The research in our lab is currently funded by the Australian Research Council (ARC, DP170100008 and DP180100144), the US Air Force Office of Scientific Research (AFOSR, FA9550-15-1-0188), and Stiftelsen Olle Engkvist Byggm&#228;stare (2016/348). We thank past members of the lab that have contributed to developing hoverfly stocks, Cederholms Lantbruk and C M &amp; T L Green &amp; Son for access to cow dung and hoverflies on their farms, and the Adelaide and Uppsala Botanic Gardens for collection permits and ongoing support.

1. Establish Captive E. tenax Colony
<ol>
	<li>Establish colony via the collection of mature larvae (Step 1.2) or alternatively via the collection of free-flying hoverflies (Step 1.3).</li>
	<li>Collection of Mature Larvae
	<ol>
		<li>Collect second and third instar larvae from manure pits at cattle farms. 
		NOTE: Mature larvae are easiest to find during the beginning of their migratory phase, as they are actively seeking a dark dry environment to pupate. This tends to be near the borders of the manure pits where damp manure is close to drier areas containing large amounts of straw. We collected under permission from a cattle farm near Uppsala, Sweden.</li>
		<li>Rear mature larvae in cow manure as outlined in Step 3.2.</li>
	</ol>
	</li>
	<li>Collection of Wild Hoverflies
	<ol>
		<li>Collect wild hoverflies by netting in the field, usually from botanic gardens and parks where there is an abundance of flowering plants. 
		NOTE: We collected under permission from several locations including any of the three Adelaide Botanic Gardens, a dairy farm in Myponga, South Australia, and at various botanic gardens and parks throughout Uppsala, Sweden.</li>
		<li>House field collected hoverflies as outlined in Step 2.</li>
	</ol>
	</li>
</ol>
2. Housing and Long-Term Maintenance of Hoverflies
<ol>
	<li>House hoverflies in 30 cm x 45 cm plastic bags for groups of 20 or less, or in an insect rearing cage (25 cm x 25 cm x 25 cm) for larger groups.</li>
	<li>Provide food and water ad libitum, in the form of 10-20 grains of bee pollen and 2-3 mL honey placed on top of several moist cotton balls. 
	NOTE: For housing in plastic bags, it is important that the cotton balls are moist but not overly saturated, as any accumulation of water inside the bag can be detrimental to survival rates. Conversely, for housing in insect rearing cages, the mesh sides allow for significant evaporation to occur. Cotton balls should therefore be placed in a shallow container and be fully saturated.
	<ol>
		<li>Leave hoverflies to feed for 6 hours at room temperature.</li>
		<li>Place hoverflies with food and water in their housing, in the fridge at 8-10 &#176;C and keep in complete darkness. 
		NOTE: Storing hoverflies at 8-10 &#176;C in darkness causes the hoverflies to enter a state of hibernation, with a reduction in activity and metabolic rate.</li>
	</ol>
	</li>
	<li>Every 3-4 days remove hoverflies from the fridge, thus breaking the artificial hibernation and allowing both feeding and grooming to occur.
	<ol>
		<li>Transfer hoverflies to a new plastic bag or clean insect cage with fresh food and water. This transfer can either be done manually, for small numbers of flies, or by utilizing phototaxis, for larger numbers.</li>
		<li>To utilize phototaxis, join a clean insect cage to the old one, ensure there is an opening for the hoverflies to move freely between the two without escaping. Cover the old insect cage with an opaque fabric. The hoverflies will move towards the light and into the clean insect cage.</li>
	</ol>
	</li>
	<li>Allow hoverflies to feed and groom at room temperature for 6 h.</li>
	<li>Return hoverflies, with the food and water in their housing, into the fridge at 8-10 &#176;C in complete darkness. This recommences the artificial hibernation of the hoverflies.</li>
	<li>Continue the cyclical breaking of hibernation, every 3-4 days, to ensure the health and longevity of hoverflies for the duration of their captivity.</li>
</ol>
3. Laboratory Rearing of E. tenax
<ol>
	<li>Rear mature larvae collected from cattle farms (Step 3.2) or alternatively rear eggs laid by gravid wild caught females (Step 3.3).</li>
	<li>Laboratory Rearing of Mature Larvae from Cattle Farms
	<ol>
		<li>Place larvae collected in cow manure from cattle farms in 30 L buckets.</li>
		<li>Position the bucket, containing mature larvae, within a larger box or bag (minimum volume of 50-60 L) and place 20-30 L wood shavings up to the height of the rim of the bucket. 
		NOTE: This allows 3rd instar larvae to crawl into the wood shavings and pupate.</li>
		<li>Hang a double layered mosquito net from the ceiling allowing it to drape over the boxes and/or bags, thus ensuring that neither larvae nor any emerging hoverflies can escape. 
		NOTE: As a precaution, double-sided sticky tape can be used to surround the setup, as any escaping larvae will get stuck and pupate on this tape. If they do, remove the pupae before eclosion.</li>
		<li>House both larvae and pupae at room temperature (21.5 &#177; 2.5 &#176;C), and expose to either indirect sunlight as well as room lights during office hours or keep at a light:dark cycle of 12 h light:12 h dark. 
		NOTE: Exposure to 24 h light can be detrimental to survival rates. For larvae collected from cattle farms pupation time will vary from 1- 20 days after collection depending on their maturity at the time of collection.</li>
		<li>Provide food and water within the hanging mosquito net enclosure (as prepared in Step 2.2) prior to the expected eclosion date and replace every 2-3 days. Eclosion will occur 7-10 days after pupation.</li>
		<li>Allow emerging hoverflies to feed for 6 hours at room temperature before placing them in housing as outlined in Step 2.</li>
	</ol>
	</li>
	<li>Laboratory Rearing of Eggs laid by Gravid Wild Caught Females
	<ol>
		<li>Check hoverfly housing for eggs, both before changing housing (Step 2), and prior to returning them to the fridge. Oviposition by wild-caught gravid females occurs at both 8-10 &#176;C and at room temperature.
		<ol>
			<li>Place eggs in a 100 mm x 20 mm Petri dish containing 70 mL of tap water and keep at room temperature until hatching occurs, usually 2-3 days after oviposition.</li>
		</ol>
		</li>
		<li>Place hatched larvae in a 2.3 L bucket containing 1 L of fresh rabbit feces and 1 L of tap water.
		<ol>
			<li>Check slurry every 2-3 days and add additional tap water as required, to ensure that the slurry does not dry out before the 3rd instar larvae emerge.</li>
		</ol>
		</li>
		<li>Position the bucket, containing the larvae in the rabbit feces slurry, within a larger box (minimum volume 30 L) containing 20 L of wood shavings. Ensure the wood shavings are up to the height of the rim of the bucket. 
		NOTE: This allows 3rd instar larvae to crawl into the wood shavings and pupate.
		<ol>
			<li>Place a double layered mosquito net over the box to ensure that neither larvae nor any emerging hoverflies can escape.</li>
		</ol>
		</li>
		<li>House both larvae and pupae at room temperature (21.5 &#177; 2.5 &#176;C), and expose to either indirect sunlight as well as room lights during office hours or keep at a light:dark cycle of 12 h light:12 h dark. 
		NOTE: Exposure to 24 h light can be detrimental to survival rates.</li>
		<li>Expect pupation to occur after 15-20 days. Collect pupae and place in an insect cage, allowing hoverflies to eclose there.</li>
		<li>Provide food and water (as prepared in Step 2.2) prior to the expected eclosion date - eclosion will occur 6-10 days after pupation - and replace every 2-3 days.</li>
		<li>Allow emerging hoverflies to feed for 6 hours at room temperature before placing them in housing, as outlined in Step 2. 
		NOTE: Both the pupation of larvae and the eclosion of pupae can be delayed by storage in darkness at 8-10 &#176;C. For this purpose, store larvae in the rabbit feces slurry and pupae in wood shavings.</li>
	</ol>
	</li>
</ol>

