, beating the previous records from the Middle East by some 100 million years. The arthropods are one fly and two mites (one of which is pictured above), with more still to come in future papers.

Some more cool insect preservation papers published this year include:

The original colours of fossil beetles details how preservation of beetle cuticle allows us to reconstruct the colour of fossil beetles – after all, the metallic sheen that some beetles have isn’t due to pigments, but due to the nanostructure of the cuticle playing tricks with the light. THE CONTROLS ON THE PRESERVATION OF STRUCTURAL COLOR IN FOSSIL INSECTS outlines the details of how cuticle preservation affects recovered colour.

WIDESPREAD PYRITIZATION OF INSECTS IN THE EARLY CRETACEOUS JEHOL BIOTA shows that the insects from Jehol – the famous lacustrine fossil locality that has yielded many feathered dinosaurs – are pyritised with the help of bacterial acitvity.

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1. On Dorsal Prothoracic Appendages in Treehoppers (Hemiptera: Membracidae) and the Nature of Morphological Evidence. [OA]

In 2011, Prud’homme et al. published an intriguing paper with developmental and some morphological evidence that the helmet of treehoppers, pictured above, is actually a cooption of an ancestral wing-like structure. This is obviously a very extraordinary claim, and this paper reviews all the evidence and comes up with alternative scenarios that show flaws in the Prud’homme et al. paper. It gets the top spot not only for the subject matter, but also for being a prime example of the scientific method in action.

A morphology-only critique of the Prud’homme et al. paper was done very early in the year by Yoshizawa [OA].

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Jump to: Botany; Developmental Biology; Ecology; Environmental; Evolution; Geology; Historical Geology; Human Evolution; Palaeontology; Zoology.

Top Books of 2012: Zoology

Jump to another list: Environmental and Climate Change; Evolution; Historical Geology; History of Science; Human Evolution and Anthropology; Palaeontology.

These are books about animals. Some are layman-oriented, others are complete academic texts, so your mileage may vary. And with this post, the Top Books of 2012 comes at an end… but the series continues tomorrow with the real meat: the top discoveries of 2012, outlined in a total of 11 posts.

Cardé & Resh (eds.). A World of Insects: The Harvard University Press Reader. (Harvard University Press)
	This is an excellent anthology of some of the groundbreaking studies in entomology. It’s compiled not for the entomologist, but for the layman, so it’s also very accessible. If you’re at all interested in insects, or have children who are into nature, or are a science/biology teacher, this books is perfect for you.

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Hughes, Brodeur & Thomas (eds.). Host Manipulation by Parasites. (Oxford University Press)

Who doesn’t think behaviour-manipulating parasites are awesome? No reader of this blog, I can guarantee that based on readership statistics of this post. This book is an authoritative review of the phenomenon, written by leading researchers, with the contributions of behavioural ecologists to help integrate the organismal, neurobiological, and evolutionary aspects with behavioural and ecological aspects of this sort of parasitism. It’s academically-oriented, but if you can get through my posts, you should have no problems reading the book. And I really do recommend it, because the subject matter is absolutely lovely.

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Brunetta & Craig. Spider Silk: Evolution and 400 Million Years of Spinning, Waiting, Snagging, and Mating. (Yale University Press)
	A 2012 paperback release of a 2010 hardback, this book is worth re-advertising because it’s really great. If you’ve ever wondered about the diversity of silk in spiders, the myriad different uses of silk in spiders, and just how silk use has evolved to fulfil all these uses, then this book is exactly what you need. Highly recommended for any arachnophiliac of any level.

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Gould & Gould. Animal Architects: Building and the Evolution of Intelligence. (Basic Books)
	One of my favourite pet topics is animal intelligence (e.g.). If you’re also interested in it, then you should look into getting this book. It’s a compendium of architecture produced from all over the animal kingdom, examined in the light of behavioural biology and intelligence.

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Land & Nilsson. Animal Eyes. (2d ed.; Oxford University Press)
	An excellent resource for anyone interested in anatomy and sensory physiology. All types of animal eyes are  detailed in this book, along with their efficiency and nervous integration (how they’re used to make images), and evolutionary histories. It’s written by two leading authorities on the subject, so if you want a comprehensive overview of animal vision, this is it.

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Fortey. Horseshoe Crabs and Velvet Worms: The Story of the Animals and Plants That Time Has Left Behind. (Knopf)

This book is a natural history of various “living fossils” – not to worry, the fallacies of the term “living fossil” are explained in the book. Written by Richard Fortey, a veteran invertebrate palaeontologist who’s written a lot of other popular science books, all of which rank among my favourites. I could quibble that this one is a bit low on science compared to his other books, but it’s still a wonderful read.

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Sagarin. Learning From the Octopus: How Secrets from Nature Can Help Us Fight Terrorist Attacks, Natural Disasters, and Disease. (Basic Books)

Whenever people ask me what the point to being a zoologist is, or what the purpose of my research is, I struggle for an answer – I’m one of those hedonistic scientists, doing science just for the sake of satisfying my curiosity and thirst for knowledge. After reading this book, I don’t have to make stuff up anymore. Sagarin goes through various adaptations that organisms have, and uses them as inspiration for suggestions on how to improve our own ways of doign things in politics, security, and much more. It’s a light-hearted book, not an academic one, so I fully recommend it for an easy and fun read.

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Waldbauer. How Not to Be Eaten: The Insects Fight Back. (University of California Press)
	From hiding to playing dead to spitting acid to blowing themselves up, insects have many ways to defend themselves – and their predators have coevolved to dispatch those defences. This book is all about this fascinating stuff, and is aimed at the lay public, so get it if you want interesting tales and factoids from the world of insect natural history.

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Weis. Walking Sideways: The Remarkable World of Crabs. (Cornell University Press)

I know for a fact, from eating together with lesser beings who have not studied invertebrate zoology, that there is a huge demand for general books about invertebrate groups, similar to the myriad ones we have for charming vertebrates. This one covers it for crabs. Aimed at the lay reader, it goes through all the basics of crab biology and ecology. Reading it is like having a seafood lunch/dinner with me. It’s also suitable for more advanced readers, since it has a good literature list.

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Krasnov. Functional and Evolutionary Ecology of Fleas: A Model for Ecological Parasitology. (Cambridge University Press)
	A 2008 hardback re-published in 2012 as a paperback. My list, my rules: this counts as a new book. It’s all about fleas, but using them as a case study for the evolutionary and ecological aspects of parasitology. So it’s really of interest to anyone whos tudies parasitology. It’s an academic textbook though, not quite easy reading for a non-biologist.

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Jump to another list: Environmental and Climate Change; Evolution; Historical Geology; History of Science; Human Evolution and Anthropology; Palaeontology.

