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 border="0" cellpadding="0" cellspacing="0" style="width:903px;" width="902">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Bee Pollen</td>
			<td style="width:211px;">Forest Super Foods</td>
			<td style="width:119px;"></td>
			<td style="width:357px;">any brand of bee pollen is suitable</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Honey</td>
			<td style="width:211px;">Bramwells</td>
			<td style="width:119px;"></td>
			<td>any brand of liquid honey is suitable</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Rabbit Faeces</td>
			<td style="width:211px;"></td>
			<td style="width:119px;"></td>
			<td>can be substituted with cow or pig manure made into a slurry</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">BugDome</td>
			<td style="width:211px;">Australia Entomological Supplies</td>
			<td style="width:119px;">EM42222</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Plastic Bags</td>
			<td style="width:211px;">Woolworths Homebrand</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Mosquito netting</td>
			<td style="width:211px;">Clas Ohlson</td>
			<td>34-1113</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Cotton Balls</td>
			<td style="width:211px;">Woolworths Select</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Fridge</td>
			<td>Hisense</td>
			<td style="width:119px;"></td>
			<td>fridge needs to maintain a stable 8-10&#176;C&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Buckets (2-3L)</td>
			<td style="width:211px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Large plastic tubs (30L)</td>
			<td style="width:211px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Wood shavings</td>
			<td style="width:211px;">Pollards Sawdust Supplies</td>
			<td>MaxiFlake (75)&#160;</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Bag clips</td>
			<td style="width:211px;">IKEA</td>
			<td style="width:119px;">Bevara 303.391.70</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Petri Dish (100mm x 20mm)</td>
			<td style="width:211px;">Corning</td>
			<td style="width:119px;">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 ...

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13.3K Views.  Flinders University. The overall goal of these procedures is to establish, maintain and refresh a captive population of Eristalis tenax in a research setting.

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

Waste Water Derived Electroactive Microbial Biofilms: Growth, Maintenance, and Basic Characterization

The growth of anodic electroactive microbial biofilms from waste water inocula in a fed-batch reactor is demonstrated using a three-electrode setup controlled by a potentiostat. Thereby the use of potentiostats allows an exact adjustment of the electrode potential and ensures reproducible microbial culturing conditions. During growth the current production is monitored using chronoamperometry (CA). Based on these data the maximum current density (jmax) and the coulombic efficiency (CE) are discussed as measures for characterization of the bioelectrocatalytic performance. Cyclic voltammetry (CV), a nondestructive, i.e. noninvasive, method, is used to study the extracellular electron transfer (EET) of electroactive bacteria. CV measurements are performed on anodic biofilm electrodes in the presence of the microbial substrate, i.e. turnover conditions, and in the absence of the substrate, i.e. nonturnover conditions, using different scan rates. Subsequently, data analysis is exemplified and fundamental thermodynamic parameters of the microbial EET are derived and explained: peak potential (Ep), peak current density (jp), formal potential (Ef) and peak separation (&#916;Ep). Additionally the limits of the method and the state-of the art data analysis are addressed. Thereby this video-article shall provide a guide for the basic experimental steps and the fundamental data analysis.

Chronoamperometric growth of anodic electroactive microbial biofilms in a fed-batch reactor using a three-electrode setup, controlled by a potentiostat, is demonstrated. The extracellular electron transfer is characterized using cyclic voltammetry in the presence of the electron donor and in the absence of the electron donor. Fundamental data analysis is demonstrated.

The growth of anodic electroactive microbial biofilms from waste water inocula in a fed-batch reactor is demonstrated using a three-electrode setup ...