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Genetic Variation in the dronefly Eristalis tenax.</a> Heredity. 42 (2), 223-236 (1979).</li><li>Ireland, S., Turner, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+effects+of+larval+crowding+and+food+type+on+the+size+and+development+of+the+blowfly,+Calliphora+vomitoria.">The effects of larval crowding and food type on the size and development of the blowfly, Calliphora vomitoria.</a> Forensic Science International. 159, 175-181 (2006).</li><li>Dolley, W. L., Golden, L. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+effect+of+sex+and+age+on+the+temperature+at+which+reversal+in+reaction+to+light+in+Eristalis+tenax+occurs.">The effect of sex and age on the temperature at which reversal in reaction to light in Eristalis tenax occurs.</a> Biology Bulletin. 92 (3), 178-186 (1947).</li></ol>

The authors have nothing to disclose.

Using our techniques (Figure 1) hoverflies have been maintained in the laboratory for a period of over 1 year and successfully used in behavioral experiments after 7 months in captivity39. Indeed, even if it seems counterintuitive, keeping the hoverflies in a more natural environment, under 12 h light:12 h dark conditions, at room temperature, substantially decreases their life expectancy to 2-3 months. Maintaining E. tenax in our artificial hibernation cycle for over a year is significantly longer than previous attempts using different protocols (77 days33, 4 months9, 18 weeks30). The main factor influencing this increased longevity is likely the use of artificial hibernation at 8-10 &#176;C. By cyclically breaking the hibernation, every 3-4 days (Step 2, Figure 1), we allow hoverflies to both feed and self-groom, thus maintaining the nutritional state and wellbeing of the hoverflies, as evidenced by an observed increase in weight (Figure 4) and no change in locomotor activity even after long periods in captivity (Figure 3, and see39). Indeed, reports in the literature of unsuccessful attempts at artificial hibernation do not break the hibernation cycle, thus leading to increased mortality and the presence of mold9.
Additionally, there is some controversy throughout the literature as to the provision of pollen as a food source. Several papers state that bee pollen is not sufficient, specifically for oviposition, and only the provision of dry or indeed fresh pollen is suitable9,29. Our findings indicate that by complementing bee pollen with honey and water, we see both longevity and oviposition, even after long periods of captivity, with an increase in weight seen in both sexes (Figure 4) and oviposition still occurring in females after more than 5 months in captivity39. This increased longevity allows us to study the behaviors of hoverflies at all life stages.
In the field, female hoverflies are fertilized prior to seasonal hibernation and remain in reproductive diapause, where sperm is stored and oocytes remain underdeveloped, until spring28. Given that a typical female is capable of laying 3000 eggs in 60 days29, rearing of these eggs is therefore a quick and efficient way to increase our captive population. However, our current understanding of the factors that lead to oocyte development after a period of hibernation are limited. Temperature, humidity, light intensity and nutritional state have all been suggested as playing a role in controlling reproductive diapause28,40. Experimental manipulation of such factors may lead to a greater governance of oviposition timing and rates.
Similarly, we have successfully delayed the development of larvae, as well as the eclosion of pupae, by storage in darkness at 8-10 &#176;C for 2 weeks, although viability may be much longer. Indeed, Heal30&#160;reported a pupal duration increase of up to 37 days when the pupal temperature was dropped from 25 &#176;C to 10 &#176;C. Employing these strategies and delaying the production of eggs and/or the development of pupae would allow for a greater manipulation of the demographics of the captive population.
While temporal consistency of supply is of much greater importance for our requirements than large yield, this may be more important for other uses, such as pollination in greenhouses. We found that when using our technique with rabbit feces, we got 163 &#177; 34 eclosed hoverflies from each clutch of eggs (N=3). Given that a typical female lays up to 200 eggs40, we might be able to increase this yield by either decreasing overcrowding and food competition, or by adjusting the temperature, as these have been implicated as significantly affecting larval growth9,31,40,41. However, there is no indication that the basis of the media greatly influences yield32. In addition, as opposed to feces from other vertebrates29,30,31,42, rabbit feces is relatively odor free, allowing the colony to be kept under normal laboratory conditions without the need for additional ventilation. Decreasing the density of larvae in the media, or adding nutritional supplements such as yeast, as well as keeping a constant temperature between 20-25 &#176;C, is probably sufficient to fully optimise yield31,32,40.
The practicalities of collecting sufficient numbers of freely flying hoverflies, or sustaining a genetically diverse captive population, are both impractical and time restrictive for small scale research projects. Therefore, rearing the offspring of wild mated females, and supplementing supplies by harvesting mature larvae7, allow the most practical options for year-round use of E. tenax in a research setting. As these methods are limited by the seasons in which collection can occur, there is a need for both ensuring the longevity of adult hoverflies and rearing any eggs laid by captured gravid females.

