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.


Introduction
Hoverflies are emerging as useful models for investigating a range of scientific questions, including flight behavior 1 , neural mechanisms underlying motion vision 2 , pollination efficiency 3,4,5,6 and microbial immunity 7 . However, as opposed to some other dipteran models, such as Drosophila 8 , 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 studies 9,10,11 . 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 groups 12,13 . There are about 6000 hoverfly species worldwide 14,15 , with more than 300 species in 75 genera in Sweden 16 and more than 300 species in 69 genera in India 17,18,19 . 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 Asia 6,16,17,18,19,20,21,22,23,24,25 . 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 hoverflies 26,27 . In the field, Eristalis can be found at any time of the year in Mediterranean climates 11 , 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 March 28 .
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 autumn 26,27 . 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 streams 10,11 . 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 matter 9,10,29,30,31,32,33 . However, larvae collection is limited by season, and is only a viable collection tool from late spring to the start of autumn . When working on vision, or other senses used in courtship and mating, we thus recommend maintaining genetic diversity, either by reestablishing 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 territories 34 . 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 hoverflies 35,36 . 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 production 37 , and reduced rates of male courtship behaviors 38 . 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.

Establish Captive E. tenax Colony
1. Establish colony via the collection of mature larvae (Step 1.2) or alternatively via the collection of free-flying hoverflies (Step 1.3).

Collection of Mature Larvae
1. 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. 2. Rear mature larvae in cow manure as outlined in Step 3.2.

Collection of Wild Hoverflies
1. 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. 2. House field collected hoverflies as outlined in Step 2. 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. 1. Leave hoverflies to feed for 6 hours at room temperature. 2. Place hoverflies with food and water in their housing, in the fridge at 8-10 °C and keep in complete darkness. NOTE: Storing hoverflies at 8-10 °C in darkness causes the hoverflies to enter a state of hibernation, with a reduction in activity and metabolic rate.

Housing and Long-Term Maintenance of Hoverflies
3. Every 3-4 days remove hoverflies from the fridge, thus breaking the artificial hibernation and allowing both feeding and grooming to occur. 1. 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. 2. 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.
4. Allow hoverflies to feed and groom at room temperature for 6 h. 5. Return hoverflies, with the food and water in their housing, into the fridge at 8-10 °C in complete darkness. This recommences the artificial hibernation of the hoverflies. 6. Continue the cyclical breaking of hibernation, every 3-4 days, to ensure the health and longevity of hoverflies for the duration of their captivity. 3. 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. 4. House both larvae and pupae at room temperature (21.5 ± 2.5 °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. 5. 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. 6. Allow emerging hoverflies to feed for 6 hours at room temperature before placing them in housing as outlined in Step 2.

Laboratory Rearing of Eggs laid by Gravid Wild Caught Females
1. Check hoverfly housing for eggs, both before changing housing (Step 2), and prior to returning them to the fridge. Oviposition by wildcaught gravid females occurs at both 8-10 °C and at room temperature. 1. 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.
2. Place hatched larvae in a 2.3 L bucket containing 1 L of fresh rabbit feces and 1 L of tap water. 1. Check slurry every 2-3 days and add additional tap water as required, to ensure that the slurry does not dry out before the 3 rd instar larvae emerge.
3. 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 3 rd instar larvae to crawl into the wood shavings and pupate.
1. Place a double layered mosquito net over the box to ensure that neither larvae nor any emerging hoverflies can escape.
4. House both larvae and pupae at room temperature (21.5 ± 2.5 °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. 5. Expect pupation to occur after 15-20 days. Collect pupae and place in an insect cage, allowing hoverflies to eclose there. 6. 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. 7. 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 °C. For this purpose, store larvae in the rabbit feces slurry and pupae in wood shavings.

Representative Results
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 m 3 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 ± 34 (mean ± 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 previously 39 . 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 ± 7 days for 5 females and 79 ± 4 days for 11 males). When the hoverflies were kept in artificial hibernation they could live in excess of 12 months.

Discussion
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 captivity 39 . 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 days 33 , 4 months 9 , 18 weeks 30 ). The main factor influencing this increased longevity is likely the use of artificial hibernation at 8-10 °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 see 39 ). 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 mold 9 .
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 suitable 9,29 . 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 captivity 39 . 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 spring 28 . Given that a typical female is capable of laying 3000 eggs in 60 days 29 , 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 diapause 28,40 . 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 °C for 2 weeks, although viability may be much longer. Indeed, Heal 30 reported a pupal duration increase of up to 37 days when the pupal temperature was dropped from 25 °C to 10 °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 ± 34 eclosed hoverflies from each clutch of eggs (N=3). Given that a typical female lays up to 200 eggs 40 , 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 growth 9,31,40,41 . However, there is no indication that the basis of the media greatly influences yield 32 . In addition, as opposed to feces from other vertebrates