Design and Construction of an Urban Runoff Research Facility

As the urban population increases, so does the area of irrigated urban landscape. Summer water use in urban areas can be 2-3x winter base line water use due to increased demand for landscape irrigation. Improper irrigation practices and large rainfall events can result in runoff from urban landscapes which has potential to carry nutrients and sediments into local streams and lakes where they may contribute to eutrophication. A 1,000 m2 facility was constructed which consists of 24 individual 33.6 m2 field plots, each equipped for measuring total runoff volumes with time and collection of runoff subsamples at selected intervals for quantification of chemical constituents in the runoff water from simulated urban landscapes. Runoff volumes from the first and second trials had coefficient of variability (CV) values of 38.2 and 28.7%, respectively. CV values for runoff pH, EC, and Na concentration for both trials were all under 10%. Concentrations of DOC, TDN, DON, PO4-P, K+, Mg2+, and Ca2+ had CV values less than 50% in both trials. Overall, the results of testing performed after sod installation at the facility indicated good uniformity between plots for runoff volumes and chemical constituents. The large plot size is sufficient to include much of the natural variability and therefore provides better simulation of urban landscape ecosystems.

This paper describes the design, construction, and function of a 1,000 m2 facility containing 24 individual 33.6 m2 field plots equipped for measuring total runoff volumes with time and collection of runoff subsamples at selected intervals for quantification of chemical constituents in the runoff water from simulated home lawns.

As the urban population increases, so does the area of irrigated urban landscape. Summer water use in urban areas can be 2-3x winter base line water use ...

Physical, Chemical and Biological Characterization of Six Biochars Produced for the Remediation of Contaminated Sites

The physical and chemical properties of biochar vary based on feedstock sources and production conditions, making it possible to engineer biochars with specific functions (e.g. carbon sequestration, soil quality improvements, or contaminant sorption). In 2013, the International Biochar Initiative (IBI) made publically available their Standardized Product Definition and Product Testing Guidelines (Version 1.1) which set standards for physical and chemical characteristics for biochar. Six biochars made from three different feedstocks and at two temperatures were analyzed for characteristics related to their use as a soil amendment. The protocol describes analyses of the feedstocks and biochars and includes: cation exchange capacity (CEC), specific surface area (SSA), organic carbon (OC) and moisture percentage, pH, particle size distribution, and proximate and ultimate analysis. Also described in the protocol are the analyses of the feedstocks and biochars for contaminants including polycyclic aromatic hydrocarbons (PAHs), polychlorinated biphenyls (PCBs), metals and mercury as well as nutrients (phosphorous, nitrite and nitrate and ammonium as nitrogen). The protocol also includes the biological testing procedures, earthworm avoidance and germination assays. Based on the quality assurance / quality control (QA/QC) results of blanks, duplicates, standards and reference materials, all methods were determined adequate for use with biochar and feedstock materials. All biochars and feedstocks were well within the criterion set by the IBI and there were little differences among biochars, except in the case of the biochar produced from construction waste materials. This biochar (referred to as Old biochar) was determined to have elevated levels of arsenic, chromium, copper, and lead, and failed the earthworm avoidance and germination assays. Based on these results, Old biochar would not be appropriate for use as a soil amendment for carbon sequestration, substrate quality improvements or remediation.

Biochar is a carbon-rich material used as a soil amendment with the ability to sustainably sequester carbon, improve substrate quality and sorb contaminants. This protocol describes the 17 analytical methods used for the characterization of biochar, which is required prior to large scale implementation of these amendments in the environment.

The physical and chemical properties of biochar vary based on feedstock sources and production conditions, making it possible to engineer biochars with ...

Administering and Detecting Protein Marks on Arthropods for Dispersal Research

Monitoring arthropod movement is often required to better understand associated population dynamics, dispersal patterns, host plant preferences, and other ecological interactions. Arthropods are usually tracked in nature by tagging them with a unique mark and then re-collecting them over time and space to determine their dispersal capabilities. In addition to actual physical tags, such as colored dust or paint, various types of proteins have proven very effective for marking arthropods for ecological research. Proteins can be administered internally and/or externally. The proteins can then be detected on recaptured arthropods with a protein-specific enzyme-linked immunosorbent assay (ELISA). Here we describe protocols for externally and internally tagging arthropods with protein. Two simple experimental examples are demonstrated: (1) an internal protein mark introduced to an insect by providing a protein-enriched diet and (2) an external protein mark topically applied to an insect using a medical nebulizer. We then relate a step-by-step guide of the sandwich and indirect ELISA methods used to detect protein marks on the insects. In this demonstration, various aspects of the acquisition and detection of protein markers on arthropods for mark-release-recapture, mark-capture, and self-mark-capture types of research are discussed, along with the various ways that the immunomarking procedure has been adapted to suit a wide variety of research objectives.