Hoverflies are emerging as useful models for investigating a range of scientific questions, including flight behavior1, neural mechanisms underlying motion vision2, pollination efficiency3,4,5,6&#160;and microbial immunity7. However, as opposed to some other dipteran models, such as Drosophila8, there are no standardized protocols for lab rearing of hoverflies for use in scientific research. Indeed, even if the current literature describes methods for breeding the hoverfly Eristalis tenax, many of these are developed for the mass cultivation of hoverflies for crop pollination, bio-degradation of organic waste, or anatomical studies9,10,11. Thus, they do not address the need for an easy protocol that provides a consistent supply of healthy robust hoverflies, whilst maintaining the genetic fitness of the population.
Following bees and bumblebees, hoverflies are one of the most important wild, generalist pollinator groups12,13. There are about 6000 hoverfly species worldwide14,15, with more than 300 species in 75 genera in Sweden16&#160;and more than 300 species in 69 genera in India17,18,19. For example, the agriculturally important marmalade hoverfly Episyrphus balteatus and the drone fly, Eristalis tenax, which we focus on here, are found across Europe, America and Asia6,16,17,18,19,20,21,22,23,24,25. Hoverflies are not equally active throughout the year, nor throughout the day. Indeed, not only the season, and the time of day, but also fluctuations in light intensity, temperature, humidity and wind velocity, affect the activity patterns of hoverflies26,27. In the field, Eristalis can be found at any time of the year in Mediterranean climates11, but the numbers of active hoverflies are much lower in winter. Conversely in cold temperate climates, Eristalis hibernate over winter and are not found actively behaving in the field from around October through to March28.
Freely flying hoverflies can be collected by netting in the field. Indeed, in temperate climates they are found in the greatest abundance in the mid to late morning, on calm sunny days, at the end of summer and throughout autumn26,27. Alternatively, mature E. tenax larvae, second or third instar, can be identified and harvested from decaying organic matter, such as manure heaps or organically polluted streams10,11. Indeed, published techniques for lab rearing of E. tenax are all based on raising larvae in organically polluted water, either via some form of vegetative or fecal matter9,10,29,30,31,32,33. However, larvae collection is limited by season, and is only a viable collection tool from late spring to the start of autumn11. Furthermore, the abundance of larvae is affected by local weather patterns, as changes in ambient temperature can affect both the occurrence of oviposition and larval development rates9,28.
Therefore, strategies to maintain healthy stocks of hoverflies by rearing larvae and eggs within the laboratory are needed to ensure that experiments can be conducted year-round, irrespective of the season or local weather events. Importantly, the technique described here breeds the hoverflies from only wild-mated females.This is important as a study by Francuski, et al.10&#160;found that the genetic diversity of a laboratory bred population of hoverflies, originally established from 120 mature larvae, was rapidly lost. They therefore suggested that to maintain genetic diversity in colonies to be used for commercial crop pollination purposes, these need to be replenished, or even completely re-established, with field collected individuals each spring10.
When working on vision, or other senses used in courtship and mating, we thus recommend maintaining genetic diversity, either by re-establishing the colony or by replenishing the colony with field collected individuals, regularly. This is important as sexual selection affects the genetic drift of the population. Indeed, in the wild, male hoverflies need to identify and intercept suitable mates, as well as compete with other males for mating rights by defending their territories34. This process ensures that males with the best vision and spatial attention are likely to be the most successful in mating, and hence these traits are passed on to the next generation. The resultant effect of these ongoing processes is, in part, demonstrated by the presence of sexual dimorphisms in the visual pathway of hoverflies35,36. In captivity males do not have the same obstacles to successful mating as in the field: firstly, females are readily available, and secondly the small, confined enclosure negates the effect of territorial behaviors, which act to deter the mating access of other competitive males. The experimental removal of sexual selection in Drosophila melanogaster, has been shown to have a significant effect on captive populations with a decrease in overall body size, testes size and sperm production37, and reduced rates of male courtship behaviors38. Thus, a captive breeding program, without any consideration of sexual selection, may have a profound effect on both the visual and behavioral studies subsequently conducted.
We here describe a simple and cost-effective solution that provides a consistent supply of healthy hoverflies. The protocol is flexible and easy to re-start and/or upscale, depending on the research demands.