How do insects breathe? An outline of the tracheal system

Hexapods (including insects) have two pairs of openings on their thorax, called spiracles. These open into chitinous tubes called tracheae which then further subdivide until becoming less than 1 mm in diameter tracheoles, 2-3 µm away from metabolically-active tissues, forming a network all through the body of the insect, as seen in a larval Lestes dragonfly above (Kennedy, 1922). Oxygen diffuses from the outside through the spiracles and into this tracheal system – this is how an insect breathes, by diffusion. It’s also one of the major reasons behind the limit of insect size, and the converse is that the smallest insects don’t need a full tracheal system (e.g. the system of tiny ptiliid beetles is highly reduced) because the surface area:volume ratio doesn’t need to get expanded for diffusion to occur.

The spiracles are muscular valves in the insect body, and their opening can be controlled, mostly in order to regulate water loss, since the tracheal system is always saturated with water vapour. In fact, if control over them is lost (e.g. by nervous system malfunction), the insect will die of dehydration (Mellanby, 1935). Diagrammed above (Gullan & Cranston, 2004) is a cross-section of a typical spiracle, with the opening having hair for filtering and the valve for controlling air flow.

The tracheal system is very easy to look at by dissecting any large insect. Cockroaches are the classics, but if your students are squeamish, grasshoppers or bees should do the trick (even if they’re a bit harder to dissect for the inexperienced). Important is to keep the insect alive, anaesthetised with chloroform, because when the tracheae are filled with air, they’re silvery and easy to see. If you need a guide to such a dissection, contact me!

Development of the Tracheal System

The tracheal system derives from the invagination of the external cuticle of the embryonic insect – in effect, it’s nothing more than an extension of the exoskeleton. This is why tracheal tubes are always surrounded by cuticle. The precise details of the morphogenesis of the tracheal system is an area of intense research because of the tracheal system’s interesting branching pattern (similar to capillaries and lungs); see Affolter & Cassinus (2008) for a review. This research isn’t just pure, but has potential applications in regenerative medicine by informing us about how lungs are designed (Nichols et al., 2009).

The quick summary of the tracheal system’s development, from Drosophila larvae, is as follows: there are 10 pairs of placodes (~40 cells each) derived by budding off from the cuticle under the action of the genes trachealess and tango (Ghabrial et al., 2003), one in each embryonic segment corresponding to the adult 3nd thoracic segment to 8^th abdominal segment. Each of these then undergoes the exact same process independently. They bud off sideways, to form 6 primary branches that then join together. Then, the left and the right sides connect, unifying all the branches together into the tracheal system; further branching then takes place to form tracheoles (Samakovlis et al., 1996), and the end result is what you see in the diagram above (Capinera, 2010). It’s an efficient repetitive process, one that is controlled by 200 genes (Ghabrial et al., 2011) regulated by metabolic and oxygen levels around the body (Centanin et al., 2010).

The tracheal system doesn’t scale up equally with size, as discussed in this post. The tracheal system increases in relative mass the larger an insect gets, a phenomenon called tracheal hypermetry. In tenebrionid beetles at least, this mass increase is most noticeable in the legs (Kaiser et al., 2007); whether this is a general trait for insects still needs to be researched. The size of individual tracheae depends on oxygen concentrations during development (Loudon, 1988), with lower concentrations making wider tracheae.

Gas Exchange

Carbon dioxide is excreted by pumping the thorax and/or abdomen, either up and down or by scrunching it along its length, increasing the pressure to push the CO₂ out, as in a ventilation system. This also serves to increase the amount of oxygen that can enter the tissues, since it pushes the oxygen out of the tracheoles at the times when gas exchange is needed the most (movement and especially flight). When the insect is resting, this doesn’t occur (discontinuous gas exchange) – the spiracles are typically closed, opening only temporarily to let oxygen in and the built-up CO₂ out (Chown et al., 2006).

More ventilatory pressure can be produced by the circulation system, which forms a “valve” at the junction between thorax and abdomen. The insect circulatory system is unique in that blood flow is occasionally reversed. When this happens, these valves contract, causing the tracheal system to get compressed and then relaxed – a ventilation effect.

Another method some insects have to increase the rate of gas exchange is called passive suction. They store CO₂ as bicarbonate ions in the haemocoel (“bloodstream”), creating a vacuum when oxygen goes into the tissues but isn’t replaced by CO₂, thereby drawing in more oxygen from the outside (basic laws of physics). The bicarbonate ions are occasionally converted to CO₂ and released.

Westneat et al. (2003) reported a brand new mechanism of breathing in which the tracheae in the thorax and thorax compress and expand in fast cycles. This is hypothesised to be a form of active breathing, but its importance and extent is still under dispute.

Modifications of the Tracheal System

The tracheal system is modified according to the specific ecology of the insects, something most strikingly seen in aquatic larvae. Most of them have a closed tracheal system, meaning the spiracles are sealed off, with oxygen diffusing through that seal. To increase the surface area and thus increase the rate of oxygen diffusion, some have extended their tracheal system to outside the body, forming elaborate gills, best seen on the abdomen of larval mayflies, as diagrammed above (Bradley, 2009). The most unique system is that of chironomid larvae who are apneustic, getting their oxygen entirely by absorbing it through their cuticle and through gills. A similar set-up is found in Trichoptera (caddisflies), where this system is synapomorphic.

Other aquatic insects may not have special adaptations in the tracheal system, but will preserve a bubble of air around their spiracles (held up usually by specialised hairs), the plastron, and use it as a lung, maintaining it by surfacing occasionally. A tangential point: the limitations of the tracheal system are one of the reasons why there are no marine insects. These insects would have to stay near the surface, where they would get eaten to extinction by fish (whereas crustaceans migrate through the depths to avoid predation). Also, they would be too buoyant with air-filled tracheae to actually sink. A plastron would collapse from pressure at depth. Finally, pressure makes the total amount of air in the tracheal system limited, and the hypothetical marine insect wouldn’t be able to exchange gases fast enough to live.

The most interesting use of the tracheal system is for making sound, a function that evolved in the famous Madagascar hissing cockroach for use as social signals (Nelson, 1979); it’s also found in other members of the Gromphadorhina genus. The sound is achieved by the modification of the spiracle valve being enlarged, and the connected end of the trachea being similarly enlarged; in effect, it’s no different than a horn. Sounds are changed as the air flow in the horn changes (due to thorax pumping), and by how open the spiracle valve is.

Another accoustic use for the tracheal system is with the tympanal organ of those insects that can hear, e.g. grasshoppers and other orthopterans. In these insects, there is a trachea in the tympanal organ modified to become much larger or to form a tracheal sac, and it’s this that serves as the membrane which the sound waves vibrate (Heinrich et al., 1993), analogous to the three bones in our ear. A similar system exists in hemipterans, except there it’s called the tymbal organ, not the tympanal organ.