Proteins are often used to mark arthropods for dispersal research. Methods for administering internal and external protein marks on arthropods are demonstrated. Protein mark detection techniques, either through indirect or sandwich enzyme-linked immunosorbent assays (ELISAs), are also illustrated.

Monitoring arthropod movement is often required to better understand associated population dynamics, dispersal patterns, host plant preferences, and other ...

Clean Sampling and Analysis of River and Estuarine Waters for Trace Metal Studies

Most of the trace metal concentrations in ambient waters obtained a few decades ago have been considered unreliable owing to the lack of contamination control. Developments of some techniques aiming to reduce trace metal contamination in the last couple of decades have resulted in concentrations reported now being orders of magnitude lower than those in the past. These low concentrations often necessitate preconcentration of water samples prior to instrumental analysis of samples. Since contamination can appear in all phases of trace metal analyses, including sample collection (and during preparation of sampling containers), storage and handling, pretreatments, and instrumental analysis, specific care needs to be taken in order to reduce contamination levels at all steps. The effort to develop and utilize "clean techniques" in trace metal studies allows scientists to investigate trace metal distributions and chemical and biological behavior in greater details. This advancement also provides the required accuracy and precision of trace metal data allowing for environmental conditions to be related to trace metal concentrations in aquatic environments.
This protocol that is presented here details needed materials for sample preparation, sample collection, sample pretreatment including preconcentration, and instrumental analysis. By reducing contamination throughout all phases mentioned above for trace metal analysis, much lower detection limits and thus accuracy can be achieved. The effectiveness of "clean techniques" is further demonstrated using low field blanks and good recoveries for standard reference material. The data quality that can be obtained thus enables the assessment of trace metal distributions and their relationships to environmental parameters.

Special care using "clean techniques" is required to properly collect and process water samples for trace metal studies in aquatic environments. A protocol for sampling, processing, and analytical procedures with the aim of obtaining reliable environmental monitoring data and results with high sensitivity for detailed trace metal studies is presented.

Most of the trace metal concentrations in ambient waters obtained a few decades ago have been considered unreliable owing to the lack of contamination ...

Use of a Filter Cartridge for Filtration of Water Samples and Extraction of Environmental DNA

Recent studies demonstrated the use of environmental DNA (eDNA) from fishes to be appropriate as a non-invasive monitoring tool. Most of these studies employed disk fiber filters to collect eDNA from water samples, although a number of microbial studies in aquatic environments have employed filter cartridges, because the cartridge has the advantage of accommodating large water volumes and of overall ease of use. Here we provide a protocol for filtration of water samples using the filter cartridge and extraction of eDNA from the filter without having to cut open the housing. The main portions of this protocol consists of 1) filtration of water samples (water volumes &#8804;4 L or &gt;4 L); (2) extraction of DNA on the filter using a roller shaker placed in a preheated incubator; and (3) purification of DNA using a commercial kit. With the use of this and previously-used protocols, we perform metabarcoding analysis of eDNA taken from a huge aquarium tank (7,500 m3) with known species composition, and show the number of detected species per library from the two protocols as the representative results. This protocol has been developed for metabarcoding eDNA from fishes, but is also applicable to eDNA from other organisms.

We describe a protocol for filtration of water samples with a filter cartridge and extraction of environmental DNA (eDNA) without having to cut open the housing to remove the filter. This protocol is developed for metabarcoding eDNA from fishes, but is also applicable to eDNA from other organisms.