<table><tbody><tr><td>Bee Pollen</td><td>Forest Super Foods</td><td></td><td>any brand of bee pollen is suitable</td></tr><tr><td>Honey</td><td>Bramwells</td><td></td><td>any brand of liquid honey is suitable</td></tr><tr><td>Rabbit Faeces</td><td></td><td></td><td>can be substituted with cow or pig manure made into a slurry</td></tr><tr><td>BugDome</td><td>Australia Entomological Supplies</td><td>EM42222</td><td></td></tr><tr><td>Plastic Bags</td><td>Woolworths Homebrand</td><td></td><td></td></tr><tr><td>Mosquito netting</td><td>Clas Ohlson</td><td>34-1113</td><td></td></tr><tr><td>Cotton Balls</td><td>Woolworths Select</td><td></td><td></td></tr><tr><td>Fridge</td><td>Hisense</td><td></td><td>fridge needs to maintain a stable 8-10&amp;deg;C&amp;nbsp;</td></tr><tr><td>Buckets (2-3L)</td><td></td><td></td><td></td></tr><tr><td>Large plastic tubs (30L)</td><td></td><td></td><td></td></tr><tr><td>Wood shavings</td><td>Pollards Sawdust Supplies</td><td>MaxiFlake (75)&amp;nbsp;</td><td></td></tr><tr><td>Bag clips</td><td>IKEA</td><td>Bevara 303.391.70</td><td></td></tr><tr><td>Petri Dish (100mm x 20mm)</td><td>Corning</td><td>430167</td><td></td></tr></tbody></table>

rearing and long-term maintenance of eristalis tenax hoverflies for research studies

With an estimated 6000 species worldwide, hoverflies are ecologically important as alternative pollinators to domesticated honeybees. However, they are also a useful scientific model to study motion vision and flight dynamics in a controlled laboratory setting. As the larvae develop in organically polluted water, they are useful models for investigating investment in microbial immunity. While large scale commercial breeding for agriculture already occurs, there are no standardized protocols for maintaining captive populations for scientific studies. This is important as commercial captive breeding programs focusing on mass output during peak pollination periods may fail to provide a population that is consistent, stable and robust throughout the year, as is often needed for other research purposes. Therefore, a method to establish, maintain and refresh a captive research population is required. Here, we describe the utilization of an artificial hibernation cycle, in addition to the nutritional and housing requirements, for long term maintenance of Eristalis tenax. Using these methods, we have significantly increased the health and longevity of captive populations of E. tenax compared to previous reports. We furthermore discuss small scale rearing methods and options for optimizing yields and manipulating population demographics.