Evolution of the Tracheal System

Discussing the origin of the tracheal system requires a higher-level phylogeny of the arthropods. One of the hypotheses is that hexapods and myriapods (centi- and millipedes, and a couple of smaller groups) are sister groups forming the Tracheata clade (also known as Atelocerata, “without horns”, in reference to the lack of a secondary antenna). They’re united by their common possession of a tracheal system (among other things), meaning the tracheal system in hexapods is symplesiomorphic.

However, the validity of this taxon is under question with the Pancrustacea hypothesis gaining ground. This one states that Crustacea is paraphyletic and the Hexapoda are nested within it. This would mean that the tracheal system could be a hexapodan apomorphy. The pros and cons of each hypothesis fall outside the scope of this post.

The evolution of the tracheal system was a pivotal moment in the history of the insects, as it allowed insects to colonise both the land (by providing a way to breathe outside of water) and the air (by supporting aerobic metabolism in-flight (Komai, 2001)).

Outside of the hexapods, arthropods with a tracheal system include:

• Myriapods: Interestingly, they also have haemocyanins (the arthropod equivalent of haemoglobin), even if they don’t transport oxygen in their haemolymph (“blood”);
• Opilionids (harvestmen): one pair of spiracles on second opisthosomal segment;
• Pseudoscorpions: Spiracles on first two opisthosomal segments;
• Solifugae (camel spiders): Very extensive, insect-like tracheal system;
• Many araneomorphs: Modify either the second or both book lungs into tracheae;
• Ricinulei (hooded tickspiders): Spiracles at posterior of prosoma;
• Acari (mites, ticks): Spiracles can be anywhere;
• Onychophora (velvet worms): Same remark as myriapods, evidence from Kusche et al. (2002);
• Oniscoidea (woodlice);
• Tardigrada (water bears).

What is thus clear is that a tracheal system can emerge independently pretty easily in terrestrial arthropods, and that it is lacking in marine ones. Many arachnids have a tracheal system, and this is most likely to be a derived character originating from the book gills of a xiphosuran-like ancestor, not shared with the last common ancestor with the hexapods/tracheates (Bromhall, 1987). It’s thus obvious that it’s a key innovation enabling successful terrestrialisation in arthropods.

This is also a major hit against a possible homology between myriapodan and hexapodan tracheal systems, and thus also against the Tracheata hypothesis. However, as is often the case with such open questions in phylogeny, a homology can be reasonably argued for, as Klass & Kristensen (2001) do with a (biased) appeal to the phylogenetic value of spiracle positioning, muscularity, and innervation. I am personally against a homology, since I’m a big fan of the Pancrustacea concept, and a look at the entire tracheal system besides the spiracles reveals more differences than commonalities anyway.

And such rampant convergent evolution is interesting in and of itself. If I had a lab with lots of funding (a pipe dream at this point), one of the main projects will be to look at the tracheal system from an evo-devo perspective. How does a strikingly similar system arise independently in all arthropods that go on land? Are there any deep homologies to be found, or is it simply the easiest way to get a breathing function with an arthropodan body type? If I had to, I would use isopods as a model organism, since the Oniscoidea, a terrestrial isopod superfamily (woodlice et al.), has relatives that are all marine, and so marine and terrestrial can be easily compared to each other. But one can also look deeper in the arthropod by looking at onychophores and tardigrades – since the cuticle and its formation is essentially the same in all these groups, there is a reasonable case to be made for a similar pathway to tracheal system formation. If a reader happens to have a lab, get on it! (I’m not aware of any such research programs happening atm.)

References:

Affolter M & Caussinus E. 2008. Tracheal branching morphogenesis in Drosophila: new insights into cell behaviour and organ architecture. Development 135, 2055-2064.

Bradley TJ. 2009. Animal Osmoregulation.

Bromhall C. 1987. Spider tracheal systems. Tissue and Cell 19, 793-807.

Capinera J. 2010. Insects and Wildlife: Arthropods and their relationships with wild vertebrate animals.

Centanin L, Gorr TA & Wappner P. 2010. Tracheal remodelling in response to hypoxia. Journal of Insect Physiology 56, 447-454.

Chown SL, Gibbs AG, Hetz SK, Klok CJ, Lighton JRB & Marais E. 2006. Discontinuous gas exchange in insects: a clarification of hypotheses and approaches. Physiological and Biochemical Zoology 2, 79.

Ghabrial AS, Luschnig S, Metzstein MM & Krasnow MA. 2003. Branching morphogenesis of the Drosophila tracheal system. Annual Review of Cell and Developmental Biology 19, 623-647.

Ghabrial AS, Levi BP & Krasnow MA. 2011. A Systematic Screen for Tube Morphogenesis and Branching Genes in the Drosophila Tracheal System. PLoS Genetics 7, e1002087.

Gullan PJ & Cranston P. 2004. The Insects: An Outline of Entomology, 3^rd ed..

Heinrich R, Jatho M & Kalmring K. 1993. Acoustic transmission characteristics of the tympanal tracheae of bushcrickets (Tettigoniidae). II: Comparative studies of the tracheae of seven species. Journal of the Acoustical Society of America 93, 3481-3489.

Kaiser A, Klok CJ, Socha JJ, Lee W-K, Quinian MC & Harrison JF. 2007. Increase in tracheal investment with beetle size supports hypothesis of oxygen limitation on insect gigantism. PNAS 104, 13198-13203.

Kennedy CH. 1922. The Homologies of the Tracheal Branches in the Respiratory System of Insects. Ohio Journal of Science 22, 84-89.

Komai Y. 2001. Direct measurement of oxygen partial pressure in a flying bumblebee. Journal of Experimental Biology 204, 2999-3007.

Klass K & Kristensen NP. 2001. The ground plan and affinities of hexapods: recent progress and open problems. Annales de la Société Entomologique de France 37, 265-298.

Kusche K, Ruhberg H & Burmester T. 2002. A hemocyanin from the Onychophora and the emergence of respiratory proteins. PNAS 99, 10545-10548.

Loudon C. 1988. Development of Tenebrio molitor in low oxygen levels. Journal of Insect Physiology 34, 97-103.

Mellanby K. 1935. THE EVAPORATION OF WATER FROM INSECTS. Biological Reviews 10, 317-333.

Nelson MC. 1979. Sound production in the cockroach, Gromphadorhina portentosa: The sound-producing apparatus. Journal of Comparative Physiology A 132, 27-38.

Nichols JE, Niles JA & Cortiella J. 2009. Design and development of tissue engineered lung: Progress and challenges. Organogenesis 5, 57-61.

