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I'm gonna make a presumptive statement: It -seems- as though I get bitten by mosquitos and other biting insects in -exactly- the same places over and over. I mean within a few mm. Is that true?
If true, that implies that insects have a mechanism to detect certain preferred spots in order to get blood or whatever they want. (Location of blood vessels? Sweat glands?)
And if that is so, what -is- that mechanism? What are they going for? And how does this detection work?
IOW: How do they know to bite me exactly on the side of my right knee over and over and over… but never on other spots?
There are at least two sources that influence patterns of behavior: the individual that is doing the biting, and the individual that is being bitten.
The example of lice in one of the comments is a good example. Three categories of lice that get on humans are: head lice, body lice, and pubic lice. They are related species (possibly the same species in some cases - https://www.insidescience.org/content/are-head-and-body-lice-same-species/550), but each type has their own preference for habitat and feeding. There are all kinds of reasons for a species to have a preference, and some stick to those preferences quite strongly.
As for your mosquitoes, their absolute preferences are still not known but blood seeking insects do have their senses attuned to clues such as carbon dioxide and heat (that help identify potential hosts) as well smells and other influences on their senses. There might be something special about the side of your right knee as far as mosquito senses go, but not every individual in a species sees, smells, hears, etc. the same way. My brother has a gene that makes broccoli taste very bitter, but I am fortunate to not have those taste buds. Color blindness is another example of differences in perception, but even without being color blind, what you see as red is probably not how someone else sees red due to natural variations in the patterns of sensory cells in the eye. For a consistent pattern of behavior from the mosquitoes, in spite of individual differences we still need to consider another source of variation that might lead to this mosquito preference: the host.
If the variation of individual mosquitoes is trumped by a consistent pattern of predation, perhaps the mediating factor is the host. Your knee could have a special smell, or other sensory flag that attracts mosquitoes. There are efforts to understand how a mosquito's perception directs who gets bitten: http://www.scientificamerican.com/video/future-repellents-mess-with-mosquit2013-12-23/ . Not enough is known yet about the preferences of mosquitoes to say for sure what makes your knee so good. There is also a chance that your knee is not special to the mosquito because it appears the most appetizing.
Host behavior is another possible influence. The knee is just far enough away from the reach of your hands. Conscious or absentminded grooming would both be limited at that distance. The knee may also be better than an ankle or a foot, because of what is worn or because the knee moves less than the lower leg during walking and other movements. If you are bitten on the lateral (outside) side of your knee, this would add evidence to this hypothesis of your movement influencing mosquito behavior, because the inside of your knee more frequently brushes against the other leg. The frequency of the inside of the knee being a safe place for a meal is much lower.
As for why it's the right knee rather than the left, there is possibility that that is also a reflection of you, rather than the pests. Humans display preferences in dominance for both hands and feet. One side of your body may be selected more often than another because of you being right- or left-handed, or right- or left-footed. This could affect grooming habits (how you swat at the bugs), how your cross your legs, which side you carry items or lean against other objects which might brush against the body, influence how well you spray yourself with bug repellent or who knows what else. (Just for curiosity's sake, ¿are you right-handed or are you left-handed?)
In short: There's a lot of evidence to collect before we could come to a definitive answer. There are a lot of possibilities, including that your knee may not be the tastiest spot, but rather a relatively safe spot for dinner.
For more unempirical thoughts on the relationship of insects to the lower limbs, please see: http://fishboyridesagain.blogspot.com/2005/11/bugs-and-trousers.html
Parasitic Mites of Humans
Mites are very small arthropods which are closely related to ticks. Mite larvae have six legs whereas the nymphal and adult stages have eight. Most species of mites are pests of agricultural crops. However, certain types of mites are parasitic on humans.
Chiggers are the larvae of a family of mites that are sometimes called red bugs. The adults are large, red mites often seen running over pavement and lawns. Chiggers are extremely small (0.5 mm) and are difficult to see without magnification. The six-legged larvae are hairy and yellow-orange or light red. They are usually encountered outdoors in low, damp places where vegetation is rank and grass and weeds are overgrown. Some species also infest drier areas, however, making it difficult to predict where an infestation will occur.
Chiggers overwinter as adults in the soil, becoming active in the spring. Eggs are laid on the soil. After hatching, the larvae crawl about until they locate and attach to a suitable host. The larvae do not burrow into the skin, but inject a salivary fluid which produces a hardened, raised area around them. Body fluids from the host are withdrawn through a feeding tube. Larvae feed for about 4 days and then drop off and molt to nonparasitic nymphs and adults. Chiggers feed on a variety of wild and domestic animals, as well as humans. The life cycle (from egg to egg) is completed in about 50 days.
Most people react to chigger bites by developing reddish welts within 24 hours. Intense itching accompanies the welts, which may persist for a week or longer if not treated. Bites commonly occur around the ankles, waistline, armpits, or other areas where clothing fits tightly against the skin. Besides causing intense itching, chigger bites that are scratched may result in infection and sometimes fever. Chiggers in North America are not known to transmit disease.
Persons walking in chigger-infested areas can be protected by treating clothing (cuffs, socks, waistline, sleeves) or exposed skin with tick repellents. Some repellents should only be used on clothing and it is important to follow label directions. People who suspect they may have been attacked by chiggers should take a soapy bath immediately and apply antiseptic to any welts. A local anesthetic will provide temporary relief from itching.
Regular mowing and removal of weeds and brush make areas less suitable for chiggers and their wild hosts. Mowing also enhances penetration and performance of miticides, should they be required. Chigger populations can be further reduced by treating infested areas with residual miticides. Applications should be thorough but restricted to areas frequented and suspected of being infested.
The sarcoptic itch mites, Sarcoptes scabei, infest the skin of a variety of animals including humans. The types of Sarcoptes inhabiting the skin of mammals are all considered forms of Sarcoptes scabei and can exchange hosts to some degree. (For example, Canine scabies can be temporarily transferred from dogs to humans, causing itching and lesions on the waist, chest and forearms.)
Human scabies mites are very small and are rarely seen. They commonly attack the thin skin between the fingers, the bend of the elbow and knee, the penis, breasts, and the shoulder blades. The mites burrow into the skin, making tunnels up to 3 mm (0.1 inch) long. When they first burrow into the skin, the mites cause little irritation, but after about a month, sensitization begins. A rash appears in the area of the burrows and intense itching is experienced.
Scabies mites are transmitted by close personal contact, usually from sleeping in the same bed. Bedridden individuals in institutions (e.g., nursing homes) may also pass the mites from caregiver to patient. The adult fertilized female mite is usually the infective life stage. She adheres to the skin using suckers on her legs and burrows into the skin where she lays her oval eggs. In 3 to 5 days these eggs hatch into larvae and move freely over the skin. Soon they transform into nymphs and reach maturity 10 to 14 days after hatching.
A scabies infestation should be handled as a medical problem and is readily diagnosed and treated by most physicians. (Confirmation requires isolating the mites in a skin scraping.) The first step to control a scabies infestation usually involves softening the skin with soap and water to make sure the pesticide treatments can penetrate well. An evening bath followed by overnight treatment works best. A total body (neck- down) application of topical pesticide medication should remain for 8-12 hours before showering in the morning. Commonly used products include lindane (Kwell (tm)), permethrin (Elimite (tm)) and crotamiton (Eurax (tm)). Follow directions on the product package carefully.
Because the symptoms of scabies mite infestations are delayed by about a month, other members of the household besides those showing symptoms may be harboring the mites. It is important that everyone in the infected family or living group go through the treatment regime. A second treatment may be necessary to eliminate an infestation of scabies mites, but patients should avoid overzealous pesticide treatment since itching may persist for a week or more after treatment and does not necessarily indicate treatment failure.
Scabies mites cannot live off of a human host for more than 24 hours. Therefore, insecticide treatment of premises is not warranted. It is recommended, however, that coincident with treatment, the clothing and bedding from infested individuals be washed in hot water or dry cleaned.
Bird and Rodent Mites
Parasitic mites that occasionally infest buildings are usually associated with wild or domestic birds or rodents. Bird and rodent mites normally live on the host or in their nests, but migrate to other areas of the structure when the animal dies or abandons the nest. Rodent mites often become a nuisance after an infestation of mice or rats has been eliminated. People usually become aware of the problem when they are attacked by mites searching for an alternate food source. Their bites cause moderate to intense itching and irritation. Rodent and bird mites are very tiny, but usually can be seen with the naked eye. They are about the size of the period at the end of this sentence.
The first step in controlling bird or rodent mites is to eliminate the host animals and remove their nesting sites. Often, the nests will be found in the attic, around the eaves and rafters, or in the gutters or chimney. Gloves should be used when handling dead animals. A respirator should also be worn when removing nest materials to avoid inhaling fungal spores and other potential disease-producing organisms associated with the droppings.
After nests are removed, the areas adjacent to the nest should be sprayed or dusted with a residual insecticide such as those products labeled for flea control. Space or ULV treatments with non-residual materials (e.g., synergized pyrethrins) can be used in conjunction with residual sprays. Space treatments are especially useful when the mite infestation has dispersed widely from the nesting site. In this case, more extensive treatment with residual and non-residual insecticides may also be necessary in other areas of the structure where mites are observed. A vacuum cleaner or cloth moistened with alcohol can be used to eliminate mites crawling on open surfaces.
CAUTION! Pesticide recommendations in this publication are registered for use in Kentucky, USA ONLY! The use of some products may not be legal in your state or country. Please check with your local county agent or regulatory official before using any pesticide mentioned in this publication.
Of course, ALWAYS READ AND FOLLOW LABEL DIRECTIONS FOR SAFE USE OF ANY PESTICIDE!
Sensing Polarized Light in Insects
Evolution has produced vast morphological and behavioral diversity amongst insects, including very successful adaptations to a diverse range of ecological niches spanning the invasion of the sky by flying insects, the crawling lifestyle on (or below) the earth, and the (semi-)aquatic life on (or below) the water surface. Developing the ability to extract a maximal amount of useful information from their environment was crucial for ensuring the survival of many insect species. Navigating insects rely heavily on a combination of different visual and non-visual cues to reliably orient under a wide spectrum of environmental conditions while avoiding predators. The pattern of linearly polarized skylight that results from scattering of sunlight in the atmosphere is one important navigational cue that many insects can detect. Here we summarize progress made toward understanding how different insect species sense polarized light. First, we present behavioral studies with “true” insect navigators (central-place foragers, like honeybees or desert ants), as well as insects that rely on polarized light to improve more “basic” orientation skills (like dung beetles). Second, we provide an overview over the anatomical basis of the polarized light detection system that these insects use, as well as the underlying neural circuitry. Third, we emphasize the importance of physiological studies (electrophysiology, as well as genetically encoded activity indicators, in Drosophila) for understanding both the structure and function of polarized light circuitry in the insect brain. We also discuss the importance of an alternative source of polarized light that can be detected by many insects: linearly polarized light reflected off shiny surfaces like water represents an important environmental factor, yet the anatomy and physiology of underlying circuits remain incompletely understood.
The periodic illumination by sunlight is essential for directly fueling plant life on the earth, which in turn serves as an indispensable food source for all animal life. Sunlight is also crucial for synchronizing circadian behaviors across the animal kingdom, but it also enabled the evolution of a variety of photosensitive eye structures crucial for the successful adaptation to both nocturnal and diurnal lifestyles across a wide range of light intensities (Land & Fernald, 1992). Just like all other senses, visual systems across phyla evolved the ability to extract, distinguish, and quantify different modalities of incident or reflected sunlight, like intensity (brightness) or wavelength composition (color). Complex neural circuits underlying these processes have been characterized using behavioral, anatomical, and physiological approaches. Across species, striking similarities exist between the neuronal arrangements processing a specific feature, such as the local intensity increments or decrements associated with the perception of moving objects. Recently, understanding the computations performed by these circuits down to a cellular level has started to come within reach (Borst, 2014).
