Maintaining Aedes aegypti Mosquitoes Infected with Wolbachia

Summary

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.

Abstract

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.

Introduction

Aedes aegypti mosquitoes are responsible for transmitting some of the most important arboviruses in the world, including dengue, Zika and chikungunya¹. These viruses are becoming an increasing threat to global health as the widespread distribution of Ae. aegypti in the tropics continues to expand^2,3,4. Female Ae. aegypti preferentially feed on human blood⁵ and thus tend to live in close proximity to humans, particularly in urban areas where populations are most dense. Through this close association with humans they have also adapted to breed in artificial habitats, including tires, pots, gutters and water tanks^6,7. Ae. aegypti also readily adapt to laboratory environments where they can be maintained without any special requirements after being collected directly from the field, unlike some other species in the Aedes genus^8,9,10. Their ease of maintenance has seen them studied widely in the laboratory in a broad range of fields, ultimately aiming to control the diseases mosquitoes may transmit.

Traditionally, arboviral control relies heavily on the use of insecticides to reduce mosquito populations. However, there is increasing interest in approaches where modified mosquitoes are reared in the laboratory and then released into natural populations. Released mosquitoes may be modified genetically^11,12,13, biologically^14,15, through irradiation¹⁶, chemical treatment^17,18, or with combined techniques¹⁹ to either suppress populations of mosquitoes or replace them with mosquitoes that are refractory to arboviral transmission²⁰.

Wolbachia are bacteria that are currently being used as a biological control agent for arboviruses. Several strains of Wolbachia were recently introduced into Ae. aegypti experimentally using embryonic microinjection^21,22,23,24. These strains reduce the capacity of arboviruses to disseminate and replicate in the mosquito, diminishing their transmission potential^{23,25,26,27,28}. Wolbachia infections are transmitted from mother to offspring, however certain strains induce sterility when infected males mate with uninfected females²². Wolbachia-infected males can therefore be released in large quantities to suppress natural mosquito populations, as recently demonstrated in other Aedes species^15,29. However, since Wolbachia also inhibit arboviral transmission in Ae. aegypti, mosquitoes can also be released to replace native populations with poorer vectors. Ae. aegypti infected experimentally with Wolbachia are now being released into the field in several countries using this latter approach^14,30,31.

Wolbachia-based approaches for arboviral control rely on a sound understanding of the interactions between Wolbachia, the mosquito and the environment. Wolbachia occur naturally in a broad range of insects, and the strains introduced into mosquitoes are diverse in their effects³². As new Wolbachia infection types are introduced into Ae. aegypti²⁴, it is necessary to characterize each strain for their effects on mosquito fitness, reproduction and arboviral interference under a range of conditions. Rigorous experimentation in the laboratory is therefore required to evaluate the potential for Wolbachia strains to succeed in the field.

Open field releases of Ae. aegypti with Wolbachia infections can often require thousands to tens of thousands of mosquitoes per release zone to be reared each week^14,30,31. The success of initial releases can be improved by releasing mosquitoes of a large size to maximize their fecundity³³ and mating success^34,35. Mosquitoes should also be adapted to the conditions they will experience in the field, however long-term laboratory rearing may cause changes in behavior and physiology which could impact field performance^36,37,38.

We describe a simple protocol for rearing Ae. aegypti in the laboratory using basic equipment. This protocol is suitable for both wild-type and Wolbachia-infected mosquitoes, the latter of which can require special attention as some Wolbachia strains have substantial effects on mosquito life-history traits^39,40. The rearing conditions avoid overcrowding and competition for food to produce mosquitoes of a consistent size, which is critical for vector competence and fitness experiments, and ensures that the mosquitoes are healthy for field release⁴¹. We also take precautions to minimize laboratory adaptation and inbreeding by reducing selective pressures and ensuring that the next generation is sampled from a large, random pool. However, laboratory environments are distinctly different from field conditions, and long-term maintenance under relaxed conditions could reduce the fitness of mosquitoes upon release into the field^37,42,43. We therefore cross females from laboratory lines to field-collected males periodically, resulting in colonies that are genetically similar for experimental comparisons and that are adapted to the target field population³⁹. The methods do not require any specialized equipment and can be scaled up to rear tens of thousands of individuals per week for field releases. The protocol also prioritizes the fitness of mosquitoes within and across generations, an important consideration for insects destined for establishment in natural populations. The protocol is suitable for most laboratories that require maintenance of Ae. aegypti, particularly for experimental comparisons where a consistent quality of mosquitoes and relatability to the field are important.