Samakovlis C, Manning G, Steneberg P, Hacohen N, Cantera R & Krasnow MA. 1996. Genetic control of epithelial tube fusion during Drosophila tracheal development. Development 122, 3531-3536.

Westneat MW, Betz O, Blob RW, Fezzaa K, Cooper WJ & Lee W-K. 2003. Tracheal Respiration in Insects Visualized with Synchrotron X-ray Imaging. Science 299, 558-560.

Wolbachia: The Ubiquitous Male-Killing, Feminising Parasite

I mentioned Wolbachia in my parthenogenesis post; here I will talk about it in more detail, because it’s a really cool parasite, as well as in the center of much research nowadays.

Index:

• Introduction
• Parasitism
• Transfer
• Biocontrol
• Positive Effects of Wolbachia
• Evolutionary Theory
• Practical Problems
• References

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Introduction

Wolbachia are Gram-negative ⍺-Proteobacteria (order Rickettsiales, family Anaplasmataceae; Dumler et al.,2001), systematically split into 6 supergroups, lettered from A to F (Lo et al., 2002). Those in supergroups C and D have coevolved with their nematodan hosts and are mutualistic (Bandi et al., 1998). Those in supergroups A and B are parasites in arthropods. Morphologically, they come in two groups: ~1µm large rods, and ~0.5µm diameter coccoids. They can be found in a membrane-bound vacuole in the host cell’s cytoplasm (Yen & Barr, 1971); in arthropods, this will be in egg and ovarian cells (see diagram above, from Weiss et al. (2009)), although it has also been found in other tissues, including the Malpighian tubules, and muscle and nervous tissue (Stouthamer et al., 1999). When found in sperm cell cytoplasm, it disappears during spermatogenesis (Clark et al., 2002).

Note that in this post and in the literature, what’s referred to as Wolbachia is Wolbachia pipientis. Before DNA sequencing, when bacteria were classified morphologically, W. persica and W. melaphagi were also thought to be related, but these have nothing to do with the famous Wolbachia (W. persica is a γ-proteobacterium, W. melaphagi is a rhizobacterium).

Being present endosymbiotically in 20+% of all known insect species (Werren & Windsor, 2000), and given that some estimate that over half of all insect species are infected by it (Hilgenboecker et al., 2008), it’s obviously extremely successful, although it should be noted that the virulence of each Wolbachia strain is different (Min & Benzer, 1997), and each species is has a different susceptibility. In Australian and Panamanian fig wasps, Wolbachia has been found to infect over 70% of all species (Haine & Cook, 2005). A host database can be found here.

Parasitism

The biggest reason for its success is its parasitic effect: it’s always transferred maternally in the cytoplasm of the eggs, and in order to ensure maximal transfer rates, it alters its host’s reproductive abilities to favour their own reproductive success (Charlat et al., 2003). It has four ways to do this, in contrast to the one or two ways present in other such parasites:

• Inducing thelytokous parthenogenesis in haplodiploid organisms, e.g. hymenopterans, thus making sure that only female offspring are produced (e.g. Stouthamer et al., 1993). It occurs by forcing the fusion of the two nuclei of the first mitotic division (Huigens et al., 2000).
• Feminizing males, i.e. males reproduce as females (Wilkinson, 1998). The mechanism for this varies; for example, in the pillbug, Wolbachia blocks the formation of the androgenic gland which produces the masculinising androgenic hormone (Martin et al., 1990).
• Killing males, either in the embryonic stages or later (Hurst, 1991). The mechanisms of action are still largely unknown, although recent pioneering research by Riparbelli et al. (2012) shows that it happens due to Wolbachia (purposely?) messing around with male chromosomes at various stages in development, leading to defective embryos and death.
• Inducing cytoplasmic incompatibility (CI) between infected males and uninfected females (Hurst & Werren, 2001), causing sterility (Bordenstein & Werren, 2007) and eventually reproductive isolation within a species.

The latter was the first of Wolbachia‘s effects to be discovered, by Yen & Barr (1971) in mosquitoes; Wolbachia itself was first spotted by Hertig & Wolbach (1924). What happens is that the sperm enters the cell as normal, but its chromosomes don’t decondense and fuse with maternal chromosomes due to a delay in the breakdown of the nuclear envelope in the male’s pronucleus (Tram & Sullivan (2002); see Landmann et al. (2009) for more molecular details), and so can’t enter the first mitosis, meaning they get discarded (Lassy & Karr, 1996). The embryo either becomes a haploid female (in haplodiploid organisms), or it dies. In evolutionary theory terms, in a population susceptible to Wolbachia-induced CI, uninfected females become more unfit, therefore giving a fitness advantage to infected females (Bourtzis et al., 2003), hence explaining how the phenotype persists.

Which manipulation happens cannot be predicted even within the same species, as demonstrated by Hornett et al. (2008), who found that in their North American Hypolimnas bolina butterflies, Wolbachia was a male killer while in Southeast Asian ones, it was a CI-inducer. The difference came from a dominant allele in the SE Asian butterfly genome suppressing the male-killing. Charlat et al. (2007) showed that such a mutation can become fixed in a population in under 10 generations due to the extreme selection pressure to maintain a decent sex ratio, a testament to the ecological power of Wolbachia.

However, in some cases, Wolbachia can be so prominent that the entire affected population gets a very female-biased sex ratio, which in the case of the butterfly Acarea encedon has led to females reversing their sexual roles and behaving like males (Jiggins et al., 2000). In other cases, Wolbachia is counteracted by other elements, for example the B chromosome in the parasitoid wasp Trichogramma kaykai which turns Wolbachia-feminised males back into regular males (Stouthamer et al., 2001).

Transfer

It must be mentioned that part of their success is their ability to be transmitted horizontally across different species (Raychoudhury et al., 2009) – they aren’t host-specific. This has been shown as happening through parasitoids (Heath et al., 1999) or through the environment (e.g. by sharing a common food source (Huigens et al., 2000)). This also means that a single individual may have multiple Wolbachia species (or, better said, strains) co-existing and mingling inside it; the largest number I know of is eight, in the fire ant Solenopsis daguerrei (Dedeine et al., 2005). Note that this ability only comes in the arthropod-associated Wolbachia, whose genomes are more plastic, with recombination and phage-derived elements (Wu et al., 2004), none of which are characteristics present in their nematodan counterparts (Foster et al., 2005).

One study also reported the possibility of Wolbachia having transferred part or all of its genome to its hosts, albeit with only 2% of the genes able to be transcribed and none of them having any described effect (Hotopp et al., 2007).

