We acknowledge Heng Lin Yeap, Chris Paton, Petrina Johnson and Clare Doig for their contributions to the development of our colony maintenance methods, and three anonymous reviewers for their suggestions which helped to improve the manuscript. Our research is supported by a program grant and fellowship to AAH from the National Health and Medical Research Council and a translation grant from the Wellcome Trust. PAR is a recipient of an Australian Government Research Training Program Scholarship.

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 &#177; 0.5 &#176;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 development44,45,46. 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 unpublished].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).
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>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&#160;during this period to ensure development is synchronous and body size is consistent, otherwise the results of experiments may be confounded (see Representative Results).</li>
	<li>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.</li>
</ol>
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 &#176;C (see Representative Results). Larval development is typically unaffected by Wolbachia infections when ample food is provided23,39,49.
<ol>
	<li>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 &#34;Blood Feeding and Oviposition&#34; 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.</li>
	<li>Prepare the adult emergence cages (Figure 1C) by providing two cups of 10% sucrose solution (Figure 1F) and two cups of moist cotton&#160;wool to prevent desiccation (Figure 1E).</li>
	<li>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.</li>
</ol>
3. Pupal Sexing for Outcrossing
<ol>
	<li>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.</li>
	<li>Adults will emerge into the containers; confirm their sex&#160;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.</li>
	<li>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.
	<ol>
		<li>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).</li>
	</ol>
	</li>
	<li>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.</li>
</ol>
4. Blood Feeding and Oviposition
<ol>
	<li>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 longevity22,24,49. Remove the sugar cups the day prior to feeding to improve feeding rates.
	<ol>
		<li>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.</li>
		<li>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 oviposition51,52 and females will lay their eggs more synchronously.</li>
	</ol>
	</li>
</ol>
5. Egg Collection and Conditioning
<ol>
	<li>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.</li>
	<li>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).&#160;</li>
	<li>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).</li>
	<li>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.</li>
	<li>For long-term storage, keep the eggs in a sealed container at a high (&#62; 80%) humidity at around 20 &#176;C. Under these conditions, eggs without Wolbachia can be stored&#160;for several months while maintaining high hatch rates53,54.</li>
	<li>As some Wolbachia infections greatly reduce the viability of eggs with age49,55, 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.</li>
</ol>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Day</td>
			<td colspan="3">Step</td>
		</tr>
		<tr>
			<td>0</td>
			<td colspan="3">Hatch eggs</td>
		</tr>
		<tr>
			<td>1</td>
			<td colspan="3">Count larvae into trays</td>
		</tr>
		<tr>
			<td>7</td>
			<td colspan="3">Transfer larvae and pupae to colony cages</td>
		</tr>
		<tr>
			<td>17</td>
			<td colspan="3">Blood feed female adults</td>
		</tr>
		<tr>
			<td>21</td>
			<td colspan="3">Begin collecting eggs</td>
		</tr>
		<tr>
			<td>25</td>
			<td colspan="3">Finish collecting eggs</td>
		</tr>
		<tr>
			<td>28</td>
			<td colspan="3">Hatch eggs</td>
		</tr>
	</tbody>
</table>
Table 1: Overview of the Ae. aegypti colony maintenance schedule at 26 &#176;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.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56124/56124fig1.jpg" /> 
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. <a href="//cloudfront.jove.com/files/ftp_upload/56124/56124fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56124/56124fig2.jpg" /> 
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. <a href="//cloudfront.jove.com/files/ftp_upload/56124/56124fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56124/56124fig3.jpg" /> 
Figure 3. Four day old Ae. aegypti eggs under different conditions. (A) Intact eggs on sandpaper strips maintained at a high (&#62; 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. <a href="//cloudfront.jove.com/files/ftp_upload/56124/56124fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

The authors declare that they have no competing financial interests.

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...

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 expand2,3,4. 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 tanks6,7. 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 genus8,9,10. 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 genetically11,12,13, biologically14,15, through irradiation16, chemical treatment17,18, 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 microinjection21,22,23,24. These strains reduce the capacity of arboviruses to disseminate and replicate in the mosquito, diminishing their transmission potential23,25,26,27,28. 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 species15,29. 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 approach14,30,31.
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 week14,30,31. The success of initial releases can be improved by releasing mosquitoes of a large size to maximize their fecundity33 and mating success34,35. 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 performance36,37,38.
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 traits39,40. 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 field37,42,43. 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.