Figure 1: Polarized light sources in the environment. Direct sunlight is unpolarized, meaning all e-vector orientations are equally represented (symbolized by the white, double headed arrows. Depicted are four different sources of polarization (from left to right): (1) reflection off water, leading to linearly polarized light with a horizontally oriented e-vector (2) reflection off of shiny leaves, can lead to an oblique e-vector orientation (3) reflections off the body surface of a scarab beetle lead to circular polarization with a rotating e-vector (4) celestial pattern of linearly polarized light is created by scattering of sunlight in the atmosphere, which is strongest at an angle of 90 degrees from the sun. Therefore, e-vector orientation and degree of polarization vary across the sky (symbolized by thickness and orientation of the bars) in a predictable pattern that is centered around the sun.
In order to maximize an animal’s chance of survival, its nervous system had to evolve the ability to constantly extract specific features from the visual world that are useful for its lifestyle, filter them for relevance, and then combine them in a meaningful way to ensure an optimal behavioral output. It is therefore no surprise that many animal species developed very specialized sensitivities to specific visual modalities. One specific example is the perception of linearly polarized light, an ability that can be found across a large number of species involving vertebrates (birds, fishes), as well as marine invertebrates (Cephalopods, Crustaceans), and many flying or crawling insects (Cronin et al., 2003 Nilsson & Warrant, 1999). Although many of these species may use the information encoded by a stimulus containing polarized light for different purposes, they all detect the same physical property— the e-vector orientation of incident light (Wehner, 2001). Normally, sunlight is unpolarized, meaning its e-vectors are distributed randomly. However, different phenomena can result in the overrepresentation of certain e-vector orientations (summarized in Figure 1). First, there is scattering of sunlight in the atmosphere, resulting in linearly polarized skylight. Interestingly, both degree of polarization as well as e-vector orientation vary across the sky, creating a celestial polarization pattern that is centered around the sun and therefore changes over time from an observer’s perspective. Second, reflection of sunlight off of shiny surfaces results in linearly polarized reflections with defined e-vector orientations (horizontal, in the case of water bodies). Ponds or lakes, as well as shiny leaves or even moist skin or fur can create such linearly polarized reflections. Third, reflection of sunlight from certain materials, like the body surface of certain invertebrates, can result in circular or elliptical polarization, meaning the e-vector orientation changes in a circular fashion as the light beam propagates. Therefore, all three sources of polarized light provide crucial cues to many animals, especially insects. Polarized light can serve as: (1) information about the position of the sun (which itself might be obstructed from view), providing an additional celestial orientation cue baring important information to facilitate navigational tasks (2) a signal indicating the presence of water as a potentially attractive or repulsive surface (depending on the behavioral context) (3) polarized reflections off different objects that may encode attractiveness as a food source or as suitable substrate for egg laying or (4) polarized reflections off the body surface a conspecific individual signalizing availability for mating or to signal competition.
In the following sections we will summarize the existing work covering the different roles polarized lights can play for different insects, focusing on progress from more recent studies. We group the text into three broad research areas, which, when taken together, form a synergistic picture of the neuroethology of e-vector detection: first, we focus on the lessons that can be learned from the careful quantitative description of an animal’s behavioral responses to celestial and reflected polarized light stimuli. Second, we describe the anatomical techniques used to characterize both polarization sensitive photoreceptor neurons, as well as their synaptic connectivity. Third, we describe how targeted and unbiased physiologic characterization of polarization-sensitive neurons from the retina to the central brain results in a growing cellular model of polarization circuitry in the insect brain.
The huge morphological and behavioral diversity within the insect world provides a wide range of stunning examples of how different insect species utilize polarized light stimuli in order to adapt to their environment. Classic examples are true point-to-point navigators like honeybees and desert ants as well as species that use polarized light to improve their orientation skills, especially dung beetles, crickets, desert locusts, butterflies, and flies. Behavioral responses to polarized reflections from water are less well understood, although a growing number of reports have described this behavior. However, the importance of polarized reflections off other substrates remains much less well understood.
Polarized Light and Navigation Behavior of Central Place Foragers
Figure 2: Honeybees deduce solar position from perceived e-vector orientations. Summary of experiments where honeybees were trained to fly through a tunnel toward a feeder. A polarizing foil mounted onto the tunnel ceiling resulted in the bees experiencing exclusively axial vectors (A–C), orthogonal e-vectors (D–F), or one followed by the other (G–I). The sun was always blocked off and could not be used as a landmark by the animals. (B) During the consecutive waggle dance, the bees communicated the direction to the food source by dancing a straight line at a defined angle to the sun. Individual waggle dance traces (red) plotted after flights through a tunnel with axial e-vectors were directed predominantly at orthogonal angles to the solar meridian (mean axis in turquoise). (C) Schematic summarizes the natural relationship between axial e-vectors in the zenith and the solar position associated with it (solar meridian is orthogonal to the bees path). (E) Waggle dance traces (red) are oriented along the solar meridian when orthogonal e-vectors were perceived, which is in agreement with the natural relationship between these stimuli (see F). The perception of both axial and orthogonal e-vectors arranged consecutively within the same tunnel led to dance directions (mean axis in turquoise) along the four diagonal axes (dotted black lines). (H) The four dance directions observed under a natural sky, after training in atunnel with two e-vectors (G), can be explained by the summation of two vectors (I). (Adapted with permission from Evangelista et al., 2014.)
The importance of linearly polarized light as a guidance cue for navigating insects was first demonstrated in 1949 , when Karl von Frisch was experimenting with honeybees, which need to reliably find their way from the hive to a food source and back (von Frisch, 1949). Therefore, they serve as an excellent insect model system for truly navigating “central place foragers.” In von Frisch’s experiments, honeybee workers were able to correctly indicate to their nestmates the angular direction toward a food source during the “waggle dance,” even when the sun was occluded from their field of view. Through a series of tests, von Frisch identified the celestial polarized light pattern (which is always centered around the sun) as the source of this behavior, a discovery that was later confirmed and elaborated on by Rossel and Wehner (1986 for review see Rossel, 1993). These experiments revealed that honeybees can deduce the position of the sun (which serves as the reference for the “waggle” dance) from the perceived polarization pattern. This is particularly useful in situations when the sun is not visible, for instance, when hidden behind a cloud. Hence, by using both solar position (if available) and the celestial polarization pattern, the honeybee internal compass becomes much more reliable. In a more recent reexamination of this topic, it was shown that honeybees indeed compute polarized light information during flight by extracting a heading angle from the perceived e-vectors (summarized in Figure 2). For these experiments, honeybees were trained to fly through a 12-m tunnel toward a feeder while experiencing either axial or perpendicular e-vectors from above (Evangelista, Kraft, Dacke, Labhart, & Srinivasan, 2014 Kraft, Evangelista, Dacke, Labhart, & Srinivasan, 2011). As expected, the resulting heading orientation encoded by the linear phase of the “waggle dances” differed by 90 degrees between the two experiments: an axial e-vector resulted in perpendicular dances (Figure 2A–C), while a perpendicular e-vector resulted in axial dance orientations (Figure 2D–F) (waggle dance directions are always mirror symmetric, due to the inherent directional ambiguity of a polarized light stimulus: 0 and 180 degrees are the same). These waggle dance orientations were in good agreement with the natural solar position under these conditions: perpendicular e-vectors in the zenith occur when the sun is behind or in front of the animal, and axial e-vectors in the sun coincide with solar positions toward +90 or −90 degrees. Interestingly, when the flight tunnel was divided into two consecutive halves of first axial and then perpendicular e-vectors, the resulting waggle dances pointed toward all four diagonal directions, which in fact is the sum of all possible directional vectors the animal could have deduced from summing the two orthogonal e-vector orientations it had experienced (Figure 2G–I). In the absence of a solar landmark, the honeybees therefore make correct decisions by using the celestial e-vector pattern, which heavily biases their navigational decisions.
Figure 3: Integration of sun and polarized light information by desert ants. (A) Classic example for path integration of the desert ant Cataglyphis : after an unbiased exploration run (blue), the ant returns to the nest in a straight line (red). To achieve this goal, the ant must constantly update information segments traveled to determine the homing vector to the nest. (With permission from Wehner, 2003.) (B) Summary of experiments where the ant was walking through a tunnel toward a feeder, while experiencing defined e-vector orientations that were created by mounting a polarizing foil onto the tunnel. When presenting the animals only one of three different stimuli during training and test (solar position, single e-vector, or celestial polarization pattern), they could correctly deduce solar position from e-vectors and vice versa. The animal therefore has an innate prediction of the relationship between these stimuli, and information can be transferred between the sensory systems perceiving them. Homing responses after training with perpendicular e-vectors is less accurate due to interference from phototaxis. (Adapted from Lebhardt & Ronacher, 2015.) (C) Top: By allowing the ant to see the sun while traveling through the tunnel, stimulus combinations were produced where polarization pattern and solar positions were in conflict. Bottom: The ants appeared to consider both stimuli equally reliable, setting of an intermediate homing vector (red) through averaging the vectors suggested by solar position (yellow), and polarization pattern (purple). (Adapted from Lebhardt & Ronacher, 2014.)
These experiments raise exciting questions about the integration of solar and celestial compass information, for instance, in cases of apparent conflict between a solar positions with an “unnatural” e-vector orientation. These questions have been addressed using another central place forager model system, the desert ant Cataglyphis . Over the last decades, the work of Rüdiger Wehner and colleagues has demonstrated the ability of these desert ants to efficiently navigate in an arid environment mostly void of any visual landmarks, thereby relying heavily on the celestial pattern of polarized light (for review, see Wehner, 2003). After finding food during an unbiased exploration run, the desert ants return to their nest entrance in a straight line rather than retracing their steps (which would result in an unnecessary waste of energy) (Figure 3A). In order to achieve this task, the ant is constantly updating its homing vector as it proceeds by measuring the direction of individual path segments as well as the distances traveled. This sophisticated navigational process, which was termed “path integration,” therefore combines the internal compass and the odometer system of the ant. Previous work investigating the ant’s internal compass system demonstrated that the celestial polarized light information appeared to dominate over the perceived solar position when the two were in conflict (Wehner & Müller, 2006). More recently, a series of experiments by Lebhardt and colleagues added new insights to the computation of these cues (Lebhardt, Koch, & Ronacher, 2012 Lebhardt & Ronacher, 2014, 2015): similar to the honeybee experiments, desert ants were trained to walk toward a feeder through a tunnel covered with polarization filters while the sun was occluded from the field of view. In the following test runs, the ants were walking under the open sky, witnessing the entire celestial e-vector pattern as well as the position of the sun. In agreement with the honeybee experiments, the ants trained under a single e-vector orientation correctly associated the solar position with the e-vector orientation, leading to strongly directed homing vectors: perpendicular e-vectors during training signaled solar positions front or back, whereas axial e-vectors signaled solar positions at orthogonal angles (Lebhardt et al., 2012). Once again, linear summation of orthogonal e-vectors during training resulted in diagonal homeward trajectories. Interestingly, it was also shown that polarized light information dominates over idiothetic cues (e.g., proprioceptive signals from mechanoreceptors), since the change of direction enforced by orthogonal bends in the tunnel were ignored by the ant if the presented e-vectors signaled an invariant celestial pattern. Another study by the same authors investigated the transfer of information between the solar detection system and the sensory system for detecting celestial polarization. For these experiments, desert ants experienced only one stimulus—either solar position or single e-vectors—during their training runs through the tunnel (Lebhardt & Ronacher, 2015). For the consecutive homing runs, the ants then had to extract the correct heading vector exclusively from the cue they were not trained with (the celestial polarization pattern or solar position) (Figure 3B). In all cases, the ants made correct navigational decisions. It appears therefore that the ant brain is capable of integrating directional information from solar and polarization detection systems, although they depend on different retinal detectors. Finally, Lebhardt and colleagues investigated the navigational decisions of Cataglyphis when solar and compass information were in direct conflict. In these experiments, desert ants performed training runs while presented single e-vectors as before, but the animals could also see the sun at the same time (Lebhardt & Ronacher, 2014). In all cases where the directional information provided by the e-vector and the solar compass disagreed during training, the ants chose an intermediate homing vector during the consecutive test under the open sky (Figure 3C). It appears therefore that inputs from both compass systems are summed roughly equally, which is surprising given previous findings on the dominance of the sky compass. It is possible that this choice of a mean direction applies to situations where both solar and sky compass information are considered equally reliable, resulting in their summation, rather than hierarchical override by one system (Wehner, Hoinsville, Cruse, & Cheng, 2016). It remains to be seen which factors determine dominance versus summation. In summary, both honeybees and desert ants can extract navigational information from detecting linearly polarized light or the solar position, and they have an innate understanding of the geographical relationship between these stimuli as they occur in nature.