Biocontrol

The negative effects of Wolbachia are obviously of great interest for biocontrol of pests and disease vectors. For example, Alam et al. (2011) discuss the possibility of using Wolbachia to control the tsetse fly, a vector of trypanosomiasis; Atyame et al. (2011) do the same for mosquitoes. Such biocontrol would work by allowing a chosen genotype to dominate the population by infecting the undesired genotype (e.g. Xi et al. (2005)), or by shortening lifespans to prevent sexual maturity (e.g. Moreira et al. (2009)). Wolbachia can also lead to population bottlenecks with very few individuals becoming able to reproduce (Nice et al., 2009), which is another way to control a pest population.

In non-pest studies, Wolbachia leads to increased susceptibility to parasitoids in Drosophila (Fytrou et al., 2006). It also leads to a less effective immune system in the pillbug Armadillidum vulgare, as seen by a lower density of haemocytes and higher density of bacteria (Bracquart-Vanier et al., 2008). These would be other avenues for pest control if a similar effect is seen in pest groups.

Positive Effects of Wolbachia

Interestingly, the effects of Wolbachia aren’t all negative. In the Cimicidae (bed bugs), Wolbachia is a mutualist; getting rid of it with antibiotics reduces the amount of food the host gets (Hosokawa et al., 2010). In mosquitoes, Wolbachia was found to boost their immune system and cause resistance to dengue virus (Bian et al., 2010). Pinto et al. (2012) describe how this happens at the genetic level. This is another potential use of Wolbachia as a biocontrol agent for disease vectors. In Drosophila, Wolbachia has been shown to confer resistance to several RNA viruses (Teixeira et al., 2008). In a Drosophila lab culture, Weeks et al. (2007) showed that the Wolbachia went from being a parasite to being a mutualist within two decades.

At the extreme end, nematode-infecting Wolbachia are needed for nematode development and fertility (Foster et al., 2005), so Wolbachia antibiotics could be used to control their populations (Taylor & Hoerauf, 1999). This is useful knowledge, given that some of the affected nematodes are vectors for very serious diseases like elephantiasis and onchocerciasis. It’s a similar story with the wasp Asobara tabida, wherein no ovocyte can even be produced when Wolbachia isn’t present (Dedeine et al., 2001) because its absence promotes excessive apoptosis in the ovarioles (Pannebakker et al., 2007).

Other effects of Wolbachia on sexual physiology have been documented, for example an increase in sperm competition in Tribolium beetles (Wade & Chang, 1995), or changes to the spermathecal duct in female Allonemobius crickets (Marshall, 2007).

Evolutionary Theory

An interesting point can be made about the process of molecular evolution in Wolbachia. As I said in the introductory paragraph, Wolbachia is a nematode mutualist and arthropod parasite (generally speaking). One of the Wolbachia genes involved in interaction with the host is wsp, which codes for cell membrane proteins. It was found to be undergoing divergent selection when it is in a parasitic relationship, but not when it’s in a mutualistic relationship (Jiggins et al., 2002). This is in line with what we expect: wsp being involved in host recognition means it theoretically should experience heightened evolutionary rates, and this is confirmed by the empirical data.

Many, if not all, negative and positive effects of Wolbachia have evolved by natural selection in order to maximise the transmission of the strain, either by allowing the bacterium to survive in the host (depressed immune system), or reducing competition by blocking the transmission of other pathogens (as Teixeira et al. (2008) suggest for the viral resistance effect). By extension, this means that parthenogenic arthropods aren’t expected to be Wolbachia hosts, since the manipulations are useless there. In terms of evolutionary theory, they can be treated as nothing more than selfish genetic elements.

When I first heard of Wolbachia, my intuition was that it played a sizeable role in speciation, since it promotes reproductive isolation, or by selecting for subdivided populations (Hatcher et al., 2000). Some analyses showed it not to be true (Rousset & Raymond, 1991), but more and more recent studies are supportive of the idea (Bordenstein, 2003), so it’s accepted as a cause of speciation. It definitely has been demonstrated (Thompson, 1987), and in some cases has also induced rapid speciation (Bordenstein et al., 2001).

On a general evolutionary synthesis level, Wolbachia is pretty interesting as a very recognisable case of inheritable symbiosis, one of the few proper examples that lend credence to the view that symbioses are a driving force behind evolution.

Milder effects of large-scale Wolbachia infection and sex ratio-skewing include altering dispersal ability – many insects have dispersing females and non-dispersing males, or vice versa. On a more influential level, there is also evidence that they can play a role in sexual selection (Jiggins et al., 2000), since sexual conflict gets reduced when levels of polyandry fall (Arnqvist & Rowe, 2005).

Practical Problems

Wolbachia can sometimes present a methodological stumbling block for molecular phylogenies based on mitochondrial DNA, since mtDNA will also hitchike maternally, favouring the maternal mtDNA haplotype, eventually leading to the entire dataset being worthless; see Ballard & Rand (2005) for more information. However, Arthofer et al. (2010) tested this idea using infected bark beetles and found no significant effect from Wolbachia, so it is still unsure just how significant this slight inaccuracy is.

Where it is definitely a problem is in barcoding initiatives using mtDNA. For successful barcoding of a species, a stable molecular marker needs to be used that is guaranteed not to vary across individuals, populations, or ecomorphs. However, there are some studies that show that Wolbachia causes divergences in mtDNA sequences even among individuals of the same species, e.g. in the butterfly genus Hypolimnas (Galtier et al., 2009). The reason for this is that mitochondrial genes are transferred only maternally, so only the maternal set plays a role in evolution. Given the ubiquity of Wolbachia, this is definitely a large problem that should be studied carefully before proceeding with mtDNA barcoding.

Wolbachia alone can’t be cultivated, but it is possible to keep a Wolbachia line using host cell lines (Noda et al., 2002), so experimental evolution studies are possible with them.

For the entomologists among you, make sure to check any colonies for Wolbachia infections, as they could invalidate your results, especially if you’re doing population biology. They can be gotten rid of using any antibiotic. I hear that tetracycline is recommended; if that’s not possible, high heat is enough, since Wolbachia is sensitive to temperature. If you’re sequencing your insects as well, using DNA from the legs is probably the safest way to avoid getting contaminating Wolbachia DNA amplified (this is standard procedure anyway).

Other symbionts that alter the reproduction of their arthropod hosts include Buchnera and Cardinium – but I’ll leave them for other posts.

References:

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Research Blogging necessities :)

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Cypriot Biodiversity Captured in 3 Minutes

I went on a fieldtrip up to the mountains yesterday, exploring some areas I hadn’t been to before. Among the things I discovered was this abandoned house with a blooming apple tree in front of it.

Apple trees are commonly cultivated or found wild at high altitudes – and I was only 200 meters below Mt. Olympus, the peak of the island. The house itself was inhabited by roaches, rats, and some bird (sounded like a pigeon). What follows are pictures I snapped quickly in 3 minutes – I was travelling light, with only a killing jar and the camera body with standard lens (no extension tubes or tripod). The aim of this post is to show you that insects are very easy to observe in nature – all you need to do is make sure the weather’s cool (not too hot, not too cold), and find somewhere where there’s a strong scent or strong primary colours – blooming trees or flower fields are the best.