Protocol

Blood feeding of mosquitoes on human subjects was approved by the University of Melbourne Human Ethics Committee (approval #: 0723847). All volunteers provided informed written consent.

1. Larval Rearing

NOTE: Mosquitoes are held at 26 ± 0.5 °C and 50-70% relative humidity, with a 12:12 h (light:dark) photoperiod for this colony maintenance protocol. These conditions are similar to the average climatic conditions in Cairns, Australia and within the optimal thermal range for Ae. aegypti survival and development^44,45,46. High temperatures can result in the loss of Wolbachia infections from mosquito colonies and should be avoided⁴⁷. We maintain at least 500 individuals per population to minimize inbreeding; maintaining colonies of a smaller size can have fitness consequences [Ross et al. unpublished]. Under these conditions and assuming adequate nutrition, the average generation time is 28 days (see Table 1).

1. Submerge the eggs on substrate in trays (Figure 1A) containing 3 L of water (reverse osmosis water or aged tap water, generated by leaving tap water in trays for 24 h prior to use), ~300 mg of fish food (one crushed tablet, see Table of Materials) and a few grains of active dry yeast to induce hatching⁴⁸.
2. One day after hatching, use a glass pipette to transfer approximately 500 larvae to trays containing 4 L of water (Figure 1B), count using a clicker counter. Add two crushed fish food tablets to each tray. If needed, use containers of different sizes for rearing larvae (Figure 1A), but keep larval densities below 0.5 larvae/mL to avoid overcrowding.
3. Check the trays daily to ensure that the larvae have sufficient food; add approximately two food tablets to the trays every two days. Provide food ad libitum but ensure that 0.5 mg/larva/day is available during this period to ensure development is synchronous and body size is consistent, otherwise the results of experiments may be confounded (see Representative Results).
4. Take care to avoid the overfeeding of larvae, particularly in smaller rearing containers with less water surface area and volume. If the water looks cloudy or if there is significant larval mortality, replace it with fresh water; mortality should be negligible if larvae are fed optimally.

2. Adult Emergence

NOTE: Larvae will begin to pupate from five days after hatching if well fed and the majority should pupate by seven days after hatching. Adults will begin emerging approximately two days after pupation if maintained optimally at 26 °C (see Representative Results). Larval development is typically unaffected by Wolbachia infections when ample food is provided^23,39,49.

1. Seven days after hatching pour the entire contents of the tray through a fine mesh (pore size 0.4 mm). Keep the filtered larval water for later use in ovicups (see "Blood Feeding and Oviposition" section). Invert the mesh and dip it into a plastic container with 200 mL of water to transfer the pupae. Provide additional food if any larvae remain.
2. Prepare the adult emergence cages (Figure 1C) by providing two cups of 10% sucrose solution (Figure 1F) and two cups of moist cotton wool to prevent desiccation (Figure 1E).
3. If the pupae do not need to be sorted by sex, place the lidded containers of pupae into the cage and leave the lid slightly ajar to allow the adults to emerge into the cage. Alternatively, place an inverted funnel over the container to minimize drowning. Ensure all adults have emerged before removing the container from the cage to prevent selecting against slow developers.

3. Pupal Sexing for Outcrossing

1. If the pupae do need to be sorted by sex (e.g., for outcrossing), pipette the pupae from larval trays and separate the sexes (Figure 2) into plastic containers (Figure 1A) with 200 mL of water every 24 h until the desired number of each sex has been reached. Place lids on the containers and leave them closed.
2. Adults will emerge into the containers; confirm their sex before releasing into cages (Figure 2C). Remove any adults sexed incorrectly with an aspirator within 24 h of emergence before they reach sexual maturity. Once the sexes have been confirmed, release the adults into cages every 24 h.
3. To obtain Wolbachia-infected colonies of a similar genetic background to a natural population, outcross by adding Wolbachia-infected females from laboratory colonies to cages of uninfected males derived from eggs collected by ovitraps in the field³⁹, maintaining the prescribed density of 500 individuals per population.
   1. Repeat the outcrossing for at least three consecutive generations to produce colonies that are at least 87.5% similar genetically to the field population³⁹. Critical: Ensure that the sexes are correct at this stage (see step 3.1).
4. Female Ae. aegypti are usually refractory to further insemination within hours of mating⁵⁰. When outcrossing colonies, allow the females and males to mature in separate cages for two days and then aspirate the females into the male cage to provide an equal opportunity to all males.