We have developed a three-way strategy that maintains a healthy population for both visual and behavioral studies (summarized in Figure 1). Our method starts with collection of hoverflies from the wild (Step 1, Figure 1). In our lab, hoverflies are housed in insect cages or plastic bags, under an artificial hibernation cycle (Step 2, Figure 1), substantially prolonging their life span. For increased numbers, offspring can be reared from wild-mated females (Step 3, Figure 1).
We have found that catching vast numbers of wild hoverflies is a time intensive endeavor, even when environmental conditions are favorable. In contrast, the successful rearing of mature larvae harvested from the manure pits of cattle farms is a far more efficient way to source large numbers of wild hoverflies (Step 1, Figure 1), with us collecting up to 700 larvae in 0.03 m3&#160;of manure. Additionally, our techniques to rear eggs laid by captured gravid females have proved to be successful (Step 3, Figure 1). Females captured in a Mediterranean climate (Adelaide) during the autumn and winter months laid several batches of eggs, with 24 clusters observed from 19 females in a period of 20 weeks. Of these egg batches, 10 were placed into water, all of which were fertile and resulted in the hatching of larvae. 3 groups of larvae were then taken beyond this point, and placed in the rabbit feces slurry, resulting in 163 &#177; 34 (mean &#177; SD, N=3) emerged hoverflies, with no observed gender bias (Figure 2).
The health of these lab reared hoverflies was determined by a comparison of the weight and locomotor activity of female hoverflies compared to field captured individuals. General locomotor activity was assessed using a Locomotor Activity Monitoring system (LAMS), as described previously39. No significant differences in weight (Figure 3A) or activity (Figure 3B) were observed between lab reared and wild caught hoverflies after 4 months in captivity under our artificial hibernation cycle. When E. tenax were maintained in the laboratory without the use of an artificial hibernation cycle we saw a significant decrease in longevity, with a lifespan of 2.5 - 3 months (73 &#177; 7 days for 5 females and 79 &#177; 4 days for 11 males). When the hoverflies were kept in artificial hibernation they could live in excess of 12 months.
Additionally, the effect of long term maintenance, using our described methods, was further assessed by a comparison of weights over time for both sexes of lab reared hoverflies. We observed a significant increase in weight over a period of 4 months for both sexes, with females consistently weighing more than their male counterparts (p &#60; 0.0001, two-way ANOVA, N=12, Figure 4).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57711/57711fig1.jpg" /> 
Figure 1:&#160;Flow diagram outlining the methods for maintenance of healthy captive population of E. tenax. (1) The procedure described here starts with collection of either mature larvae from cow dung (Step 1.2) or freely flying hoverflies (Step 1.3). (2) The hoverflies are housed in insect cages or plastic bags, dependent on numbers. They are kept in an artificial hibernation cycle at 8-10 &#176;C, which is broken every 3-4 days. (3) Collected larvae are kept in their cow dung (Step 3.2). Eggs laid in the lab are placed in a rabbit feces slurry (Step 3.3). When reaching maturity, 3rd instar larvae crawl into the surrounding saw dust where they pupate. Eclosion occurs after 6-10 days, and the eclosed hoverflies are placed in the housing (Step 2). <a href="https://www.jove.com/files/ftp_upload/57711/57711fig1large.jpg">Please click here to download this file.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57711/57711fig2v3.jpg" /> 
Figure 2:&#160;Number and sex ratio of E. tenax successfully reared from individual egg batches. The data show the numbers of E. tenax that eclosed from pupae developed from 3 batches of eggs laid in our lab. The eggs were laid by wild-caught females. The data are color coded for the sex of the flies. There is no significant difference in the ratio.&#160;
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/57711/57711fig3v3.jpg" /> 
Figure 3:&#160;Assessment of population health in laboratory reared and field collected female hoverflies after long term captivity. (A) Weight comparison between lab reared and field captured female hoverflies after 4 months in captivity under artificial hibernation (N=12). (B) Activity levels of lab-reared and field captured female hoverflies after 4 months in captivity under artificial hibernation.&#160;The hoverflies&#39; locomotor activity was measured in a Locomotor Activity System by them breaking an infrared beam during movement. As previously, we averaged the activity across 6.7&#160;hours in the middle of the day, on the second full day in the Locomotor Activity Monitor System39&#160;(Nfield caught=9, Nlab reared=12). The central mark of each boxplot shows the median, the edges of the box the 25th to 75th percentiles of the data, and the whiskers extend from the minimum to the maximum of the data.
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/57711/57711fig4v3.jpg" /> 
Figure 4:&#160;Comparison of the effect of long term maintenance on the weight of lab reared hoverflies of either sex. The data show the hoverfly weight as a function of time kept in captivity under artificial hibernation. As all hoverflies were reared from eggs laid in our lab (N=12 at each data point) and t=0 equals time of pupal hatching, the time in captivity is the same as the age of the animals. The central mark of each boxplot shows the median, the edges of the box the 25th to 75thpercentiles of the data, and the whiskers extend from the minimum to the maximum of the data.&#160;

Watch this Scientific Journal Video about Rearing and Long-Term Maintenance of Eristalis tenax Hoverflies for Research Studies at JoVE.com

Rearing and Long-Term Maintenance of Eristalis tenax Hoverflies for Research Studies

The overall goal of these procedures is to establish, maintain and refresh a captive population of Eristalis tenax in a research setting.

Rearing and Long-Term Maintenance of Eristalis tenax Hoverflies for Research Studies

With an estimated 6000 species worldwide, hoverflies are ecologically important as alternative pollinators to domesticated honeybees. However, they are ...

rearing-long-term-maintenance-eristalis-tenax-hoverflies-for-research

Research

JoVE Journal

Environment

13.4K Views.  Flinders University. Het algemene doel van deze procedures moet opzetten, onderhouden en vernieuwen van een gevangenschap bevolking van Eristalis tenax in de instelling van een onderzoek.