All pictures have been cropped and downgraded to save webspace (also, not having the equipment means the pictures are not that good in the first place). I will also leave IDs off. You all can try and identify, or ask for tips :)

This being the mountains, coccinellids were pretty common. While they’re commonly known as biocontrol agents in agriculture, many coccinellids also have a reputation as cold-weather and high-altitude specialists. The following picture, for example, shows a coccinellid I took a picture of in March, found on a rock cropping out from the blanket of snow covering the mountain (temperature at the time ~1°C).

So, if you’re interested in observing ladybirds, your best bet is to go in the colder habitats.

The tree was inhabited by many beetles. Here’s two of them; such beetles are very easily collectable simply by placing a sheet under the tree, or holding a tray under the branch, and shaking. They all fall off and you can pick them off quickly before the fly off (usually they walk around in a daze, best time to grab them).

This one’s a pathetically out of focus multi-order image, and multi-trophic level image too: you have the carnivorous wasp (Hymenoptera), the beetles, and the pentatomid (Heteroptera) hiding underneath.

Some more beetles. The big brown one is especially attracted to colour – I was wearing a light blue t-shirt and always had at least two or three climbing on it.

I like to imagine this pentatomid feeling triumphant.

Here you can see the pentatomid. At all temperate times of the year, pentatomids invade the pine tree regions and can be found forming such congregations. On the left you see a game of hide-and-seek between the carnivorous ladybird and its herbivorous would-be prey (I don’t know whether it got eaten – this was all in three minutes, remember?).

Where there are flowers, there are bumblebees. You can tell this one’s pretty large.

Some more multi-order goodness. What the black thing under the coccinellid is is anyone’s guess. The out-of-depth-of-field brown blob is a hemipteran (of the leafhopper planthopper variety I would never dream of getting mixed up with).

And, moving away from the tree and its diverse fauna (much of which was way too small to photograph using a standard 35 mm lens), here’s a fly washing its hands.

Coming back to the car, I noticed many brown masses on the hood and roof. My car was a very welcome source of heat in the otherwise cool surroundings, and was quickly colonised by adventurous insects, like this water beetle look-alike. And by quickly, I mean a spider had built a web between the antenna and the roof in the five minutes I was gone.

And all this was merely during a 5 minute quick look in the area (I actually took a wrong turn and stumbled on this place). For parents or teachers reading, if you want to get your kids/students interested in insects or wildlife, simply find a fragrant tree in bloom and sit in front of it and observe.

And if you want more posts like this, please support my Petridish project, Toxic Geology: A Cause of Insect Endemism in Cyprus. It aims to investigate such habitats. You can find more details on the project page. Sharing it around your friends would be greatly appreciated; donating as well. Thanks!

Cixiidae (Hemiptera: Fulgoromorpha)

The Cixiidae Spinola 1839 are a cosmopolitan family of fulgoromorph (planthopper), comprising of over 1500 species in over 170 genera (Holzinger et al., 2002). Drawn above is Oteana (source: Hoch (2006)).

It is accepted as being relatively basal in the Fulgoromorpha (Yeh & Yang, 1999), and the only thorough cladistic analysis to date has recovered it as monophyletic (Ceotto & Bourgoin, 2008), with the following combination of apomorphies; none of these are unique to the Cixiidae, but their combination is unique:

• Flagelliform aedeagus;
• Sixth and seventh abdominal sternites formed by two sclerites;
• Six spines on the hind tibia’s apex;
• Tubercles on the forewing veins.

However, there is a lot of resistance to the idea of a monophyletic Cixiidae, since the level of homoplasy is high enough to cause problems for morphological analyses (Ceotto & Bourgoin, 2008). A particularly problematic family with respect to the Cixiidae is the Achilixiidae, a family which some studies place inside the Cixiidae (Liang, 2001), a position rejected by other studies (Urban & Cryan, 2006). Ceotto et al. (2008) conducted a thorough molecular analysis that recovered the Cixiidae as paraphyletic, with the Delphacidae arising from within the family. While the exact relationships need more work, it is mostly accepted that the family is paraphyletic.

The current higher-level classification used is in need of revision, precisely because of this paraphyly (see the Ceotto et al. (2008) phylogeny above). They are typically divided into three subfamilies: the Bothriocerinae Muir 1923, Borystheninae Emeljanov 1989, and Cixiinae Spinola 1839.

I sometimes confuse them with cicadas in the field; they basically look like very small cicadas. You can distinguish them with the following characters, which will tell you that you’re looking at a fulgoromorph, not a cicada: the antennal pedicel is enlarged or bulbous, the mesothorax has tegulae, and the bases of the middle coxae are widely separated. So if you’re going to collect them, make sure you have a hand lens with you. They can be recognised as cixiids instead of other fulgoromorphs by their clear, membraneous wings (see opening drawing) held at a very low angle to the body. Females can be recognised by their immoveable tibial spur; the sword-shaped ovipositor is also easily-recognisable, but is also found in delphacids.

Interestingly, some cixiids have patterns on their wings, namely dark patterns at the rear of the wings (Scherbakov & Popov, 2002). These serve to distract a predator by making it think the rear of the wing is the head of the animal.

Adults can be found on the surface of plants, and it’s easy to distinguish the sexes. Females have a distinct sword-shaped ovipositor, using it to lay eggs either in the soil or in plant tissue. If you want to catch them passively, a Malaise trap set up to capture inter-plant movement is the best choice. If you want an estimation of predation pressure on them, you can check out bird droppings: their hindlegs are often found there, and are easily recognisable (Ralph et al., 1985).

Nymphs commonly live at the base of plants on the surface, or between the roots (they commonly have flattened tibiae, possibly as digging adaptations). Some may be found on fungi on rotten wood, or on ferns; some are even associated with ants. In all though, the nymphs are hard to see because of their cryptic colouration and habits. Host plants or specificity aren’t known. Nymphs are very hard to identify to species-level, so you’ll most likely want to capture them alive and rear them to adulthood for ID.

Mating involves using vibrational signals: males send messages, females reply (Mazzoni et al., 2010); this is a cooption of their regular way of communication. Copulation can be pretty interesting: Hoch & Remane (1985) describe how Hyalesthes males have to be upside down and the females the right side up facing the opposite direction (if we humans were to try it like this, then the woman would end up with the man’s feet in her face while penetration was happening); Sforza & Bourgoin (1998) describe how the partners copulate side-by-side while facing opposite directions (in other words, the male has to twist his genitalia considerably to get them in; I don’t want to think about the human equivalent).