Spontaneous Behavioral Responses to Polarized Light Guiding Orientation Behavior
Most insects do not travel back and forth between food sources and a fixed location like a nest or a hive. Therefore, reproducing the navigational decisions of these animal becomes more difficult. In this case, the investigation of polarized light vision relies on spontaneous behavioral responses, like the setting of certain compass headings, the alignment of the body axis with respect to the incident e-vector, an increased turning tendency in response to a rotating polarization filter, or keeping a straight course over time. All these spontaneous responses can be summarized as “polarotactic” behaviors. In all cases mentioned above, the animal might use the celestial polarization pattern as an orientation cue to plan and execute certain short- or long-range migratory tasks by avoiding to walk or fly in circles. Some of these experiments can be performed in the wild. Alternatively, polarotactic responses are characterized in laboratory experiments where an insect is tethered either over an air-suspended ball or on a wire in a flight arena. By turning the polarization filter above the animal, changes in the angular velocity and heading can be measured. For instance, polarotactic responses of walking crickets were described (Brunner & Labhart, 1985), and behavioral performance under different stimulus conditions were measured, by simulating decreasing degrees of polarization using quarter wave plate, which produces elliptically polarized light with varying ellipticity (and which is perceived as unpolarized in the case of circular polarization, since all e-vectors are equally represented over time Henze & Labhart, 2007). Interestingly, the behavioral response remained stable down to a (simulated) degree 5% polarization, after which the polarotactic response abruptly ceases, thereby demonstrating how stable this behavior is under natural conditions. The polarotactic responses of desert locusts in tethered flight were used in a similar way to demonstrate the importance of a prominent anatomical structure in the insect brain, the anterior optic tract (AOT), that connects the optic lobe with the anterior optic tubercle (AOTU), a more centrally located structure belonging to the optic glomeruli (Mappes & Homberg, 2004). Bilateral surgical lesion of the AOT completely abolishes polarotaxis in tethered desert locusts, thereby confirming its importance for polarization vision (Mappes & Homberg, 2007). One study on monarch butterflies suspended on a wire also found polarotactic responses (Reppert, Zhu, & White, 2004), which is important given that this species performs long distance migrations over several thousand kilometers across the North American continent, while another one disagreed (Stalleicken et al., 2005). Polarotactic responses are also common in flies ( Musca , Drosophila ), for which they were demonstrated both for single flies walking on a ball as well as for flies suspended in a flight arena (Vonphilipsborn & Labhart, 1990 Weir & Dickinson, 2012 Wolf, Gebhardt, Gademann, & Heisenberg. 1980). Additionally, using real-time computer tracking of walking populations of Drosophila further demonstrated their spontaneous tendency to align their body axis with the incident e-vector when no other visual stimuli were presented (Velez, Gohl, Clandinin, & Wernet, 2014 Velez, Wernet, Clark, & Clandinin, 2014 Wernet et al., 2012). In the latter case, polarotactic responses were described both for celestial stimuli as well as for linearly polarized stimuli presented to the ventral half of the eye.
Figure 4: Dung beetles set a bearing based on a snapshot of the visual scenery. Summary of three experiments investigating the rolling direction of the diurnal dung beetle Scarabaeus lamarcki . A specific combination of visual cues (celestial body or “Ersatz sun,” green polarization pattern, UV) was presented during initial dancing and/or ball rolling, followed by a second experiment with a different combination of stimuli. (A) When only the celestial body was presented during the first trial (dance and roll), polarized UV light did not orient the second run (right: circular plot of change in direction between consecutive runs is random). Hence, no internal matched filter exists for transferring orientation information between solar position and celestial polarization pattern. (B) Both cues were presented only during the first “dance” of the animal on top of the ball, yet only polarized UV light was presented during a second experiment (dance and roll). Clustering of changes in rolling direction around zero and 180 degrees suggests the position of the celestial body was remembered, albeit solar and antisolar orientation being indistinguishable, when using the polarized light pattern alone. (C) Only visual cues present during the first dance (in this case, the celestial body) can be used for orienting ball rolling during experiment 2. Taken together, the rolling performance of the dung beetle during experiment shows that a snapshot of the visual scenery can only be taken during the first dance period. (Adapted from el Jundi et al., 2016.)
Particularly powerful model systems for investigating polarotactic behaviors are nocturnal and diurnal dung beetles. In most cases, the dung beetle forms a ball of dung and rolls it away from the food source, usually following a straight line (Baird, Byrne, Scholtz, Warrant, & Dacke, 2010). To achieve this task, the beetles use a combination of visual cues. Amazingly, the nocturnal species Scarabaeus zambesianus uses the polarization pattern of the moonlit sky, which originates from reflection of sunlight off the moon and is one million times dimmer than the daylight pattern (Dacke, Nilsson, Scholtz, Byrne, & Warrant, 2003 Dacke, Nordstrom, & Scholtz, 2003). In addition, the beetle can also use the moon as a landmark to set a straight course (Dacke, Byrne, Scholtz, & Warrant, 2004), and even the Milky Way can be used as an orientation cue (Dacke, Baird, Byrne, Scholtz, & Warrant, 2013) in the case of Scarabaeus satyrus . Diurnal dung beetles like Scarabaeus lamarcki can use an even wider selection of visual cues for their orientation, including the solar position, spectral cues, intensity gradients, and the celestial polarization pattern, the latter being the dominant cue (Dacke, el Jundi, Smolka, Byrne, & Baird, 2014 el Jundi, Foster, Byrne, Baird, & Dacke, 2015 el Jundi, Smolka, Baird, Byrne, & Dacke, 2014). Interestingly, these beetles appear to ignore landmarks (Dacke, Byrne, Smolka, Warrant, & Baird, 2013). But how does the dung beetle process the mixture of visual cues? An important clue was provided by the beetle’s behavior: just prior to rolling the ball away from the dung pile, it climbs onto the ball and performs a characteristic “dance” during which it rotates around its vertical axis (Baird, Byrne, Smolka, Warrant, & Dacke, 2012). Interestingly, the dances also occur whenever the beetle loses control of the ball, after they lose contact with it, when deviations from the set course are induced, or when visual cues change. It was therefore proposed that the dance serves to establish a roll bearing and to return to this bearing once a disturbance has been experienced. One recent study tested this hypothesis, revealing the mechanism underlying these orientation dances (el Jundi et al., 2016). Using an experimental arena, in which artificial stimuli could be presented to the dung beetles (sun, polarization pattern, spectral cues), it was first revealed that unlike honeybees or desert ants, the dung beetle has no innate ability to correctly predict the natural relationship between these cues. Instead, it appears that the dance is used to take a “celestial snapshot” of the current stimulus ensemble, even if this combination of solar azimuth and e-vector orientation never occurs in a natural sky. This internal representation is then used to set a fixed bearing. Interestingly, the snapshot must be taken during the dance, since a specific visual cue that has been absent while dancing cannot be used for accurate ball rolling (Figure 4). This strikingly simple strategy serves as an efficient way to efficiently use available visual cues to walk in a straight line away from the dung pile where predators or competitors may aggregate.
The Detection of Polarized Reflections: Finding or Avoiding Water Surfaces?
When sunlight is reflected off a water surface, it becomes horizontally polarized, with a maximum degree of polarization (100%) at a particular angle of incidence known as Brewster’s angle (53° for an air/water interface). Some flying insects rely on it to identify bodies of water and many semi-aquatic and aquatic insects appear to detect water surfaces via polarized reflections (Wehner, 2001) (Figure 5A). In most cases, polarized reflections appear to be attractive, yet exceptions exist, for instance, flying locust swarms usually avoid flying over polarized surfaces, probably to avoid crashing into the sea—a fact that can be exploited to repel them (Shashar, Sabbah, & Aharoni, 2005). One classic example for insects attracted by polarized surfaces are dragonflies: male dragonflies approach polarized surfaces to establish an aquatic territory, whereas females will attempt oviposition on what they assume to be the water surface (Wildermuth, 1998). It has been proposed that different dragonfly species known to show a preference for dark versus bright ponds might use the degree of polarization to distinguish between these habitats, yet it remains to be shown how insects would be able to perform this computationally difficult task (Bernath, Szedenics, Wildermuth, & Horvath, 2002). Numerous accounts exist where female insects erroneously oviposit onto shiny surfaces they mistakenly take for water, like parked cars or black gravestones (Horvath, Bernath, & Molnar, 1998 Horvath, Malik, Kriska, & Wildermuth, 2007 Kriska, Horvath, & Andrikovics, 1998), while other insects are attracted by glass buildings (Kriska, Malik, Szivak, & Horvath, 2008). Many of the extremely short-lived mayflies (Ephemeropta) show strong attraction to water (Kriska, Bernath, & Horvath, 2007). The so-called compensatory upstream flights of female mayflies before oviposition appear to be guided by horizontally polarized reflections (Farkas et al., 2016). Interestingly, these important dispersion maneuvers are severely disrupted by unpolarized light pollution, like bridge lamps (Szaz et al., 2015). Another species for which the attraction to polarized surfaces has been demonstrated in great detail is the hemipteran back swimmer Notonecta glauca , a bug that spends a considerable portion of its life hanging under the water surface. However, during dispersal flights between water bodies, it also visually identifies water surfaces, resulting in a characteristic diving reaction (Figure 5B) (Schwind, 1984b). Providing horizontal platforms emitting linearly polarized UV light are sufficient to induce Notonecta ’s diving reaction (Schwind, 1983a). It should be pointed out that the glare resulting from horizontally polarized reflections can be problematic for some aquatic insects, for instance, when observing underwater objects from above the water surface, or for contrast enhancement while living near or under water (Wehner, 2001). Specific behavioral adaptations could therefore also aim at avoiding this stimulus (Sharkey, Partridge, & Roberts, 2015). Finally, it should also be noted that some insect species for which one would intuitively expect a strong attraction to linearly polarized reflections, based on their visual ecology centered around aquatic habitats, fail to display clear signs of such behaviors. For instance, so far there is no evidence that those honeybee workers with the dedicated task of water collection use polarized reflections to detect water sources. Also, it remains unclear whether female mosquitoes use vision at all for identifying oviposition sites after a blood meal, since they seem to rely mostly on olfactory cues (Bernath, Horvath, Gal, Fekete, & Meyer-Rochow, 2008 Bernath, Horvath, & Meyer-Rochow, 2012). In contrast, non-biting midges (Chironomidae) with a similar lifestyle appear to rely heavily on visual cues for the detection of water surfaces, thereby revealing important differences in the behavioral strategies used by these related species (Horvath, Mora, Bernath, & Kriska, 2011 Lerner et al., 2008). Taken together, a growing number of reports strengthens the importance of linearly polarized water reflections as an important environmental cue for many insect species. However, further quantitative characterization of these responses is needed for a better understanding of the different behaviors elicited by this stimulus.