Video: Rearing and Long-Term Maintenance of Eristalis tenax Hoverflies for Research Studies

Early Metamorphic Insertion Technology for Insect Flight Behavior Monitoring

Early Metamorphosis Insertion Technology (EMIT) is a novel methodology for integrating microfabricated neuromuscular recording and actuation platforms on insects during their metamorphic development. Here, the implants are fused within the structure and function of the neuromuscular system as a result of metamorphic tissue remaking. The implants emerge with the insect where the development of tissue around the electronics during pupal development results in a bioelectrically and biomechanically enhanced tissue interface. This relatively more reliable and stable interface would be beneficial for many researchers exploring the neural basis of the insect locomotion with alleviated traumatic effects caused during adult stage insertions. In this article, we implant our electrodes into the indirect flight muscles of Manduca sexta. Located in the dorsal-thorax, these main flight powering dorsoventral and dorsolongitudinal muscles actuate the wings and supply the mechanical power for up and down strokes. Relative contraction of these two muscle groups has been under investigation to explore how the yaw maneuver is neurophysiologically coordinated. To characterize the flight dynamics, insects are often tethered with wires and their flight is recorded with digital cameras. We also developed a novel way to tether Manduca sexta on a magnetically levitating frame where the insect is connected to a commercially available wireless neural amplifier. This set up can be used to limit the degree of freedom to yawing &#8220;only&#8221; while transmitting the related electromyography signals from dorsoventral and dorsolongitudinal muscle groups.

We present a novel surgical procedure to implant electrodes in Manduca sexta during its early metamorphic stages. This technique allows mechanically stable and electrically reliable coupling with the neuromuscular tissue to study flight neurophysiology dynamics. We also present a novel magnetic levitation platform for tethered studies of insect yaw.

Early Metamorphosis Insertion Technology (EMIT) is a novel methodology for integrating microfabricated neuromuscular recording and actuation platforms on ...

Behavior

Flying Insect Detection and Classification with Inexpensive Sensors

An inexpensive, noninvasive system that could accurately classify flying insects would have important implications for entomological research, and allow for the development of many useful applications in vector and pest control for both medical and agricultural entomology. Given this, the last sixty years have seen many research efforts devoted to this task. To date, however, none of this research has had a lasting impact. In this work, we show that pseudo-acoustic optical sensors can produce superior data; that additional features, both intrinsic and extrinsic to the insect&#8217;s flight behavior, can be exploited to improve insect classification; that a Bayesian classification approach allows to efficiently learn classification models that are very robust to over-fitting, and a general classification framework allows to easily incorporate arbitrary number of features. We demonstrate the findings with large-scale experiments that dwarf all previous works combined, as measured by the number of insects and the number of species considered.

We proposed a system that uses inexpensive, noninvasive pseudo-acoustic optical sensors to automatically and accurately detect, count, and classify flying insects based on their flying sound.

An inexpensive, noninvasive system that could accurately classify flying insects would have important implications for entomological research, and allow ...

Bioengineering

Mass-Rearing and Molecular Studies in Tortricidae Pest Insects

Tortricidae (Lepidoptera), commonly known as tortrix or leafroller moths, comprises many agricultural and forestry pests, which cause serious agricultural losses. To understand the biology of such pest moths, fundamental techniques have been in high demand. Here, methods for mass-rearing, observations, and molecular studies are developed using two tea tortrix, Homona magnanima and Adoxophyes honmai (Lepidoptera: Tortricidae). Insects were mass-reared with sliced artificial diet and maintained by inbreeding for over 100 generations by considering their biological characteristics. Insects have various sex dimorphisms; hence it is difficult to distinguish the sex during the developing stages, which have prevented subsequent assays. The present work highlighted that the sex of tortricids larvae could be determined by observing testes or lactic-acetic orcein staining to visualize the female-specific W chromosome. Moreover, using the sex determination methods, the present study enabled nucleic acid extractions from sex determined embryos and application toward high throughput sequencing. These tips are applicable for other pest insects and will facilitate further morphological and genetic studies.

The present protocol describes the rearing method of tortricid pest insects in the laboratories. The procedures to distinguish insects' sex and extract nucleic acids for high throughput sequencing are established using two tortricid pests.

Tortricidae (Lepidoptera), commonly known as tortrix or leafroller moths, comprises many agricultural and forestry pests, which cause serious agricultural ...

A Proboscis Extension Response Protocol for Investigating Behavioral Plasticity in Insects: Application to Basic, Biomedical, and Agricultural Research

Insects modify their responses to stimuli through experience of associating those stimuli with events important for survival (e.g., food, mates, threats). There are several behavioral mechanisms through which an insect learns salient associations and relates them to these events. It is important to understand this behavioral plasticity for programs aimed toward assisting insects that are beneficial for agriculture. This understanding can also be used for discovering solutions to biomedical and agricultural problems created by insects that act as disease vectors and pests. The Proboscis Extension Response (PER) conditioning protocol was developed for honey bees (Apis mellifera) over 50 years ago to study how they perceive and learn about floral odors, which signal the nectar and pollen resources a colony needs for survival. The PER procedure provides a robust and easy-to-employ framework for studying several different ecologically relevant mechanisms of behavioral plasticity. It is easily adaptable for use with several other insect species and other behavioral reflexes. These protocols can be readily employed in conjunction with various means for monitoring neural activity in the CNS via electrophysiology or bioimaging, or for manipulating targeted neuromodulatory pathways. It is a robust assay for rapidly detecting sub-lethal effects on behavior caused by environmental stressors, toxins or pesticides.
We show how the PER protocol is straightforward to implement using two procedures. One is suitable as a laboratory exercise for students or for quick assays of the effect of an experimental treatment. The other provides more thorough control of variables, which is important for studies of behavioral conditioning. We show how several measures for the behavioral response ranging from binary yes/no to more continuous variable like latency and duration of proboscis extension can be used to test hypotheses. And, we discuss some pitfalls that researchers commonly encounter when they use the procedure for the first time.