4. Blood Feeding and Oviposition

1. Wait for at least three days after the last female has emerged before blood feeding to allow ample time to mature. Blood feed the females within two weeks of emergence to prevent excessive mortality, particularly for mosquitoes with Wolbachia infections that adversely affect longevity^22,24,49. Remove the sugar cups the day prior to feeding to improve feeding rates.
   1. Ask a volunteer to insert their forearm into the cage to allow the female mosquitoes to feed. Most females should feed to repletion within 5 min, but to reduce selection against slow feeders, leave the forearm in the cage for 15 min, or until all females are visibly engorged; a latex glove to protect the hand from bites is optional but recommended.
   2. Two days after blood feeding, place two plastic cups containing larval rearing water and lined with a strip of sandpaper (Figure 1G) (or filter paper (Figure 1H)) into the cage for females to lay eggs. Partially submerge the sandpaper strip in the water to keep it moist. Remove other sources of water to prevent the females from laying their eggs outside the oviposition cup.
      NOTE: Tap water may be used in the cups, but larval-rearing water encourages oviposition^51,52 and females will lay their eggs more synchronously.

5. Egg Collection and Conditioning

1. The females will lay eggs on the sandpaper just above the water line; collect and replace the strips of sandpaper daily until no more eggs are laid. Note that oviposition may continue for up to one week.
2. Partially dry the sandpaper strips by gently blotting them on a paper towel for 30 s, taking care not to dislodge the eggs. Then, wrap the strips in a sheet of dry paper towel and place it into a sealable plastic bag (Figure 1I). 
3. Check the condition of the eggs under a dissecting microscope (Figure 3). If the sandpaper strips are too wet, eggs may hatch before being submerged in water (Figure 3B), but if dried too harshly, eggs may collapse (Figure 3C).
4. The eggs can be hatched at any time beyond three days post-collection; hatch all eggs from each colony, collected across all days, in the same container of water to ensure that the next generation is sampled from a large, random pool of individuals.
5. For long-term storage, keep the eggs in a sealed container at a high (> 80%) humidity at around 20 °C. Under these conditions, eggs without Wolbachia can be stored for several months while maintaining high hatch rates^53,54.
6. As some Wolbachia infections greatly reduce the viability of eggs with age^49,55, hatch the eggs from Wolbachia-infected lines within one week of collection to prevent excessive mortality for the relevant strains. Blood feed the females again after one week if more eggs are needed.

Day	Step
0	Hatch eggs
1	Count larvae into trays
7	Transfer larvae and pupae to colony cages
17	Blood feed female adults
21	Begin collecting eggs
25	Finish collecting eggs
28	Hatch eggs

Table 1: Overview of the Ae. aegypti colony maintenance schedule at 26 °C. The timing of the female blood-feeding and the hatching of eggs is flexible, but long durations at these stages should be avoided, particularly for mosquitoes infected with Wolbachia, in order to minimize mortality. Following this schedule minimizes selection against mosquitoes that are fast or slow to develop or mature at all life stages, provided that larvae are fed optimally.

Figure 1: Equipment used for rearing Ae. aegypti in the laboratory. (A) Plastic containers used for hatching eggs or rearing larvae with volumes of 500, 750 and 5,000 mL (from left to right). (B) Trays used for rearing larvae at a controlled density, usually 500 larvae in 4 L of water. (C) 19.7 L and (D) 3 L cages used for housing adults. A density of 25 adults or less per liter should be maintained to provide sufficient space. (E) 35 mL cup with moist cotton wool provided as a source of water to adults. (F) 35 mL cup with sucrose solution provided through a cord or dental wick as a source of sugar. (G-H) Cups filled with larval rearing water and lined with an oviposition substrate of sandpaper or filter paper (G and H, respectively). (I) Zip-lock bag used for storage of sandpaper strips or filter paper. Black spots on sandpaper are mosquito eggs. Please click here to view a larger version of this figure.

Figure 2: Lateral (A) and dorsal (B) views of pupae and adult Ae. aegypti (C) demonstrating their sexual dimorphism. Males are positioned on the left and females on the right in each panel. When optimally fed, male and female pupae are distinguished by size; females are larger than males (A) and have a relatively bulbous cephalothorax compared to males which have flatter sides (B). Male adults are easily distinguished from females under all rearing conditions, primarily by their plumose antennae and long palps. Please click here to view a larger version of this figure.

Figure 3. Four day old Ae. aegypti eggs under different conditions. (A) Intact eggs on sandpaper strips maintained at a high (> 80%) humidity but without any visible moisture. Hatch rates should be above 90% for wild-type Ae. aegypti if maintained correctly. (B) Eggs that hatch before being submerged in water (precocious hatching) are distinguished by a detached egg cap and visible larva. This indicates that the sandpaper strip was kept too moist. (C) Eggs that are dried too harshly may collapse, and are clearly visible by their concave appearance. If the sandpaper becomes stiff this also indicates that the eggs may be too dry. Please click here to view a larger version of this figure.