Recent studies demonstrated the use of environmental DNA (eDNA) from fishes to be appropriate as a non-invasive monitoring tool. Most of these studies ...

RGB and Spectral Root Imaging for Plant Phenotyping and Physiological Research: Experimental Setup and Imaging Protocols

Better understanding of plant root dynamics is essential to improve resource use efficiency of agricultural systems and increase the resistance of crop cultivars against environmental stresses. An experimental protocol is presented for RGB and hyperspectral imaging of root systems. The approach uses rhizoboxes where plants grow in natural soil over a longer time to observe fully developed root systems. Experimental settings are exemplified for assessing rhizobox plants under water stress and studying the role of roots. An RGB imaging setup is described for cheap and quick quantification of root development over time. Hyperspectral imaging improves root segmentation from the soil background compared to RGB color based thresholding. The particular strength of hyperspectral imaging is the acquisition of chemometric information on the root-soil system for functional understanding. This is demonstrated with high resolution water content mapping. Spectral imaging however is more complex in image acquisition, processing and analysis compared to the RGB approach. A combination of both methods can optimize a comprehensive assessment of the root system. Application examples integrating root and aboveground traits are given for the context of plant phenotyping and plant physiological research. Further improvement of root imaging can be obtained by optimizing RGB image quality with better illumination using different light sources and by extension of image analysis methods to infer on root zone properties from spectral data.

An experimental protocol is presented for assessment of soil grown plant root systems with RGB and hyperspectral imaging. Combination of RGB image time series with chemometric information from hyperspectral scans optimizes insights into plant root dynamics.

Better understanding of plant root dynamics is essential to improve resource use efficiency of agricultural systems and increase the resistance of crop ...

In Vitro Rearing of Solitary Bees: A Tool for Assessing Larval Risk Factors

Although solitary bees provide crucial pollination services for wild and managed crops, this species-rich group has been largely overlooked in pesticide regulation studies. The risk of exposure to fungicide residues is likely to be especially high if the spray occurs on, or near host plants while the bees are collecting pollen to provision their nests. For species of Osmia that consume pollen from a select group of plants (oligolecty), the inability to use pollen from non-host plants can increase their risk factor for fungicide-related toxicity. This manuscript describes protocols used to successfully rear oligolectic mason bees, Osmia ribifloris sensu lato, from egg to prepupal stage within cell culture plates under standardized laboratory conditions. The in vitro-reared bees are subsequently used to investigate the effects of fungicide exposure and pollen source on bee fitness. Based on a 2 &#215; 2 fully crossed factorial design, the experiment examines the main and interactive effects of fungicide exposure and pollen source on larval fitness, quantified by prepupal biomass, larval developmental time, and survivorship. A major advantage of this technique is that using in vitro-reared bees reduces natural background variability and allows the simultaneous manipulation of multiple experimental parameters. The described protocol presents a versatile tool for hypotheses testing involving the suite of factors affecting bee health. For conservation efforts to be met with significant, lasting success, such insights into the complex interplay of physiological and environmental factors driving bee declines will prove to be critical.

In Vitro Rearing of Solitary Bees: A Tool for Assessing Larval Risk Factors

Fungicide sprays on flowering plants may expose solitary bees to high concentrations of pollen-borne fungicide residues. Using laboratory-based experiments involving in vitro-reared bee larvae, this study investigates the interactive effects of consuming fungicide-treated pollen derived from host and non-host plants.

Although solitary bees provide crucial pollination services for wild and managed crops, this species-rich group has been largely overlooked in pesticide ...