<table width="1661">
	<tbody>
		<tr height="24">
			<td height="24" style="width:287px;height:24px;">Wild type Aedes aegypti</td>
			<td style="width:168px;"></td>
			<td style="width:119px;"></td>
			<td style="width:1087px;">Collected from field locations in Queensland, Australia, see Yeap and others39 for details</td>
		</tr>
		<tr height="24">
			<td height="24" style="width:287px;height:24px;">w Mel-infected Aedes aegypti</td>
			<td style="width:168px;"></td>
			<td style="width:119px;"></td>
			<td>Provided by Monash University. Refer to Walker and others23 for information on the strain</td>
		</tr>
		<tr height="24">
			<td height="24" style="width:287px;height:24px;">w AlbB-infected Aedes aegypti</td>
			<td style="width:168px;"></td>
			<td style="width:119px;"></td>
			<td>Provided by Monash University. Refer to Xi and others21 for information on the strain</td>
		</tr>
		<tr height="24">
			<td height="24" style="width:287px;height:24px;">w MelPop-infected Aedes aegypti</td>
			<td style="width:168px;"></td>
			<td style="width:119px;"></td>
			<td>Provided by Monash University. Refer to McMeniman and others22 for information on the strain</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:287px;height:21px;">Instant dried yeast</td>
			<td style="width:168px;">Lowan</td>
			<td style="width:119px;"></td>
			<td>Stimulates egg hatching. Found in general grocery stores. Other brands may be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:287px;height:21px;">5 L plastic tub</td>
			<td style="width:168px;">Quadrant</td>
			<td style="width:119px;">Q110950</td>
			<td>Used for hatching and rearing larvae. Other products may be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:287px;height:21px;">Fish Food (Tetramin Tropical Tablets)</td>
			<td style="width:168px;">Tetra</td>
			<td style="width:119px;">16152</td>
			<td>Provided to larvae as a source of food. Web address: https://www.amazon.com/Tetra-16152-TetraMin-Tropical-10-93-Ounce/dp/B00025Z6SE</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:287px;height:21px;">Plastic containers</td>
			<td style="width:168px;"></td>
			<td style="width:119px;"></td>
			<td>Used for rearing larvae. Any plastic container above 500 mL should be suitable</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:287px;height:21px;">Glass pipette</td>
			<td style="width:168px;"></td>
			<td style="width:119px;"></td>
			<td>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</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:287px;height:21px;">Clicker counter</td>
			<td style="width:168px;">RS Pro</td>
			<td style="width:119px;">710-5212</td>
			<td>Used to assist in the counting of larvae, pupae and eggs. Web address: http://au.rs-online.com/web/p/products/7105212/?grossPrice=Y</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:287px;height:21px;">Rearing trays</td>
			<td style="width:168px;">Gratnells</td>
			<td style="width:119px;"></td>
			<td>Used for rearing larvae. Web address: http://www.gratnells.com</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:287px;height:21px;">Nylon mesh</td>
			<td style="width:168px;"></td>
			<td style="width:119px;"></td>
			<td>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</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:287px;height:21px;">Cages</td>
			<td style="width:168px;">BugDorm</td>
			<td>DP1000</td>
			<td>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</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:287px;height:21px;">35 mL plastic cup</td>
			<td style="width:168px;">Huhtamaki</td>
			<td style="width:119px;">AA272225</td>
			<td>Used to provide water or sucrose to adult mosquitoes. Other brands may be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:287px;height:21px;">35 mL plastic cup lid</td>
			<td style="width:168px;">Huhtamaki</td>
			<td>GB030005</td>
			<td>Used to provide sucrose to adult mosquitoes. Other brands may be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:287px;height:21px;">Cotton wool</td>
			<td style="width:168px;">Cutisoft</td>
			<td style="width:119px;">71841-13</td>
			<td>Moist cotton wool is provided as a source of water to adults. Other brands may be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:287px;height:21px;">White Sugar</td>
			<td style="width:168px;"></td>
			<td style="width:119px;"></td>
			<td>Provided as a source of sugar to adult mosquitoes. Found in general grocery stores</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:287px;height:21px;">Rope</td>
			<td style="width:168px;">M Recht Accessories</td>
			<td>C323C/W</td>
			<td>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/</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:287px;height:21px;">Plastic cup (large)</td>
			<td style="width:168px;"></td>
			<td style="width:119px;"></td>
			<td>Used as an oviposition container. Any plastic cup that holds 100 mL of water should be suitable</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:287px;height:21px;">Sandpaper</td>
			<td style="width:168px;">Norton Master Painters</td>
			<td>CE015962</td>
			<td>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</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:287px;height:21px;">Filter paper</td>
			<td style="width:168px;">Whatman</td>
			<td style="width:119px;">1001-150</td>
			<td>Used as an alternative oviposition substrate. Other brands may be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="width:287px;height:21px;">Latex gloves</td>
			<td style="width:168px;">SemperGuard</td>
			<td>Z560979</td>
			<td>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/</td>
		</tr>
	</tbody>
</table>