Results

Figure 4 demonstrates the effects of suboptimal nutrition on the development of Ae. aegypti larvae. When containers are provided with 0.25 mg of food per larva per day or less, the development time increases for both males and females, and is less synchronous than in containers provided with 0.5 mg of food. If adequate food is not provided throughout the duration of larval development, this could have an adverse impact on the maintenance schedule. Sl...

Discussion

Following the protocol presented here for the maintenance of Wolbachia-infected Ae. aegypti should ensure that healthy mosquitoes of a consistent quality are produced for experiments and open field releases. In contrast to other protocols that prioritize the production of mass quantities of mosquitoes (see reference⁵⁷), the methods are focused on maximizing their fitness, both within generations by implementing relaxed rearing conditions, and across generations by minimizing inbr...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Wild type Aedes aegypti			Collected from field locations in Queensland, Australia, see Yeap and others39 for details
w Mel-infected Aedes aegypti			Provided by Monash University. Refer to Walker and others23 for information on the strain
w AlbB-infected Aedes aegypti			Provided by Monash University. Refer to Xi and others21 for information on the strain
w MelPop-infected Aedes aegypti			Provided by Monash University. Refer to McMeniman and others22 for information on the strain
Instant dried yeast	Lowan		Stimulates egg hatching. Found in general grocery stores. Other brands may be used
5 L plastic tub	Quadrant	Q110950	Used for hatching and rearing larvae. Other products may be used
Fish Food (Tetramin Tropical Tablets)	Tetra	16152	Provided to larvae as a source of food. Web address: https://www.amazon.com/Tetra-16152-TetraMin-Tropical-10-93-Ounce/dp/B00025Z6SE
Plastic containers			Used for rearing larvae. Any plastic container above 500 mL should be suitable
Glass pipette			Used for transferring larvae and pupae between containers. Web address: https://www.aliexpress.com/item/10Pcs-Durable-Long-Glass-Experiment-Medical-Pipette-Dropper-Transfer-Pipette-Lab-Supplies-With-Red-Rubber-Cap/32704471109.html?spm=2114.40010308.4.2.py4Kez
Clicker counter	RS Pro	710-5212	Used to assist in the counting of larvae, pupae and eggs. Web address: http://au.rs-online.com/web/p/products/7105212/?grossPrice=Y
Rearing trays	Gratnells		Used for rearing larvae. Web address: http://www.gratnells.com
Nylon mesh			Used to transfer larvae and pupae to containers of fresh water. Other brands may be used. Web address: https://www.spotlightstores.com/fabrics-yarn/specialty-apparel-fabrics/nettings-tulles/nylon-netting/p/BP80046941001-white
Cages	BugDorm	DP1000	Houses adult mosquitoes. Alternative products may be used. Web address: http://bugdorm.megaview.com.tw/bugdorm-1-insect-rearing-cage-30x30x30-cm-pack-of-one-p-29.html
35 mL plastic cup	Huhtamaki	AA272225	Used to provide water or sucrose to adult mosquitoes. Other brands may be used
35 mL plastic cup lid	Huhtamaki	GB030005	Used to provide sucrose to adult mosquitoes. Other brands may be used
Cotton wool	Cutisoft	71841-13	Moist cotton wool is provided as a source of water to adults. Other brands may be used
White Sugar			Provided as a source of sugar to adult mosquitoes. Found in general grocery stores
Rope	M Recht Accessories	C323C/W	Used to provide sucrose solution to adults. Other brands may be used. Web address: https://mrecht.com.au/haberdashery/braids-cords-and-tapes/cords/plaited-cord/cotton/
Plastic cup (large)			Used as an oviposition container. Any plastic cup that holds 100 mL of water should be suitable
Sandpaper	Norton Master Painters	CE015962	Provided as an oviposition substrate. Alternative products may be used, but we use this brand because it is relatively odorless. Lighter colors are used for contrast with eggs. Web address: https://www.bolt.com.au/115mm-36m-master-painters-bulk-roll-p80-medium-p-9396.html
Filter paper	Whatman	1001-150	Used as an alternative oviposition substrate. Other brands may be used
Latex gloves	SemperGuard	Z560979	Prevents mosquito bites on hands when blood feeding. Other brands may be used. Web address: http://www.sempermed.com/en/products/detail/semperguardR_latex_puderfrei_innercoated/