Experimental Protocol for Examining Behavioral Response Profiles in Larval Fish: Application to the Neuro-stimulant Caffeine

Fish models and behaviors are increasingly used in the biomedical sciences; however, fish have long been the subject of ecological, physiological and toxicological studies. Using automated digital tracking platforms, recent efforts in neuropharmacology are leveraging larval fish locomotor behaviors to identify potential therapeutic targets for novel small molecules. Similar to these efforts, research in the environmental sciences and comparative pharmacology and toxicology is examining various behaviors of fish models as diagnostic tools in tiered evaluation of contaminants and real-time monitoring of surface waters for contaminant threats. Whereas the zebrafish is a popular larval fish model in the biomedical sciences, the fathead minnow is a common larval fish model in ecotoxicology. Unfortunately, fathead minnow larvae have received considerably less attention in behavioral studies. Here, we develop and demonstrate a behavioral profile protocol using caffeine as a model neurostimulant. Though photomotor responses of fathead minnows were occasionally affected by caffeine, zebrafish&#160;were markedly more sensitive for photomotor and locomotor endpoints, which responded at environmentally relevant levels. Future studies are needed to understand comparative behavioral sensitivity differences among fish with age and time of day, and to determine whether similar behavioral effects would occur in nature and be indicative of adverse outcomes at the individual or population levels of biological organization.

Here, we present a protocol to examine larval zebrafish and fathead minnow locomotor activities and photomotor responses (PMR) using an automated tracking software. When incorporated in common toxicology bioassays, analyses of these behaviors provide a diagnostic tool to examine chemical bioactivity. This protocol is described using caffeine, a model neurostimulant.

Fish models and behaviors are increasingly used in the biomedical sciences; however, fish have long been the subject of ecological, physiological and ...

Long-term Video Tracking of Cohoused Aquatic Animals: A Case Study of the Daily Locomotor Activity of the Norway Lobster (Nephrops norvegicus)

We present a protocol related to a video-tracking technique based on the background subtraction and image thresholding that makes it possible to individually track cohoused animals. We tested the tracking routine with four cohoused Norway lobsters (Nephrops norvegicus) under light-darkness conditions for 5 days. The lobsters had been individually tagged. The experimental setup and the tracking techniques used are entirely based on the open source software. The comparison of the tracking output with a manual detection indicates that the lobsters were correctly detected 69% of the times. Among the correctly detected lobsters, their individual tags were correctly identified 89.5% of the times. Considering the frame rate used in the protocol and the movement rate of lobsters, the performance of the video tracking has a good quality, and the representative results support the validity of the protocol in producing valuable data for research needs (individual space occupancy or locomotor activity patterns). The protocol presented here can be easily customized and is, hence, transferable to other species where the individual tracking of specimens in a group can be valuable for answering research questions.

Long-term Video Tracking of Cohoused Aquatic Animals: A Case Study of the Daily Locomotor Activity of the Norway Lobster Nephrops norvegicus

Here we present a protocol to individually track animals over a long period of time. It uses computer vision methods to identify a set of manually constructed tags by using a group of lobsters as case study, simultaneously providing information on how to house, manipulate, and mark the lobsters.

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

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Chapters in this video

Waste Water Derived Electroactive Microbial Biofilms: Growth, Maintenance, and Basic Characterization

Design and Construction of an Urban Runoff Research Facility

Physical, Chemical and Biological Characterization of Six Biochars Produced for the Remediation of Contaminated Sites

Administering and Detecting Protein Marks on Arthropods for Dispersal Research

Clean Sampling and Analysis of River and Estuarine Waters for Trace Metal Studies

Use of a Filter Cartridge for Filtration of Water Samples and Extraction of Environmental DNA

RGB and Spectral Root Imaging for Plant Phenotyping and Physiological Research: Experimental Setup and Imaging Protocols

In Vitro Rearing of Solitary Bees: A Tool for Assessing Larval Risk Factors

Experimental Protocol for Examining Behavioral Response Profiles in Larval Fish: Application to the Neuro-stimulant Caffeine

Long-term Video Tracking of Cohoused Aquatic Animals: A Case Study of the Daily Locomotor Activity of the Norway Lobster (Nephrops norvegicus)