The Proboscis Extension Response (PER) conditioning protocol, developed for the honey bee (Apis mellifera), provides an ecologically-relevant and easily quantifiable means for studying several different mechanisms of learning in many insect species.

Insects modify their responses to stimuli through experience of associating those stimuli with events important for survival (e.g., food, mates, threats). ...

Neuroscience

Maintaining Aedes aegypti Mosquitoes Infected with Wolbachia

Aedes aegypti mosquitoes experimentally infected with Wolbachia are being utilized in programs to control the spread of arboviruses such as dengue, chikungunya and Zika. Wolbachia-infected mosquitoes can be released into the field to either reduce population sizes through incompatible matings or to transform populations with mosquitoes that are refractory to virus transmission. For these strategies to succeed, the mosquitoes released into the field from the laboratory must be competitive with native mosquitoes. However, maintaining mosquitoes in the laboratory can result in inbreeding, genetic drift and laboratory adaptation which can reduce their fitness in the field and may confound the results of experiments. To test the suitability of different Wolbachia infections for deployment in the field, it is necessary to maintain mosquitoes in a controlled laboratory environment across multiple generations. We describe a simple protocol for maintaining Ae. aegypti mosquitoes in the laboratory, which is suitable for both Wolbachia-infected and wild-type mosquitoes. The methods minimize laboratory adaptation and implement outcrossing to increase the relevance of experiments to field mosquitoes. Additionally, colonies are maintained under optimal conditions to maximize their fitness for open field releases.

Maintaining Aedes aegypti Mosquitoes Infected with Wolbachia

Aedes aegypti mosquitoes infected with Wolbachia are being released into natural populations to suppress the transmission of arboviruses. We describe methods to rear Ae. aegypti with Wolbachia infections in the laboratory for experiments and field release, taking precautions to minimize laboratory adaptation and selection.

Aedes aegypti mosquitoes experimentally infected with Wolbachia are being utilized in programs to control the spread of arboviruses such as dengue, ...

Biology

Laboratory Rearing of Stable Flies and Other Muscoid Diptera

Stable flies, Stomoxys calcitrans, are serious pests of livestock, humans, companion animals and wildlife worldwide. During the last 20+ years, changes in agronomic practices resulted in serious outbreaks of stable flies in several countries. These outbreaks disrupted livestock production and human recreation resulting in public demands for increasing research and management efforts for this pest. A simple and inexpensive procedure for rearing stable flies for laboratory studies is presented. The procedure uses locally available diet components, equipment and supplies. The procedure can be adapted for rearing other muscoid flies including face fly (Musca autumnalis), horn fly (Haematobia irritans), and house fly (Musca domestica). The procedure produces stable fly puparia averaging 12.5 mg and ~35% egg to adult survival. Approximately 3000 flies are produced in each pan.

A procedure for rearing stable flies (Stomoxys calcitrans) is presented. The procedure uses locally available materials for diet components, equipment and supplies.

Stable flies, Stomoxys calcitrans, are serious pests of livestock, humans, companion animals and wildlife worldwide. During the last 20+ years, changes in ...

At-Risk Butterfly Captive Propagation Programs to Enhance Life History Knowledge and Effective Ex Situ Conservation Techniques

Improving knowledge of ex situ best practices for at-risk butterflies is important for generating successful conservation and recovery program outcomes. Research on such captive populations can also yield valuable data to address key information gaps about the behavior, life history, and ecology of the target taxa. We describe a protocol for captive propagation of the federally endangered Cyclargus thomasi bethunebakeri that can be used as a model for other at-risk butterfly ex situ programs, especially those in the family Lycaenidae. We further provide a simple and straightforward protocol for recording various life history metrics that can be useful for informing ex situ methodologies as well as adapted for laboratory studies of other lepidoptera.

Here, we present protocols for 1) the laboratory captive propagation of the federally endangered Miami blue butterfly (Cyclargus thomasi bethunebakeri), and 2) assessing basic life history information such as immature development time and number of larval stadia. Both methods can be adapted for use with other ex situ conservation programs.

Improving knowledge of ex situ best practices for at-risk butterflies is important for generating successful conservation and recovery program outcomes. ...

Maintaining Laboratory Cultures of Gryllus bimaculatus, a Versatile Orthopteran Model for Insect Agriculture and Invertebrate Physiology