They’re somewhat unique in being one of only 6 hemipteran families with troglobitic, cave-dwelling representatives (Romero, 2009), with 80% of the cave-dwelling Fulgoromorpha being cixiids. The most famous cases are the genera Solanaima and Undarana in Australia, and Oliarus in Hawaii. Solanaima especially is considered to be a model system for studying evolution in caves, as all transitional stages are represented, from epigaean (surface-dwelling), to troglophilic (occasional cave-dwellers) to troglobitic (blind, flightless. unpigmented cave-dwellers) (Hoch & Howarth, 1989). Oliarus is also intensely studied, but more in relation to the time-scale involved in cavernicolous evolution. These species have independently moved into lava tubes, habitats whose formations can be precisely dated geologically, and study of them can yield important information about how fast features can be lost (Hoch & Howarth, 1999).

They’re also unique in being one of the few fulgoromorphs known to act as vectors for plant viruses and bacteria (Nault & Ammar, 1989), e.g. Hyalesthes obsoletus transmitting stolbur phytoplasmas (phloem-dwelling pathogenic bacteria) on grapevines (Maixner et al., 1995), the cause of what’s known as Vergilbungskrankheit (German-speaking wine-growing areas), Bois noir (French), or Legno nero (Italian), a disease that leads to shrivelled grapes, downward-rolling leaves, and lack of lignified shoots. Needless to say, it’s a huge harm to for any wine production, sometimes with over 50% of vines becoming infected during an epidemic, so research into associated cixiids is pretty active. The transmission is completely accidental, there is no manipulation by the phytoplasma. The larvae acquire them while feeding on stinging nettle roots, and they’re transmitted by the adults to the grapevine when they pierce the grapevine to suck the phloem out (similar to how mosquitoes inject malaria into us). The grapevine is a dead-end host for the phytoplasma.

Another disease transmitted by a cixiid, Haplaxius crudus (still often erroneously referred to as Myndus crudus due to general ignorance of taxonomy; New World Myndus were placed in Haplaxius by Emeljanov (1989)), is lethal yellowing, a disease of palm trees, very damaging in the Carribean region throughout this species’s range. Any infected palm is guaranteed to die: it’s single-handedly responsible for the elimination of coconut palms on the Carribean coast of central America. Several control methods have been tried, from standard insecticides (only good for preventing dispersal), to planting plants unsuitable for nymph growth, but the most successful way has simply been to plant different species of palm known to be more resistant to the disease; hopefully, this will lead to some sort of GM research to get their resistance-enabling genes into the more susceptible date- and coconut palms.

Besides those, they cause structural damage to plants by feeding (H. crudus is again implicated, but with sugarcane), but this is considered negligible relative to other pests.

Their fossil record is rather sparse, although they are one of the commonest groups found in Baltic amber (Szwedo et al., 2006); one is pictured above (Sczwedo & Sontag, 2009). As should be expected, the bulk of their record comes from ambers, specifically Dominican (e.g. Szwedo, 2000) and Baltic amber (Gębicki & Szwedo, 2000), as well as Lebanese amber (Szwedo, 2001). However, the oldest record comes in the form of ‘Cixius’ petrinus, preserved in real rock from the Early Cretaceous Weald Clay, UK (Fennah, 1961).

References:

Ceotto P & Bourgoin T. 2008. Insights into the phylogenetic relationships within Cixiidae (Hemiptera: Fulgoromorpha): cladistic analysis of a morphological dataset. Systematic Entomology 33, 484-500.

Ceotto P, Kergoat GJ, Rasplus J-Y & Bopurgoin T. 2008. Molecular phylogenetics of cixiid planthoppers (Hemiptera: Fulgoromorpha): New insights from combined analyses of mitochondrial and nuclear genes. Molecular Phylogenetics and Evolution 48, 667-678.

Emeljanov AF. 1989. On the problem of division of the family Cixiidae (Homoptera, Cicadina). Entomological Review 68, 54-67.

Fennah RG. 1961. The occurrence of a Cixiinae fulgoroid (Homoptera) in a Weald Clay. Annals and Magazine of Natural History 13, 161-163.

Gębicki C & Szwedo J. 2000. Kulikamia jantaris gen. et sp. n. from Baltic amber (Hemiptera: Fulogroidea: Cixiidae). Polskie Pismo Entomologisczne 69, 167-173.

Hoch H. 2006. New Cixiidae from Eastern Polynesia: Oteana gen.nov. and Manurevana gen. nov. (Hemiptera: Fulgoromorpha). Zootaxa 1209, 1-47.

Hoch H & Remane R. 1985. Evolution und Speziation der Zikaden-Gattung Hyalesthes Signoret, 1865 (Homoptera Auchenorrhyncha Fulgoroidea Cixiidae). Marburger entomologische Publikationen 2, 1-427.

Hoch H & Howarth FG. 1989. Six new cavernicolous cixiid planthoppers in the genus Solanaima from Australia (Homoptera: Fulgoroidea). Systematic Entomology 14, 377-402.

Hoch H & Howarth FG. 1999. Multiple cave invasions by species of the planthopper genus Oliarus in Hawaii (Homoptera: Fulgoroidea:Cixiidae). Zoological Journal of the Linnean Society 127, 453-475.

Holzinger W, Emeljanov AF & Kammer-Lander I. 2002. The family Cixiidae – A review. In: Holzinger W (ed.). Zikaden — Leafhoppers, Planthoppers and Cicadas (Insecta: Hemiptera: Auchenorrhyncha), Denisia 4.

Liang AP. 2001. Morphology of antennal sensilla in Achilixius sandakanensis Muir (Hemiptera: Fulgoromorpha: Achilixiidae) with comments on the phylogenetic position of the Achilixiidae. Raffles Bulletin of Zoology 49, 221-225.

Maixner M, Ahrens U & Seemüller E. 1995. Detection of the German grapevine yellows (Verglibungskrankheit) MLO in grapevine, alternative hosts and a vector by a specific PCR procedure. European Journal of Plant Pathology 101, 241-250.

Mazzoni V, Lucchi A, Ioriatti C, Virant-Doberlet M & Anfora G. 2010. Mating Behavior of Hyalesthes obsoletus (Hemiptera: Cixiidae). Annals of the Entomological Society of America 103, 813-822.

Nault LR & Ammar ED. 1989. Leafhopper and Planthopper Transmission of Plant Viruses. Annual Review of Entomology 34, 503-529.

Ralph CP, Nagata SE & Ralph CJ. 1985. Analysis of bird droppings to describe diets of small birds. Journal of Field Ornithology 56, 165-174.

Romero A. 2009. Cave Biology: Life in Darkness.