Communication via Polarized Light, False Colors, and Circular Polarization
Water bodies are not the only source of linearly polarized reflections in nature. Similar reflections off shiny leaves represent an attractive oviposition cue for certain butterflies (Kelber, 1999a, 1999b). Interestingly, information about e-vector orientation and wavelength composition appear to be mixed in this case, resulting in the perception of “false colors.” This system most likely enables the female butterfly to distinguish matte from shiny leaves by perceiving them as different colors while flying by (Kelber, Thunell, & Arikawa, 2001). This cue would be suitable to signal several important features, such as quality of the landing site (leaf orientation), food quality (for caterpillar offspring), or protection for the eggs. A more recent study on the Japanese swallowtail butterfly Papilio xuthus reported similar mixing of linear polarization and light intensity (Kinoshita, Yamazato, & Arikawa, 2011). Hence, some butterflies might also perceive differently polarized surfaces of having different brightness levels. The authors propose that this mixing of visual modalities dominates during foraging, hence equipping the butterflies with two modes of polarization vision (mixing with color versus brightness) that may be used depending on the behavioral context of the animal (oviposition versus foraging).
Figure 5: Reflected polarized light as an important visual cue for insects. (A) Same visual scene photographed with (left) and without (right) a horizontally oriented UV polarizing filter. (With permission from Schwind, 1983a.) (B) The aquatic hemipteran Notonecta glauca identifies water bodies during flight, leading to a characteristic diving reaction. The visual fields of ventral photoreceptors scan the water surface, from which linearly polarized light is reflected most strongly at the angle θ (Brewster’s angle). (Adapted with permission from Schwind, 1984a.) (C) Photographs documenting the change in appearance of the scarab beetle Chrysina gloriosa when using a leftward (left) or rightward (right) circular polarization filters. (With permission from Sharma et al., 2009.)
True flies (Diptera) show strong attraction to polarized surfaces, but not necessarily water, which was demonstrated for blood-sucking horse flies (Tabanidae) (Egri et al., 2013 Horvath, Majer, Horvath, Szivak, & Kriska, 2008). This behavior most likely serves prey detection since polarimetric imaging of horses and cattle reveals strong linearly polarized reflections off their fur (Horvath et al., 2010). Brown and black fur produces the strongest polarized reflections, while white fur appears to be the best protection against horse fly attacks. Interestingly, certain fur patterns like stripes (zebras) and spots (cows) appear to change angle and degree of polarized reflections, thereby reducing attractiveness for horse flies and might therefore serve as an additional advantage of these otherwise useful camouflage-type adaptations (Blaho, Egri, Bahidszki, et al., 2012 Egri et al., 2012). A more sophisticated ability to detect polarized light patterns was revealed in a recent study where bumblebees were successfully trained to distinguish between different linearly polarized patterns. It appears therefore that pollinators may also use polarized reflections to identify or evaluate floral targets (Foster et al., 2014).
The iridescent scales on the wings of many butterflies also produce linearly polarized reflections that can be perceived by conspecifics and therefore serve as mating signals (Stavenga, Matsushita, Arikawa, Leertouwer, & Wilts, 2012 Sweeney, Jiggins, & Johnsen, 2003 Yoshioka & Kinoshita, 2007). For instance, certain Heliconius butterflies use these reflections to increase their visibility in the midst of highly complex visual forest environment where an abundance color, detail, and great differences in light intensity makes it difficult to be seen (Douglas, Cronin, Chiou, & Dominy, 2007). In this case, the polarized reflections are therefore used to increase the perceived visual contrast independent of spectrum and intensity. Surprisingly, the body cuticle of some insects reflects circularly polarized light—i.e., the e-vector rotates clock- or counterclockwise as the light beam propagates. For most insects, such a stimulus would appear unpolarized, since all e-vector orientations are equally represented in the beam of light (Labhart, 1996 Henze & Labhart, 2007). For instance, several scarab beetles reflect circularly polarized light (Hegedus, Szel, & Horvath, 2006 Jewell, Vukusic, & Roberts, 2007 Sharma, Crne, Park, & Srinivasarao, 2009). Usually well camouflaged in their natural habitat, these animals exhibit stark black contrast when viewed through a circular polarization filter (Figure 5C). In one behavioral study using choice experiments, specific phototactic responses to circularly polarized light were reported for the scarab beetle Chrysina gloriosa , whose body surface produces strong circularly polarized reflections (Brady & Cummings, 2010). Interestingly, the closely related species Chrysina woodii , which manifests only weak circularly polarized reflections, exhibited no phototactic discrimination between linearly and circularly polarized stimuli. It must be noted that another study investigating four different scarab beetle species manifesting circularly polarized reflections off their exocuticle found no evidence for specific behavioral responses to circularly polarized light (Blaho, Egri, Hegedus, et al., 2012). Hence, more studies are needed to evaluate the importance of circularly polarized reflections. Taken together, polarized reflections from a wide variety of substrates serve many different, yet very specific purposes, thereby greatly increasing the visual ecology of different insect species across a wide range of habitats.
All insect retinas are composed of a varying number of repetitive unit eyes, or ommatidia, which usually contain eight or nine photoreceptor neurons (Wernet, Perry, & Desplan, 2015). In many cases, specialized ommatidia containing photoreceptors with increased polarization sensitivity can be found in the dorsal periphery, or the ventral periphery of the retina. The neural circuits underlying these specialized ommatidia are being investigated in many insect species via the combination of anatomical, physiological, and behavioral tests.
Localized Polarization-Sensitive Photoreceptors in the Dorsal Rim Area
Despite considerable morphological variations, specialized ommatidia at the dorsal rim of the adult eye containing polarization-sensitive photoreceptors have been identified in many insect species (for review, see Labhart & Meyer, 1999). In this dorsal rim area (DRA), the light-gathering structures called rhabdomeres of at least two photoreceptor cells always contain straight, untwisted membrane invaginations, or microvilli. The photosensitive rhodopsin molecules are aligned along the axis of these rhabdomeric membranes, leading to a preferential absorption of a certain e-vector orientation by the photoreceptor cell, sometimes referred to as its “analyzer direction” (Roberts, Porter, & Cronin, 2011). Throughout the rest of the retina, polarization sensitivity of photoreceptors is actively prevented through twisting of the rhabdomere membranes in different directions along the length of the rhabdomere, leading to no preferred e-vector tuning of the receptors (and therefore low polarization sensitivity) (Wehner & Bernard, 1993). Usually, two groups of untwisted, highly polarization-sensitive photoreceptors can be found in every DRA ommatidium, manifesting analyzer directions that are orthogonal to each other (Labhart & Meyer, 1999). Hence, if one channel is maximally excited, the other one is minimally active. This design appears to be crucial for the perception of polarized light. In all known cases, the two groups of untwisted, polarization-sensitive photoreceptors with orthogonal analyzers express the same rhodopsin molecules in order to avoid mixing between color and polarized light information. However, the wavelength sensitivity of the rhodopsins used in DRA ommatidia varies between insects: while UV-sensitive rhodopsins are always used by bees, ants, and flies (Fortini & Rubin, 1990 Labhart, 1986 Vonhelversen & Edrich, 1974), crickets and locusts express blue-sensitive rhodopsins in the DRA (Henze, Dannenhauer, Kohler, Labhart, & Gesemann, 2012 Schmeling et al., 2014), while some beetles (cockchafers) rely on green-sensitive polarization-sensitive DRA units (Labhart, Meyer, & Schenker, 1992) (Figure 6A). Hence, the detection of the celestial polarization pattern occurs at different wavelengths, depending on the species, the reasons for which are still being discussed (Barta & Horvath, 2004 Hegedus, Horvath, & Horvath, 2006). Interestingly, there also exists a great variety in the number and subtypes of photoreceptor cells contributing to the two orthogonal analyzer channels in the DRA. For instance, flies use only two photoreceptors with orthogonal microvilli called R7 and R8 (Wunderer, Seifert, Pilstl, Lange, & Smola, 1990 Wunderer & Smola, 1982a), while the remaining six photoreceptors are not involved in celestial polarization vision (Wernet et al., 2012). In contrast, all nine photoreceptors in DRA ommatidia are polarization-sensitive in some butterfly species, like the monarch (Sauman et al., 2005). The reasons for this diversity remain unclear.
Figure 6: The anatomical basis for polarized light sensing in insects. (A) Top row: Four examples of polarization-sensitive ommatidia in the dorsal rim area (DRA) of crickets, beetles, honeybees, and flies (from left to right). Polarization-sensitive photoreceptors with untwisted rhabdomeres are shown in color. Different colors symbolize spectral sensitivities: green, blue, versus UV. Bottom row: morphology of ommatidia outside the DRA, for comparison. (Adapted with permission from Labhart & Meyer, 1999.) (B) Morphology of locust transmedullary neurons (shown in orange, via Dextran tracer application to the lower unit of the anterior optic tubercle), which are presumed to be specifically post-synaptic to DRA photoreceptors (shown in blue). Overlap between photoreceptors and transmedullary neurons is obvious in the dorsal rim medulla (DRMe). (Adapted from el Jundi, Pfeiffer, & Homberg, 2011.)
Across insect species, the orthogonal analyzer directions differ between neighboring DRA ommatidia, resulting in a fan-shaped array of analyzers, which together can detect any celestial e-vector orientation (Blum & Labhart, 2000 Homberg & Paech, 2002 Strausfeld & Wunderer, 1985 Weir et al., 2016). Due to the low light intensities at night, the number of DRA ommatidia in nocturnal dung beetles is increased when compared to their diurnal counterparts, whereas true flies set aside only one single row of DRA ommatidia for polarization vision, along the dorsal head cuticle (Dacke, Nodrstrom, Scholtz, & Warrant, 2002 Smolka et al., 2016). Interestingly, DRA ommatidia of some insect species can be identified with the naked eye due to darker pigmentation (Homberg & Paech, 2002). In some cases, corneal specializations serve to severely widen the optical axis of the photoreceptors, thereby allowing the DRA to cover a large portion of the sky at the expense of acuity in this part of the eye (Aepli, Labhart, & Meyer, 1985 Meyer & Labhart, 1981). Interestingly, there always exists a sharp boundary between morphologically specialized DRA and non-DRA ommatidia, rather than a gradual change between the two morphological types (Labhart & Meyer, 1999). In Drosophila , the molecular mechanisms governing this localized specification of DRA ommatidia have been described in detail: specific expression of the homeodomain transcription factor Homothorax (Hth) in DRA R7 and R8 cells is both necessary and sufficient to induce the DRA fate (Wernet & Desplan, 2014 Wernet et al., 2003) Hth is expressed in response to the morphogen Wingless (Wg) and dorsal selector genes of the Iro-C complex. Interestingly, Wg levels specify different cell types depending on their proximity to the dorsal head cuticle (Tomlinson, 2003), and secondary release of Wg is used to sharpen the boundary between the different ommatidial fates (Kumar, Patel, & Tomlinson, 2015). To date, it is not known whether this molecular mechanism is conserved beyond flies.