Gryllus bimaculatus (De Geer)&#160;is a large-bodied cricket distributed throughout Africa and Southern Eurasia where it is often wild-harvested as human food. Outside its native range, culturing G. bimaculatus is feasible due to its dietary plasticity, rapid reproductive cycle, lack of diapause requirement, tolerance for high-density rearing, and robustness against pathogens. Thus, G. bimaculatus can be&#160;a versatile model for studies of insect physiology, behavior, embryology, or genetics.
Cultural parameters, such as stocking density, within-cage refugia, photoperiod, temperature, relative humidity, and diet, all impact cricket growth, behavior, and gene expression and should be standardized. In the burgeoning literature on farming insects for human consumption, these crickets are frequently employed to evaluate candidate feed admixtures derived from crop residues, food-processing byproducts, and other low-cost waste streams.
To support ongoing experiments evaluating G. bimaculatus growth performance and nutritional quality in response to variable feed substrates, a comprehensive set of standard protocols for breeding, upkeep, handling, measurement, and euthanasia in the laboratory was developed and is presented here. An industry-standard cricket feed has proven nutritionally adequate and functionally appropriate for the long-term maintenance of cricket breeding stocks, as well as for use as an experimental control feed. Rearing these crickets at a density of 0.005 crickets/cm3 in screen-topped 29.3 L polyethylene cages at an average temperature of 27 &#176;C on a 12 light (L)/12 dark (D) photoperiod, with moistened coconut coir serving both as hydration source and oviposition medium has successfully sustained healthy crickets over a 2-year span. Following these methods, crickets in a controlled experiment yielded an average mass of 0.724 g 0.190 g at harvest, with 89% survivorship and 68.2% sexual maturation between stocking (22 days) and harvest (65 days).

Maintaining Laboratory Cultures of Gryllus bimaculatus, a Versatile Orthopteran Model for Insect Agriculture and Invertebrate Physiology

This paper outlines basic methods to standardize important factors such as density, feed availability, hydration source, and environmental controls for the long-term rearing of laboratory cultures of the edible cricket, Gryllus bimaculatus.

Gryllus bimaculatus (De Geer)&#160;is a large-bodied cricket distributed throughout Africa and Southern Eurasia where it is often wild-harvested as human ...

Artificial Rearing and Heavy Metal Tolerance in Cabbage White Butterflies

Rearing the Cabbage White Butterfly (Pieris rapae) in Controlled Conditions: A Case Study with Heavy Metal Tolerance

The cabbage white butterfly (Pieris rapae) is an important system for applied pest control research and basic research in behavioral and nutritional ecology. Cabbage whites can be easily reared in controlled conditions on an artificial diet, making them a model organism of the butterfly world. In this paper, a manipulation of heavy metal exposure is used to illustrate basic methods for rearing this species. The general protocol illustrates how butterflies can be caught in the field, induced to lay eggs in greenhouse cages, and transferred as larvae to artificial diets. The methods show how butterflies can be marked, measured, and studied for a variety of research questions. The representative results give an idea of how artificial diets that vary in components can be used to assess butterfly performance relative to a control diet. More specifically, butterflies were most tolerant to nickel and least tolerant to copper, with a tolerance of zinc somewhere in the middle. Possible explanations for these results are discussed, including nickel hyper-accumulation in some mustard host plants and recent evidence in insects that copper may be more toxic than previously appreciated. Finally, the discussion first reviews variations to the protocol and directions for troubleshooting these methods, before considering how future research might further optimize the artificial diet used in this study. Overall, by providing a detailed video overview of the rearing and measurement of cabbage whites on artificial diets, this protocol provides a resource for using this system across a wide range of studies.

Author Spotlight: Investigating the Tolerance of Cabbage Butterflies to Urban Pollutants 

This paper presents a detailed protocol for rearing the cabbage white butterfly in controlled lab conditions with an artificial diet, which allows precise manipulations of early-life nutrition and toxin exposure. The representative results show how heavy metal toxicity can be assayed with this protocol.

The cabbage white butterfly (Pieris rapae) is an important system for applied pest control research and basic research in behavioral and nutritional ...

Bradysia coprophila Lab Protocols and Chromosomal Research

Laboratory Maintenance of the Lower Dipteran Fly Bradysia (Sciara) coprophila: A New/Old Emerging Model Organism

Laboratory stocks of the lower dipteran fly, Bradysia (Sciara) coprophila, have been maintained for over a century. Protocols for laboratory upkeep of B. coprophila are presented here. These protocols will be useful for the rapidly increasing number of laboratories studying B. coprophila to take advantage of its unique biological features, which include (1) a monopolar spindle in male meiosis I; (2) non-disjunction of the X dyad in male meiosis II; (3) chromosome imprinting to distinguish maternal from paternal homologs; (4) germ line-limited (L) chromosomes; (5) chromosome elimination (paternal chromosomes in male meiosis I; one to two X chromosomes in early embryos; L chromosomes from the soma in early embryos); (6) sex determination by the mother (there is no Y chromosome); and (7) developmentally regulated DNA amplification at the DNA puff loci in larval salivary gland polytene chromosomes. 
It is now possible to explore these many unique features of chromosome mechanics by using the recent advances in sequencing and assembly of the B. coprophila genome and the development of transformation methodology for genomic engineering. The growing scientific community that uses B. coprophila for research will benefit from the protocols described here for mating the flies (phenotypic markers for mothers that will have only sons or only daughters; details of mass mating for biochemical experiments), checking embryo hatch, feeding larvae, and other comments on its rearing.

Author Spotlight: Exploring Bradysia coprophila's Unique Biology &#8211; A Guide to Laboratory Maintenance

This paper outlines the laboratory maintenance (including mating and feeding) of the lower dipteran fly Bradysia (Sciara) coprophila.

Laboratory stocks of the lower dipteran fly, Bradysia (Sciara) coprophila, have been maintained for over a century. Protocols for laboratory upkeep of B. ...

Rearing and Long-Term Maintenance of Eristalis tenax Hoverflies for Research Studies

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