Scherbakov DE & Popov YA. 2002. Order Hemiptera Linné, 1758. The bugs, cicadas, plantlice, scale insects, etc. In: Rasnitsyn AP & Quicke DLJ (eds.). History of Insects.

Sforza R & Bourgoin T. 1998. Female genitalia and copulation of the planthopper Hyalesthes obsoletus Signoret (Hemiptera: Fulgoromorpha: Cixiidae). Annales de la Société entomologique de France (NS) 34, 63-70.

Szwedo J. 2000. Oliarus kulickae sp. n. from Dominican amber (Hemiptera: Fulgoroidea: Cixiidae). Polskie Pismo Entomologiczne 69, 161-166.

Szwedo J. 2001. A substitute name for the extinct genus Mundopoides Fennah (Hemiptera: Fulgoroidea: Cixiidae). Genus 12, 275.

Szwedo J & Sontag E. 2009. The traps of the “amber trap”. How inclusions could trap scientists with enigmas. Denisia 26, 155-169.

Szwedo J, Bourguin T & Lefebvre F. 2006. New Mnemosynini taxa (Hemiptera, Fulgoromorpha: Cixiidae) from the Palaeogene of France with notes on their early association with host plants. Zootaxa 1112, 31-41.

Urban JM & Cryan JR. 2006. Evolution of the planthoppers (Insecta: Hemiptera: Fulgoroidea). Molecular Phylogenetics and Evolution

Yeh WB & Yang CT. 1999. Fulgoromorpha phylogeny based on the 28S rDNA nucleotide sequence. Chinese Journal of Entomology 11, 87-111.

Bed Bugs (Cimicidae: Cimex lectularius)

This is a guest post by Sarah Rexman from BedBugs.org.

Bed bugs are parasites that feed on human blood. They are in the Hemiptera order and the Heteroptera suborder, and they belong to six subfamilies, including Afrociminae, Cimicinae, Cacodminae, Haematosiphoninae, Latrocimicinae, and Primicimicinae.

Bed bugs are often found in mattresses and bed frames, and often feed on people while they are sleeping – thus their common name, “bed bugs.”

Bed bugs are typically in the genus Cimex and the family Cimicidae. However, what the general population knows to be the common bed bug is the Cimex lectularius.

Adult bed bugs are light brown or reddish brown in colour, and have a flattened, oval shape resembling an apple seed. Adults can reach 4-5 mm in length and 1.5-3 mm in width. Babies (known as nymph or larvae) can be translucent and invisible to the naked eye.

They have no hind wings, and their front wings are vestigial. They have segmented abdomens and a long, segmented proboscis that extends forward when eating. The proboscis rests beneath the body when the bug is not eating, and it projects behind the legs.

History

Bed bugs have been common in the developing world, but were largely eradicated in the 1940s and 1950s. They have started to see a resurgence since the mid-1990s. Some believe that the invention of DDT helped to reduce infestations.

Bed bugs have been around for thousands of years, with mentioned made about them in ancient Greece as early as 400 BC. Some early peoples believed that bed bugs had medicinal properties.

Feeding Habits and Environment

Bed bugs are known as such for their preference for living in mattresses and box springs. However, they are also found in other furnishings, such as couches and chairs, as well as in carpets and in cracks in walls and other structures. Bed bugs can even be found on toys, luggage, or clothing.

They can live in a wide variety of climates, surviving temperatures as low as 14 degrees F and as high as 113 degrees F. They can survive in temperatures with high and low humidity.

Bed bugs typically come out at night to feed, but they are not exclusively nocturnal. Bed bugs feed on humans or animals, and they are attracted to their hosts by carbon dioxide, warmth and moisture.

Bed bugs feed on blood by sucking, and they can feed for three to five minutes at a time. It can take five to ten minutes for a bed bug to become completely engorged with blood. Feeding can cause itchy, red welts on hosts, but bed bugs can feed without being noticed.

Bed bugs usually feed every five to ten days, but some can live for as much as a year without feeding – making them hard to eradicate.

Life Cycle

Bed bugs have five immature life stages and one adult life stage. They must molt six times after a blood meal before they can become fertile adults. This development process is known as “gradual metamorphosis.”

Development through these five larval stages takes about a month under optimal conditions, including a suitable host. However, bugs can remain in the larval stage for many months without a blood meal.

The total lifespan of a bed bug depends on the species and its feeding habits.

Bed bugs mate by traumatic insemination. Though females have a reproductive tract, the males do not deposit sperm there. Instead, male bed bugs pierce the female’s abdomen and ejaculate into the body cavity. Sperm travel through the blood to sperm storage structures known as seminal conceptacles, and fertilization later takes place in the ovaries.

Females require a blood meal to develop eggs. If they have access to blood meals, they can lay eggs mostly continuously and are capable of laying 500 eggs over a lifetime.

Infestations

Bed bugs have become well-known in recent years for their infestation of homes and hotels throughout the country. They can cause a number of health problems, including rash and allergy. Bed bugs have been found to carry 28 different human pathogens, including bacteria, viruses, and disease.

Infestations occur because bed bugs can reproduce quickly and can hide in hard-to-reach locations such as inside furniture or cracks in walls. Bed bugs also spread quickly through contact with infected homes and hotels, transferring through clothing, luggage, pets, or personal items. Bed bugs can also travel through duct work or other passageways between dwellings, such as in apartment buildings.

Symptoms of bed bug infestation include dark spots on furniture or along seams, which can be from feces or dried blood. Adult bugs can also be seen on furniture, behind baseboards, in cracks, and other favourite hiding places.

Trained dogs can also be used to sniff out bed bug infestations.

Bed bugs can be eliminated by thorough cleaning of furniture, rugs and other items through vacuuming, wiping with rubbing alcohol, and steam cleaning. Extreme heat can kill bed bugs, so it is recommended that all linens and other fabric items be washed and dried on the highest temperature settings possible, and a steam cleaner be used for all other items.

Pesticide treatment can also be administered by a professional.

References

Goddard J & deShazo R. 2009. Bed bugs (Cimex lectularius) and clinical consequences of their bites. JAMA 301, 1358–66.

Reinhardt K & Siva-Jothy MT. 2007. “Biology of the Bed Bugs (Cimicidae)”. Annual Review of Entomology 52, 351–374.

“What Are Bed Bugs? How To Kill Bed Bugs”. Medical News Today. MediLexicon International Ltd. 20 Jul 2009.

Storey M. “CIMICIDAE (bed bugs)”. BioImages: The Virtual Field-Guide (UK).

About the author:

Sarah Rexman is the main researcher and writer for BedBugs.org. Her most recent accomplishment includes graduating from Florida State, with a master’s degree in environmental science. Her main focus for the site involves showing people where to purchase bed bug repellent as well as where to find reputable bed bugs exterminators.