Morphological Specializations Leading to Polarization-Sensitivity in the Ventral Retina
There does not appear to exist a specialized type of ommatidia, common to all insects, which is localized at the ventral rim of the retina, harboring polarization-sensitive photoreceptors that could mediate the response to linearly polarized reflections, for example, from shiny surfaces. Despite the numerous reports of behavioral responses to such stimuli, it remains surprising that only a handful of neural correlates for such behaviors have so far been characterized for different insect eyes. Probably the best-characterized example is the zonation of the ventral retina of the hemipteran back swimmer Notonecta glauca . The identification of different ventral zones of ommatidia harboring untwisted photoreceptors of differing microvilli orientations represents an ideal adaptation to its very specialized aquatic lifestyle above and below the water surface. The most ventrally facing ommatidia containing two fused, untwisted rhabdomeres with orthogonally oriented microvilli represent a structure perfectly adapted for detecting polarized reflections like water surfaces (Schwind, 1983b). Interestingly, such specialized adaptations have not yet been described in such detail for any other insect. Other retinal substrates possibly related to polarized reflections off water have been described in another hemipteran, the water strider Gerris lacustris , the sunburst diving beetle larvae Thermonectus marmoratus , as well as for some fly species (Schneider & Langer, 1969 Stecher, Morgan, & Buschbeck, 2010 Trujillocenoz & Bernard, 1972). In Gerris , ventral ommatidia contain untwisted rhabdomeres, often with only one predominant analyzer direction. This could serve to filter out polarized reflections, most likely resulting in the animal’s improved ability to look deeper into the water, or to increase contrast when observing animals against the glare that results from polarized reflections (Schneider & Langer, 1969). Hence, retinal specializations in the ventral retina that are related to polarized reflections may serve opposite functions, depending on their ultrastructure and the lifestyle of the insect: attraction to water via detection of horizontal e-vectors or the specific screening of such surface-reflected light.
Reflections off leaves or off the body surface of other animals provide important cues for many insects, either for oviposition, intraspecific communication, or to detect prey. For instance, the “false color” detection system of the Australian orchard butterfly Papilio aegeus results from blue- and green-sensitive photoreceptors outside the DRA retaining polarization sensitivity due to insufficient rhabdomere twist (Arikawa & Uchiyama, 1996). More importantly, their analyzer directions are orthogonal to each other (vertical versus horizontal, respectively). Upon excitation of any of these photoreceptors, information about e-vector orientation and wavelength composition will therefore be mixed, resulting in the perception of “false colors.” Another well-characterized example for the existence of polarization-sensitive photoreceptors outside the DRA is from electron microscopy–based ultrastructure of blood-sucking horse flies (Tabanidae). These studies revealed that in the midregion of the retina, both R7 and R8 cell rhabdomeres are largely untwisted, providing a putative retinal substrate for host finding via the detection of polarized light (Smith & Butler, 1991 Wunderer & Smola, 1986). Interestingly, very similar studies also identified a subtype of untwisted R8 photoreceptor in blow flies, yet such cells could not be reliably identified in closely related Drosophila (Wunderer & Smola, 1982b). Instead, systematic analysis of rhabdomere twist in this species revealed a low number of untwisted, UV-sensitive R7 cells in the ventral fly retina (Wernet et al., 2012). Together with low twisting outer photoreceptors within the same ommatidia, these cells could provide the retinal substrate for Drosophila ’s polarotactic responses to linearly polarized stimuli presented ventrally (Velez, Gohl et al., 2014 Velez, Wernet, et al., 2014 Wernet et al., 2012 Wolf et al., 1980).
The detection of circularly polarized light reflected off the exoskeleton of some scarab beetles also remains incompletely understood. The expected anatomical design of a hypothetical retinal detector has been proposed based on stomatopod crustacean examples: in their eyes, two orthogonally arranged detectors are joined by a third distal photoreceptor with a microvilli orientation of 45 degrees, which acts a quarter wave retarder (Chiou et al., 2008 Warrant, 2010). Nevertheless, such specializations have not yet been described in scarab beetles, whose body surfaces cause circularly polarized reflections, nor in any other insects.
Anatomical Characterization of Circuit Elements for Polarization Vision in the Insect Brain
Since the DRA photoreceptor input channels for celestial polarization vision can be identified unambiguously, their post-synaptic elements have also been the subject of several studies. Although electrophysiology has proven to be a much more effective technique for describing polarization-sensitive circuit elements throughout the insect brain, different anatomical techniques have also been used successfully to describe cell types that seem to specifically contact DRA photoreceptors. For instance, the injection of tracer molecules such as biotinylated dextran into the lower unit of the anterior optic tubercle (AOTU-LU, or AOTU-LUC in honeybees) in both locusts and bumblebees revealed transmedullary neurons with processes in the Dorsal Rim Medulla (MEDRA) and long projections into the AOTU-LU (Homberg, Hofer, Pfeiffer, & Gebhardt, 2003 Pfeiffer & Kinoshita, 2012 Zeller et al., 2015). Co-labeling of DRA photoreceptor terminals with another dextran probe confirmed very close contact between these cells (el Jundi, Pfeiffer, & Homberg, 2011) (Figure 6B). These transmedullary neurons therefore serve as excellent candidates for DRA input into the anterior optic tubercle that appear to be conserved between species. Another study using immunohistochemichal labeling of Cryptochrome-expressing cells in the monarch butterfly brain pointed toward connections between DRA photoreceptor terminals and the circadian clock (Sauman et al., 2005). Finally, cobalt injections into marginal ommatidia of the blowfly revealed columnar small field neurons connecting the MEDRA with distinct marginal regions of the deeper neuropils (lobula and lobula plate). From there, local assemblies of columnar neurons then connect to the same region in the optic lobe of the opposite eye, as well as to descending neurons terminating in the thoratic ganglia (Strausfeld & Wunderer, 1985).
Of course these morphological studies did not provide functional proof that the characterized cell types are involved in polarization vision, nor did they directly show that synaptic connections between them exist. Nevertheless, they provide the framework of a neural circuit for celestial polarization vision that includes the DRA, the AOTU, as well as central processing centers whose cellular organization has been described in detail using electrophysiology (see next section).
Functional methods for revealing the activity of identified neurons are crucial for understanding their role in the perception of specific visual stimuli. Since several excellent review articles on this topic already exist, this section only briefly introduces the characterization of polarization vision circuitry in insects using electrophysiology. The functional characterization of polarization-sensitive circuit elements includes photoreceptors, optic lobe cell types, and neurons in the central brain.
Polarization Sensitivity of Insect Photoreceptors
Figure 7: Functionally characterized polarization-sensitive neurons in the insect brain. (A) Modulation of activity from polarization-sensitive photoreceptors in the Drosophila DRA expressing the genetically encoded calcium indicator GCamp6. Top: Orientation of the polarizer maximum response (preferred e-vector angle, in pseudocolor) in R7 and R8 terminals. Bottom: Preferred e-vector angle plotted against horizontal position (trend line shown in black). Responses R7 and R8 at any given position along the fan-shaped array of analyzers are approximately orthogonal of each other, in good agreement with photoreceptor morphology (rhabdomere orientation). (With permission from Weir et al., 2016.) (B) Fluorescence traces for R7 and R8 regions of interests during one full rotation of the polarizing filter. Note the negative signals of both photoreceptor at e-vector orientations when the orthogonal channel is maximally excited. (With permission from Weir et al., 2016.) (C) Two models for polarization processing in a medullar column receiving input from R7 and R8 from the same ommatidium. All connections between cells are inhibitory (R7 and R8 photoreceptors are histaminergic). Out = output neuron Int = sign-inverting interneuron (Int). (With permission from Weir et al., 2016.) (D) Summary of the gradually changing compass-like representation of e-vector tunings (double arrows) in columnar CPU1 neurons of the protocerebral bridge and upper division of the central body (CBU) of the locust. LAL = lateral accessory lobe. (With permission from Homberg, 2015.) (E) Activity of a central body tangential neuron (lower division) with stimulation from above through a rotating polarizer. Spike frequency is modulated as a function of e-vector orientation (note how Φ max and Φ min spiking activity occur at orthogonal E-vectors). (With permission from Homberg, 2015.) (F) Anatomy of the polarization vision pathway in the locust, as deduced from single-unit electrophysiology recordings. Highlighted are the brain regions involved in processing sky compass cues (red), with an emphasis in the central body (gold). Abbreviations: La = lamina LoX = lobula-complex aMe = accessory medulla DRLa = dorsal rim lamina DRMe = dorsal rim medulla MB = mushroom body AL = antennal lobe POTu = posterior optic tubercle BU = bulb ALo = anterior lobula PB = protocerebral bridge CBU = central body upper unit CBL = central body lower unit LAL = lateral accessory lobes. (With permission from el Jundi, Pfeiffer, et al., 2014.)
The high polarization sensitivity (PS) of photoreceptors in the DRA has been confirmed for many insect species using electrophysiology (Hardie, 1984 Labhart, 1980, 1986 Labhart, Hodel, & Valenzuela, 1984 Stalleicken, Labhart, & Mouritsen, 2006). In most species, photoreceptors outside the DRA have significantly low PS due to rhabdomere twist, yet exceptions exist (Bandai, Arikawa, & Eguchi, 1992). These experiments also confirmed the direct link between morphology and physiology of DRA input channels resulting in their orthogonal analyzer directions. They also unambiguously revealed the spectral sensitivity of the respective photoreceptors. However, in some species, photoreceptors are too small for electrophysiology, for instance, in Drosophila . Due to the molecular genetic toolkit available for this species, genetically encoded indicators of activity have become an attractive alternative to electrophysiology. One recent study used the genetically encoded calcium sensor GCamp6 to visualize the activity of DRA inner photoreceptors R7 and R8 in response to a polarizer illuminated by a UV LED (Weir et al., 2016). One attractive advantage of this technique is the possibility to simultaneously record the activity from many R7 and R8 terminals. The fan-shaped array of analyzer directions across the DRA was thus visualized, as well as the strictly orthogonal alignment of R7 versus R8 within the same ommatidium (Figure 7A). Surprisingly, this study also revealed clear direct inhibitory interactions between R7 and R8, which had previously been reported only from few electrophysiological recordings in the DRA of larger flies (Hardie, 1984) (Figure 7B). This inhibition could result from direct synaptic contacts between R7 and R8 photoreceptors. The existence of such inter-photoreceptor synapses was recently confirmed via 3D electron microscopic reconstruction of the Drosophila optic lobes (Takemura et al., 2013 Takemura, Lu, & Meinertzhagen, 2008). Hence, these reciprocal synaptic connections could increase polarization contrast at the level of R7 and R8 photoreceptors themselves (Figure 7C). It remains to be seen whether this retinal microcircuitry is a specialized adaptation of flies or whether it represents a broader concept present across insect species.
Electrophysiological Characterization of Circuit Elements in the Insect Optic Lobes
Insect photoreceptors can be grouped into two categories, depending on the length of their axons. Most have short visual fibers (svf) projecting into a neuropil called lamina, where usually one or two photoreceptors with long visual fibers (lvf) terminate in a deeper neuropil called the medulla. Together with the lobula complex, lamina and medulla form the optic lobes of insects. Several polarization-sensitive neuron types have been characterized by electrophysiology in the medulla of different insect species. One example from crickets are the so-called polarization-opponent (POL) interneurons, which are excited by one particular e-vector orientation while being inhibited by an orthogonal e-vector (Labhart, 1988). Such opponent microcircuitry is beneficial in several ways, since it makes the system insensitive to intensity modulations and therefore to celestial intensity gradients, but it also increases the amplitude of signal modulation, thereby increasing polarization contrast. To date, it is not known whether POL neurons receive direct input from photoreceptor cells, which, at least in Drosophila , could produce very similar signals. However, it was shown via recordings from cricket POL neurons that DRA ommatidia with similar analyzer directions but located in different regions of the DRA converge onto the same post-synaptic elements (Labhart, Petzold, & Helbling, 2001). Interestingly, only three groups of POL neurons with distinct analyzer preference were identified in crickets, leading to a model in which the relative excitation of these three population could be used to encode a specific compass heading (Labhart & Wehner, 2006). In locusts, one recent study characterized several polarization-sensitive neurons in the medulla, as potential neuronal elements intercalated between the DRA and AOTU, whose connectivity remains unclear: five cell types were described with wide arborizations in the same medulla layer as well as in the MEDRA and the “accessory medulla,” the presumed circadian clock of the locust (el Jundi et al., 2011). Such a potential interaction between the polarization-vision and circadian systems is particularly interesting, due to the fact that the celestial polarization pattern changes in a predictable way as the solar stimulus moves across the sky together with the solar cue. Similar to cricket POL neurons, the medulla neurons in the locust showed cell type-specific orientation tuning to zenithal polarized light, albeit without the characteristic binning into three specific categories (el Jundi et al., 2011). Furthermore, the polarization-sensitive medulla neurons from locusts did not show any color opponency or daytime specific adjustment of sky compass signals. Such characteristics are important for correcting for the change in solar elevation and have been demonstrated for neurons in the AOTU of the locust and therefore arise at a later processing stage, beyond the medulla (Homberg, 2004 Homberg, Heinze, Pfeiffer, Kinoshita, & el Jundi, 2011 Kinoshita et al., 2007 Pfeiffer & Homberg, 2007). Hence, these neuronal cell types represent good candidates for the neuronal substrate that integrates time-compensated sky compass information in a migratory insect. Despite their precise characterization, the exact synaptic connectivity between these individual elements, as well as the DRA and AOTU, remains to be worked out.
Electrophysiological Characterization of Circuit Elements in the Insect Central Brain
A major step toward understanding how discrete compass headings are extracted from the celestial pattern of polarized light was the description of polarization-sensitive neurons in a neural structure buried deep in the brain of the locust, called the central complex. In a conserved substructure known as the “protocerebral bridge,” the preferred e-vector orientation of polarization-sensitive neurons varies systematically from one columnar structure to the next, leading to a topographic representation of celestial e-vectors deep in the brain (Heinze & Homberg, 2007) (Figure 7D,E). A defined compass heading could therefore be encoded by the relative activity of the “compass neurons” forming this internal map. In the meantime, central complex neurons with very similar properties have been identified in crickets, monarch butterflies, and dung beetles (el Jundi, Warrant, et al., 2015 Heinze & Reppert, 2011 Sakura, Lambrinos, & Labhart, 2006). Interestingly, these central complex neurons also respond to unpolarized chromatic stimuli (which are not detected by DRA ommatidia), thereby acting as an integrator of different celestial cues (for review, see el Jundi, Pfeiffer, Heinze, & Homberg, 2014). Recently, one study comparing responses of compass neurons between nocturnal and diurnal dung beetle species shed more light on this integration: on the one hand, the compass neurons from diurnal beetles always show responses when an artificial celestial body is presented under experimental conditions. On the other hand, in the nocturnal species, compass neurons usually respond to the pattern of polarized skylight during night time, which would be the most reliable navigational cue in the night sky. However, when the nocturnal beetle is forced to roll its ball during the day, its compass neurons undergo a switch by responding to the solar position (el Jundi, Warrant, et al., 2015). These findings correlate well with the behavioral decisions of these animals when they switch to using the sun as a more reliable orientation cue when forced to roll their ball during the day. Hence, there exists a good correlation between visual cue preference (and reliability) during ball-rolling behavior and the weighing of different visual inputs in the central complex of these insects. Taken together, the use of different physiological techniques across species has resulted in a growing circuit diagram for polarization vision in insects (Homberg, 2015) (Figure 7F). While the details are being worked out, it remains interesting to see what similarities and differences exist between these circuits and those mediating visual responses to other cues, like motion and color.
Not visible to the human eye, polarized light represents an important stimulus for many insects, including such ethologically important cues like linearly polarized skylight, linearly polarized reflections, and potentially also circularly polarized light. The study of navigating insects over the last few decades has resulted in a good understanding of their visual ecology, the underlying anatomical structures and the functional characterization of underlying circuit elements. Much less is known about the anatomical and physiological substrates for the detection of linearly polarized reflections. Insect models that rely on polarized reflections for tasks like oviposition, prey detection, or intraspecific communication should therefore become a priority. Just as for the detection of other visual cues like motion or color, understanding the functional connectivity of the vast number of neuronal elements involved in any of these processes remains a challenge. Further high-resolution reconstruction efforts are needed to reveal the exact synaptic connectivity between the described neuronal units. More sophisticated imaging techniques in combination with genetically encoded indicators of activity may also provide crucial information about sensory processing by visualizing many cells at once. Introducing these tools into non-genetic model organisms like locusts, dung beetles, or butterflies has now become possible, thanks to improved genome editing techniques like Crispr/Cas9 (Perry et al., 2016). Taken together, polarization vision serves a good example for a research field that successfully combines behavioral, anatomical, and physiological studies for understanding the relevance of certain visual cues to different animal species. It therefore combines the overlapping perspectives of such exciting fields as visual ecology, neuroethology, and modern circuit science.
The authors thank Fleur Lebhardt, Claude Desplan, and one anonymous reviewer for helpful comments on the manuscript’s text and figures. The authors also apologize for any work that could not be cited due to restrictions in length.
Catherine A. Hill and John F. MacDonald, Department of Entomology
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Horse and deer flies are annoying biting pests of wildlife, livestock, and humans. Their blood sucking habits also raise concerns about possible transmission of disease agents. You are encouraged to learn more about the biology of horse and deer flies to avoid being bitten and to understand the public health risk posed by these insects.
Are Horse and Deer Flies Public Health Risks?
The bites of female horse and deer flies are painful and, if numerous enough, can disrupt recreational activities and even the harvesting of some agricultural crops. Their mouthparts include two pairs of cutting “blades” that lacerate skin and cause flow of blood out of the wound, which females lap up with a sponge-like mouthpart. Males have similar, but much weaker mouthparts. They are not capable of biting and do not feed on blood.
The blood sucking behavior of females together with their possible role in the transmission of disease agents have been studied extensively. Numerous viruses, bacteria, and protozoa have been isolated from the bloody, sponge-like mouthpart of females and from their digestive system, but there are no studies showing conclusively that they are capable of transmitting disease agents to humans, with one exception. There is evidence that a deer fly in the western U.S. is involved in the transmission of a bacterium that causes the disease “tularemia,” which also is known as “deer fly fever” and “rabbit fever.” The role of deer flies in transmission is minor, however, compared to transmission by ticks and via contact with infected small game animals, especially rabbits.
How Many Types of Horse and Deer Flies Are There?
Horse and deer flies are “true” flies in the insect Order Diptera, and comprise the Family Tabanidae known as “tabanid flies” or “tabanids.” There are an estimated 4, 300 species of horse and deer flies in the world, approximately 335 of which occur in the continental U. S. Of these, over 160 species are horse flies, and over 110 species are deer flies. It is estimated that at least 45 species of horse flies and 30 species of deer flies occur in Indiana. The vast majority of horse flies are in two genera, Tabanus and Hybomitra. Nearly all deer flies are in the genus Chrysops.
How Can I Recognize a Horse Fly or Deer Fly?
Adult horse flies (Fig. 1) and deer flies (Fig. 2) are relatively large to very large (approximately 0.25 to 1.25 inches long), robust flies with a pair of huge eyes known as “compound eyes.” Those of some horse flies have colorful purple or green bands against a blue or yellowish-green background. The mouthparts are large and prominent, projecting downward and forward in front of the head. They have large, fan-shaped wings and are capable of rapid flight and flying long distances.
Figure 1. A horsefly, Tabanus sp. (Diptera: Tabanidae), adult female. (Photo by: Drees, Univ. of Texas)
Figure 2. A deer fly, Chrysops sp. (Diptera: Tabanidae), adult female. (Photo by: Drees, Univ. of Texas)
What Is the Life Cycle of Horse and Deer Flies in Indiana?
Similar to all flies, horse and deer flies develop from egg to adult via a process of “complete metamorphosis.” This means the last larval stage passes through a non-feeding pupal stage, from which the adult eventually emerges.
The summarized life cycle of horse flies (Fig. 3) and deer flies (Fig. 4) begins with the emergence of adults from late spring into summer, depending on the species. Upon becoming active, adults of both sexes feed on energy-rich sugars in nectar, plant sap, or honey dew produced by sap-sucking insects such as aphids and scale insects. Mating of the few species that have been observed takes place in flight. Females of some species are capable of developing an initial batch of eggs without taking a blood meal, otherwise blood is required for the development of eggs. Females search for a place to lay a single mass of eggs consisting of 100-800 eggs, depending on species. Egg masses of most species that have been studied are laid on the underside of leaves or along the stems of emergent vegetation growing in wetlands. Hatching occurs in approximately 2-3 days, and newly emerged larvae drop down into water or saturated soil in which they feed and develop.
Figure 3. Summarized life cycle of horse flies. (Drawing credit: Scott Charlesworth, Purdue University, based in part on Pechuman, L.L. and H.J. Teskey, 1981, IN: Manual of Nearctic Diptera, Volume 1)
Figure 4. Summarized life cycle of deer flies. (Drawing credit: Scott Charlesworth, Purdue University, based in part on Pechuman, L.L. and H.J. Teskey, 1981, IN: Manual of Nearctic Diptera, Volume 1
The sites in which horse and deer fly larvae develop are known for only about a third of the species in the U. S. Deer fly larvae appear to be limited to aquatic habitats, including marshes, ponds, and streams. Developmental sites of horse fly larvae are more varied. Larvae of most species are found in freshwater and saltwater marshes, some in streams, some in moist forest soils, and a few in moist decomposing wood. Larvae of all species of horse flies that have been studied are predators. They feed primarily on other soft-bodied animals such as insect larvae and worms, but larvae of some large species of horse flies feed on small vertebrates, including tadpoles, frogs, and toads. Horse fly larvae appear to possess a toxin in their saliva that is involved in subduing their prey. Much less is known about the feeding behavior of deer fly larvae, and there is no consensus as to whether they are predators or scavengers.
The larval stages of horse and deer flies range in number from 6-13. The last larval stage passes winter in the site in which it developed and molts into a pupa the following spring. Most species complete one generation per year. However, small species of deer flies can complete 2-3 generations per year and very large species of horse flies require 2-3 years in which to complete larval development.
What Should I Know About the Feeding Behavior of Adult Horse and Deer Flies?
Only females take a blood meal, and, with rare exception, they feed during the daytime. Unlike numerous other groups of blood sucking flies, female horse and deer flies do not enter structures and thus do not feed on humans indoors. Female horse flies feed primarily on large mammals, including stationary hosts, and they typically bite the legs and body, rarely on the head. Although there are species of horse flies that feed on humans, Indiana species rarely do. In contrast to horse flies, female deer flies typically feed on moving hosts and usually bite on the shoulders and head. They have a wide host range, attacking mammals of all sizes, including humans, and some species feed on birds and reptiles. Females of both horse and deer flies are aggressive, persistent feeders that quickly return to bite again if they are interrupted before they take a complete blood meal.
Similar to other blood sucking insects, female horse and deer flies respond to chemical and visual cues associated with a potential host. Carbon dioxide given off by warm-blooded animals provides a long-range cue, attracting females into the vicinity of a host. There, visual cues such as motion, size, shape, and dark color serve as attractants. Female horse and deer flies are deterred very little by repellents, including DEET, and humans entering infested areas have little protection against them.
How Do Humans Influence Horse and Deer Fly Development?
Humans generally do not influence horse and deer fly development because habitats that support larval development are “natural,” including freshwater wetlands, saltwater marshes, and open areas within forests. However, there is one type of habitat associated with human activity that can be a source of horse flies. Larvae and pupae of a few species are able to complete development in low areas of pastures or cultivated fields that support standing water or at least consist of heavily saturated soils.
Are There Effective Methods of Controlling Horse and Deer Flies?
Controlling horse and deer flies is nearly impossible. The use of insecticides to kill larvae is not an option because the vast majority of species develop in natural habitats in which insecticides cannot be applied due to environmental concerns. Even if they could be used, insecticides would be ineffective in controlling larvae because they are widely dispersed in a developmental site. The use of insecticides against adult horse and deer flies is not a realistic option because they are relatively large to very large and unaffected by the rate of insecticide that can be applied according to product label. At best, an insecticide application aimed at adults might produce a minor and temporary reduction in biting. A number of trapping devices have been used to capture adults, but their value is limited to sampling. At best, trapping devices produce temporary, minor relief from female horse flies.
Again, repellents, including those containing DEET, have very little or no effect in deterring adult horse and deer flies. Wearing a thick long sleeve shirt, thick pants, and a heavy hat may provide some protection against bites when entering habitats that support large numbers of adult horse and deer flies, but females can be very annoying as they attempt to take blood meals.
Where Can I Find More Information About Horse and Deer Flies?
There is surprisingly little information about horse and deer flies on university and governmental Web sites. There is, however, a recent textbook (2002) by G. Mullen and L. Durden, Medical and Veterinary Entomology, that includes an excellent chapter devoted to horse and deer flies, covering biology, behavior, and medical and veterinary risk. It also includes a section that evaluates various methods used in attempts to control horse and deer flies.
READ AND FOLLOW ALL LABEL INSTRUCTIONS. THIS INCLUDES DIRECTIONS FOR USE, PRECAUTIONARY STATEMENTS (HAZARDS TO HUMANS, DOMESTIC ANIMALS, AND ENDANGERED SPECIES), ENVIRONMENTAL HAZARDS, RATES OF APPLICATION, NUMBER OF APPLICATIONS, REENTRY INTERVALS, HARVEST RESTRICTIONS, STORAGE AND DISPOSAL, AND ANY SPECIFIC WARNINGS AND/OR PRECAUTIONS FOR SAFE HANDLING OF THE PESTICIDE.
It is the policy of the Purdue University Cooperative Extension Service that all persons have equal opportunity and access to its educational programs, services, activities, and facilities without regard to race, religion, color, sex, age, national origin or ancestry, marital status, parental status, sexual orientation, disability or status as a veteran. Purdue University is an Affirmative Action institution. This material may be available in alternative formats.
This work is supported in part by Extension Implementation Grant 2017-70006-27140/ IND011460G4-1013877 from the USDA National Institute of Food and Agriculture.
Can Biting Insects Detect Specific Places On A Host Body? - Biology
Biting midges can be a nuisance to campers, fishermen, hunters, hikers, gardeners, and others who spend time outdoors during early morning and evenings, and even during the daytime on cloudy days when winds are calm. They will readily bite humans the bites are irritating, painful, and can cause long-lasting painful lesions for some people.
A common observation upon experiencing a bite from this insect is that something is biting, but the person suffering cannot see what it is. Biting midges are sometimes incorrectly referred to as sand flies. Sand flies are insects that belong to a different biological group and should not be confused with the biting midges.
Figure 1. Culicoides furens shown next to a U.S. dime and pencil point to demonstrate the relative size of this adult biting midge species. Photograph by Roxanne Connelly, Florida Medical Entomology Laboratory, University of Florida.
Distribution (Back to Top)
There are over 4,000 species of biting midges in the Ceratopogonidae family, and over 1,000 in just one genus, Culicoides. The distribution of midges in the genus Culicoides is world-wide 47 species are known to occur in Florida. Species belonging to the genus Leptoconops occur in the tropics, sub-tropics, the Caribbean, and some coastal areas of southeast Florida.
The natural habitats of biting midges vary by species. Areas with substantial salt marsh habitat are major producers of many biting midge species. Additional sources for some species, like the bluetongue virus vector Culicoides sonorensis Wirth and Jones, include highly organic soil that is wet but not underwater such as those found with high manure loads in swine-, sheep- and cattle-farming operations. These insects do not establish inside homes, apartments, or inside humans or other animals.
Description (Back to Top)
Immature Stages: The eggs can be cigar-, banana-, or sausage-shaped and approximately 0.25 mm long. They are white when first laid but later turn brown or black. The eggs are laid on moist soil and cannot withstand drying out. Some species can lay up to 450 eggs per batch and as many as seven batches in a lifespan. Eggs typically hatch within two to 10 days of being laid time to hatch is dependent on the species and temperatures.
The larvae are worm-like, creamy white, and approximately 2 to 5 mm long. Larvae develop through four instars the first instar larvae possess a functional spine-bearing proleg. Pupal color can be pale yellow to light brown to dark brown. They are 2 to 5 mm in length with an unsegmented cephalothorax that has a pair of respiratory horns that may bear spines or wrinkles. During this stage, the insects possess a spiny integument which can be used to identify the fly to species level.
Adults: The adult no-see-ums are gray and less than 1/8-inch long. The two wings possess dense hairs and give rise to pigmentation patterns. These wing patterns are used by biologists to identify species. The large compound eyes are more or less contiguous above the bases of the 15-segmented antennae. The pedicel of the males' antennae houses the Johnston's organ. The mouthparts are well-developed with cutting teeth on elongated mandibles in the proboscis, adapted for blood-sucking in females, but not in males. The thorax extends slightly over the head, and the abdomen is nine-segmented and tapered at the end.
Figure 2. Adult biting midge, Culicoides sonorensis Wirth and Jones, showing blood-filled abdomen and the characteristic wings patterns used for species identification. Photograph by Ed T. Schmidtmann, USDA/ARS.
Life Cycle (Back to Top)
Adults: Biting midges are holometabolous, progressing from egg to larva to pupa, and finally to the adult stage. The complete cycle can occur in two to six weeks, but is dependent on the species and environmental conditions. The adults are most abundant near productive breeding sites, but will disperse to mate and to feed. The mean distance for female flight is 2 km, less than half of that distance for males.
Male Culicoides typically emerge before the females and are ready to mate when the female emerges from the pupal stage. Mating typically occurs in flight when females fly into swarms of males and the insects are oriented end to end with the ventral parts of the genitalia in contact. Some species mate without swarming instead, the males go to hosts where the female is likely to feed on blood mating occurs when she finishes feeding.
Eggs: Males and females feed on nectar, but the females require blood for their eggs to mature. The females will blood-feed primarily around dawn and dusk however, there are some species that prefer to feed during the day. Some species are autogenous and therefore may produce the first batch of viable eggs without a blood meal using reserves stored from the larval period blood meals are required for subsequent batches of eggs.
The number of eggs produced varies among species and size of bloodmeal. For example, Culicoides furens (Poey) can lay 50 to 110 eggs per bloodmeal, and C. mississippiensis Hoffman, 25 to 50 eggs per bloodmeal. The adults can live two to seven weeks in a laboratory setting, but only a few weeks under natural conditions.
Larvae: Larvae require water, air and food and are not strictly aquatic or terrestrial. They cannot develop without moisture. The larvae are present in and around salt-marsh and mangrove swamps, on shores of streams and ponds, and in muddy substrates. They feed on small organisms. Most species cannot exist more than a few inches below the air-water interface.
In the tropics, the larval habitat of many species is in rotting fruit, bromeliads, and other water-holding plants. Other larval habitats include mud, sand, and debris at edges of ponds, lakes and springs, tree holes, and slime-covered bark. The larval stage can last from two weeks to a year, depending on the species, temperatures, and geographic area.
While some larvae can develop in wet manure-contaminated areas (Mullen 2002), they do not develop inside the animal. The larvae also do not develop inside humans or other animals.
Pupae: The pupal stage typically lasts
Medical Significance (Back to Top)
In the U.S., the biting midges are primarily a nuisance and the major medical issue associated with Culicoides is allergic reactions to the bites. However, like other blood feeding Diptera, Culicoides species are vectors of pathogens that can cause disease in humans and animals. In Central and South America, western and central Africa, and some Caribbean islands, biting midges are the vectors of filarial worms in the genus Mansonella. These parasites cause infection in humans that produces dermatitis and skin lesions because the adult worms are located in the skin.
Biting midges, primarily the species Culicoides sonorensis, are responsible for transmission of bluetongue virus to sheep and cattle in the U.S. Bluetongue is a serious disease of ruminants. Bluetongue viruses are found world-wide and are transmitted by different Culicoides species in different regions. Many countries that are bluetongue free prohibit the movement of livestock from bluetongue endemic regions. The annual economic damage in lost trade is in the millions of dollars.
Other animal disease causing pathogens transmitted by the bite of infected biting midges include African Horsesickness virus in equines that is confined primarily to Africa and Epizootic Hemorrhagic Disease virus in ruminants found in North America and principally having lethal effects on deer. Some equines experience allergic reactions to the bites, resulting in equine allergic dermatitis, affecting the withers, mane, tail and ears of the animal.
Management and Prevention (Back to Top)
Historically, management methods included diking and drainage of marshlands to reduce the habitats used by the immature stages. The insecticide DDT was used to target the adult stage. Currently, larval habitats are not targeted in control efforts because of the extensive amount of area that the habitats may cover, some negative environmental impacts resulting from changing water flow patterns of large areas, and the spotty spatial distribution of larvae within a given habitat.
Applications of insecticides targeting the adult stage are not efficient. While this type of application may kill biting midges active on a given night, they are continually dispersing from the larval habitat and entering areas of human activity. It would require insecticide applications on a daily basis in some areas, and this is not efficient or environmentally sound. Many government agencies that provide mosquito control services receive complaint calls about biting midges. However, most of the programs are not mandated or allowed to respond by providing control measures.
On a large scale, removal trapping is conducted using CO2 as an attractant to lure the biting midges to an insecticide-treated target where they are killed. Research from the the University of Florida, Institute of Food and Agricultural Sciences Florida Medical Entomology Laboratory showed that biting midge populations were reduced in test areas of Vero Beach and Boynton Beach, FL, and Castaway Cay, Bahamas. This method of control is more appropriate for islands and specific inland areas where pest control personnel can make a long term commitment to this technique.
Homeowners can install proper screening for windows and patios to prevent no-see-ums from entering residences and outdoor areas used for leisure and entertaining. Most biting midges can pass through 16-mesh insect wire screen and netting, so a smaller mesh size is required. The small mesh size does limit air flow through the screens. Additionally, because no-see-ums are so small and are weak fliers, ceiling and window fans can be used at high speeds to keep no-see-ums out of small areas.
Repellents containing DEET (N,N-diethyl-meta-toluamide) typically used as mosquito repellents are also labeled for use against no-see-ums and can be applied prior to exposure to the biting midges. It is important that the directions for application that are printed on the label are followed for any product used as a repellent.
Coastal areas provide primary habitat for biting midges. Tourists and potential home and land owners can consult local maps prior to visiting or purchasing property in coastal areas, to determine the proximity to biting midge producing areas. It is prudent to research the area of geographic interest prior to making decisions that can lead to an unpleasant vacation or unhappy homeowners. Knowing the habitats, and that large scale control operations are not feasible, one can be prepared with repellents or make decisions to build, or visit, elsewhere.
Selected References (Back to Top)
- Blanton FS, Wirth WW. 1979. The sand flies (Culicoides) of Florida (Diptera: Ceratopogonidae). Arthropods of Florida and Neighboring Land Areas Volume 10. Florida Department of Agriculture and Consumer Services. Gainesville, FL. 204 pp.
- Day, JF, Duxbury, CG, Glasscock, S and Paganessi, JE. 2001. Removal trapping for the control of coastal biting midge populations. Technical Bulletin of the Florida Mosquito Control Association. 4th Workshop on Salt Marsh Management and Research. Florida Mosquito Control Association, Ft. Myers, FL. 3: 15-16.
- Eldridge, BF and Edman, JD, Eds. 2000. Medical Entomology: A Textbook on Public Health and Veterinary Problems Caused by Arthropods. Kluwer Academic Publishers, Dordrecht, The Netherlands.
- Foote RH, Pratt HD. 1954. The Culicoides of the eastern United States (Diptera, Heleidae). Public Health Monograph No. 18. Publication No. 296. U. S. Department of Health, Education and Welfare, Public Health Service. 53 pp.
- Holbrook FR. 1996. Biting midges and the agents they transmit. In Beaty BJ, Marquardt WC (Eds), The Biology of Disease Vectors. University Press of Colorado, Niwot, CO. p. 110-116.
- Mullen G. Biting midges (Ceratopogonidae). In Mullen G, Durden L (Eds). 2002. Medical and Veterinary Entomology. Elsevier Science, San Diego, CA. p. 163-183.
- Rutledge CR, Day JF. 2002. Mosquito Repellents. EDIS. University of Florida/IFAS. (15 June 2016)
Author: C. Roxanne Connelly, University of Florida, Entomology and Nematology Department
Photographs: C. Roxanne Connelly, University of Florida, Entomology and Nematology Department Ed T. Schmidtmann, United States Department of Agriculture, Agricultural Research Services
Web Design: Don Wasik, Jane Medley
Publication Number: EENY-349
Publication Date: May 2005. Latest Revision: August 2013. Reviewed: June 2019.
Fleas often breed in large numbers where pets and other animals live. Pets infested with fleas bite and scratch themselves constantly. Their coats become roughened, and the skin can become infected. Symptoms of sensitized hosts are often mistaken for mange. Cat fleas and dog fleas may be intermediate hosts for the dog tapeworm.
Some people suffer more than others from flea bites. The bites can cause intense itching, often resulting in secondary infection. The usual flea bite has a small red spot where the flea has inserted its mouthparts. Around the spot there is a red halo with very little swelling. Many people do not react to flea bites at all, while others are sensitive and suffer severe allergic reactions. Fleas may also vector such human diseases as plague, typhus, and tularemia.
Crataerina pallida, a flightless species of hippoboscid fly that parasitizes common swifts, “gives birth” to a prepuparium. Photo by Jurgens Pasi.
By Meredith Swett Walker
The Hippoboscidae, commonly known as “louse flies” or “keds,” are a family of rather bizarre flies that are probably more familiar to ornithologists, sheep ranchers, and equestrians than they are to most entomologists. You are unlikely to see one unless you have a bird in your hand, or are grooming a horse, because hippoboscid flies are obligate parasites. They feed on blood and only blood, and they stay very close to their lunch.
Hippoboscid flies are fairly particular about their hosts. Sheep keds are not found on birds or vice versa. There are more than 200 species of Hippoboscidae, and 75 percent of these parasitize birds of various types ranging from tiny swifts to huge albatrosses. Some louse-flies even exhibit distinct preferences for a particular species of bird. One species of hippoboscid is found exclusively on frigate birds and another species parasitizes only boobies. This specificity is seen even when the two seabirds nest in densely-packed, mixed colonies where it would be easy for a hippoboscid to fly from one bird to another.
Thankfully, hippoboscids do not parasitize humans. In 1931, G. Robert Coatney conducted an experiment to determine if pigeon louse flies, Pseudolynchia canariensis, would bite humans and survive on human blood. He must have been very persuasive because he convinced two friends to join him in playing host to the flies. The answer is yes — hippoboscids will bite humans when given no other choice of host, and their bites are definitely itchy. But the flies did not survive long or reproduce when fed only human blood. Granted, Coatney’s experiment was limited in sample size and scope, but hopefully no one feels the need to repeat it.
Pseudolynchia canariensis, a pigeon louse fly. Photo by Jessica Waite.
Hippoboscids are very mobile — most species can fly. But despite their mobility, they rarely spend any time off of their hosts. A fly dislodged from its host will quickly fly back to it or the next closest host. This ability to move easily from one host individual to another makes hippoboscids an effective vector for blood-borne pathogens like avian malaria. The only phase of their life cycle when they are not intimately associated with their host is during pupation. At this stage, hippoboscids are easy to find (just find the host), which makes them very useful for scientists studying disease ecology.
The most striking thing about a hippoboscid’s appearance is its shape. Their bodies are distinctly dorsoventrally flattened (from back to belly), as if someone had dropped a book on them. This flat body shape allows them to slide between the feathers and scuttle around in the fur of their hosts. Their shape and tough exoskeletons also make them hard to squish, both for their hosts and for the humans that study them. According to one scientist who has worked on hippoboscids, you can’t just smack them, you have to “roll them between your fingers” to kill them.
Two hippoboscid flies on a nesting Nazca booby in the Galapagos Islands. Photo by Iris Levin.
But the most bizarre aspect of hippoboscid biology is definitely their life cycle. Most flies lay eggs, which hatch into larvae or maggots. The larvae feed independently, developing through various stages called “instars” until forming puparia, undergoing complete metamorphosis, and emerging as adult flies. But along with tse-tse flies and bat flies, the hippoboscid flies belong to the group formerly known as the Pupipara or “pupa-bearers.” Rather than laying eggs, female “pupa-bearers” essentially lay a pupa or “prepuparium” — a late-stage larva enclosed in a shell that quickly hardens into a true pupa. Female hippoboscids produce one offspring at a time. A single egg hatches in the female’s uterus and the resulting larva develops ther,e feeding from “milk glands.” The larva does not leave its mother’s body until it is fully-grown and ready to pupate. In hippoboscid species that parasitize birds, females usually deposit the pupae in their host bird’s nest or roosting site, where the newly-emerged adult fly will easily find a new host.
Dr. Jessica Waite picks through pigeon droppings to collect dead pigeon louse flies and their pupae. Photo courtesy of Jessica Waite.
This unusual process entails an enormous energetic investment by the female fly. According to Dr. Jessica Waite, an infectious disease biologist at Penn State University who has worked extensively on pigeon louse flies, the pupa can weigh more than the mother herself since the shell encasing it is included in the weight.
Dr. Waite never really set out to study hippoboscid flies. She started out as a bird lover who was interested in avian disease ecology. Like most people who handle birds, she had no love for the louse flies. As Miriam Rothschild and Theresa Clay wrote in Fleas, Flukes and Cuckoos: A Study of Bird Parasites, “For reasons which defy analysis, louse-flies are particularly repellent insects, and most people experience a shudder of disgust at the sight of them, and are filled with a quite unreasonable feeling of horror if they happen to dart up their sleeves or into their hair while handling the host.”
For her dissertation research, Waite studied the relationships between parasites, hosts, and vectors, and she was specifically interested in the costs to each of the organisms in this cycle. The pigeon (host), avian malaria (parasite) and pigeon louse fly (vector) system turned out to be ideal for measuring costs for the vector because of the quirks of the fly’s biology. Because both sexes of pigeon louse fly feed on blood and can become infected with the malaria parasite, Waite could make comparisons that aren’t possible in other arthropod vectors, such as mosquitoes. She could measure the reductions in survival associated with malaria infection in female louse flies, which invest heavily in reproduction compared to the males, which invest relatively little.
Working on this malaria system required keeping all three organisms in the lab, and Waite performed elegant experiments that elucidated the costs inflicted by malaria infection on the louse flies and the costs inflicted by louse flies on the pigeons. The daily grind of this work involved long hours of picking through pigeon droppings, counting dead flies, and collecting pupae, and after a while she became something of an accidental expert on hippoboscid biology.
“They are really creepy, especially in the way they move, but definitely fascinating,” Waite said.
Like Waite, Dr. Iris Levin never thought she’d end up working with hippoboscid flies.
“If you had told me at the start of my dissertation work that I’d end up working on these flies, I would have laughed hysterically,” she said.
Levin, an integrative biologist now at the University of Colorado, Boulder, was studying great frigate birds and a different species of avian malaria parasite, Haemoproteus iwa, in the Galapagos Islands. She was pretty sure that the hippoboscid fly, Olfersia spinifera, was transmitting the malaria, but to confirm the fly as the vector and understand the patterns of malaria transmission, she had to prove that individual flies bit more than one bird, and then estimate how frequently this host switching occurred. Instead of marking individual flies and trying to recapture them on different birds as had been done with insect vectors in the past, Levin found a clever work-around. She dissected the flies and extracted the bird blood from the last meals in their guts. She then compared DNA in that bird blood to DNA from the bird on which the fly was captured. If they didn’t match, it proved that the fly had fed on one frigate bird and then moved on to another.
Levin was also interested in the population genetics of the flies and their hosts. She compared DNA samples from populations of frigate birds and flies breeding on different islands. Levin found that different populations of flies were more closely related to each other than were the populations of frigate birds on which they were found. This meant that despite always staying close to their hosts, the flies were more frequently breeding with flies from other islands than the birds were. Because flies are very unlikely to fly from island to island on their own, this meant that the frigate birds were congregating and swapping flies at places and times when they weren’t breeding. So looking at the population genetics of the flies told Levin something about the birds’ movements that she wouldn’t have learned by studying the birds alone.
“In a way, hippoboscid flies are the world’s cheapest geolocator,” said Levin half seriously.
There is still much to be learned about the fascinating, yet slightly disgusting, hippobscid flies. Unlike most other biting diptera, both males and females feed on blood. What can that tell us about the evolution of blood feeding in flies? Did their unusual “pupa-bearing” reproductive strategy evolve in conjunction with their parasitic habits? Entomologists interested in studying this fly may have to learn how to catch birds or handle livestock. Or better yet, collaborate with a veterinarian or an ornithologist — you may end up converting them into accidental entomologists.
What makes an ideal insect repellent?
The ideal insect repellent should aim to have the following properties:
- Active against a wide variety of biting insects
- Prolonged activity (remain effective for at least 8 hours between applications)
- Non-irritating to the skin and mucous membranes
- Cosmetically appealing (odourless or have a pleasant odour and greaseless)
- No systemic toxicity
- Resistant to abrasion, washing and sweating
- Chemically stable and doesn’t react with commonly used plastics
- Economically viable for widespread use.
Currently there are no insect repellents that meet all the criteria listed above. It is extremely difficult to find a single active chemical that is effective against the many different species of disease-carrying insects. DEET is the most broad-spectrum and most effective insect repellent that has been developed to date. However it has recently been discovered that the chief malaria-carrying mosquito, Anopheles albimanus, in the United States is becoming resistant to DEET.
Parasites of the Giant Panda: A Risk Factor in the Conservation of a Species
Tao Wang , . Robin B. Gasser , in Advances in Parasitology , 2018
2.1.3 Ixodidae (Hard Ticks)
Other ectoparasites that can affect the health of the giant panda are (blood-feeding) hard ticks. Since the first description of tick infection by Haemaphysalis warburtoni ( Wu and Hu, 1985a ), an increasing number of hard tick species have been identified on giant pandas in the last two decades ( Lai et al., 1990 Ma, 1987 Qiu and Zhu, 1987 Yu et al., 1998 ). To date, 13 species representing 3 genera of hard ticks have been proposed as recorded (from rescued, sheltered or dead, wild giant pandas). These ticks include members of the genera Haemaphysalis (9 species), Ixodes (3 species) and Dermacentor (1 species) ( Table 2 ). Of these ticks, Haemaphysalis flava has been most commonly reported in giant panda populations ( Cheng et al., 2013 Ma, 1987 Qiu and Zhu, 1987 ). Recently, molecular tools have also been employed for the genetic characterisation of ticks from the giant panda. Using mitochondrial and ribosomal DNA markers as well as key morphological characters, H. flava was identified to predominate on giant pandas in the Qinling mountain range ( Cheng et al., 2013 ). Although there is no report on mortality caused by such hard ticks, morbidity involving dermatitis and/or weight loss has been recorded in infested giant pandas (CCRCGP, unpublished records). Similar to the treatment of C. panda, ivermectin and selamectin are the compounds most commonly used against ticks in breeding centres and zoos (CCRCGP, unpublished clinical records). To date, there are no reports of any associated tick-borne diseases in the giant panda.
Project funding under the Divisional Research and Development Program (DRDP) to the Institute of Bioinformatics and Biotechnology, Savitribai Phule Pune University, Pune, India, is greatly acknowledged. The authors acknowledge Dr Purushottam Lomate, Dr Sneha Bansode, Nidhi Saikhedkar and Dr Shadab Ahmed for their critical comments. The authors also acknowledge Dr Tuli Dey for editorial assistance in revising the manuscript. The authors declare no conflict of interest.