maintaining aedes aegypti mosquitoes infected with wolbachia

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

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...

Watch this Scientific Journal Video about Maintaining Aedes aegypti Mosquitoes Infected with Wolbachia at JoVE.com

Maintaining Aedes aegypti Mosquitoes Infected with Wolbachia

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

Maintaining Aedes aegypti Mosquitoes Infected with Wolbachia

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

maintaining-aedes-aegypti-mosquitoes-infected-with-wolbachia

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14.1K Views.  Bio21 Institute and University of Melbourne. 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.

Video: Maintaining Aedes aegypti Mosquitoes Infected with Wolbachia

Wolbachia Bacterial Infection in Drosophila

My name is Voia Friedman. I am associate researcher at Princeton University. And today I'm going to tell you about a few protocols that I developed to trans infect, to I initially to isolate, to go back from the heli of infected flies, and later on help to inject the heli feed to unaffected flights.So basically, this is a very simple protocol to isolate him oly for basics. And you will use this device from, from mepo that contains a filter that goes on top of an E.So you basically needs to put you, we start from around a hundred flies. You anize the flies, you put them into the filter device, you can just use a funnel and that will add our other fit very easily on this device and wash it with 70%ethanol.You just take the, take the future away, you can throw away the ethanol, and then after three times your last wash you do with molecule water because you don't want have the bacteria from the, with traces of ethanol. Well, and then you just go to a new weapon door and, and then you spin down on a micro, real micro for two minutes And later on how you can inject that. He only feed do unaffected flies.So basically take a double stick tape and usually these double stick tapes have two side, one side is a sticker than the other. So leave the sticker side up because that's the side that you're gonna glue that flies on top of it. And basically you'll, the gas apparatus, if you have this light on top of it, you'll be no gas flow and the flies you wake up if they're on top of the slide.So you can just direct the flow, making a little hood, a little chamber using tape. Then you direct the gas over the flies. So then you put your, this light here with the tape and you, you have your flies.Stick the flies on the top of the tape, all of them the same orientation, and put their head towards the direction that you'll be bringing the needle from. Because then you can just over the, the wing, you make sure since they fly, you don't want to fly. If they woke up during the procedure, don't want them to go away.So make sure that you take a little tip and pressure against the, the, the wings of the fly on the tail. And it's okay if they wings get damaged. And when you get rid, when you take the flies up the injection, most likely they'll lose their wings anyway, so they'll survive.That can, that can be useful because you're not gonna lose the flies that, so he puts a mineral oil on the micrometer for the back of the needle, then the needle injection polar, and then the, you can just adjust the pressure to bring the material up to the tip of the needle. I, I did to inject Somewhere between 10 to 15 nanoliters, but as, as long, as long as I'm consistents. Okay, so one important detail there is a cushion of this experiment is that the, the fly, once the apply is being injected when you saw the needle needs to come as far as you can through the upper part of the abdomen.Because here, basically when the needle penetrates, you have a, a, a, a, a bigger space to go up and down. And basically if it's close, closer to the cuticle, it's less likely they're gonna penetrate vital organs in the flight. You, it is very likely unlikely that you're gonna punch a hole in your gut, for instance, if that is gonna kill the flight that you are performing the surgery.My name is Voia Friedman. I am associate researcher at Princeton University in the lab of Eric Hels. And today I would like to tell you about this protocol that you just saw, the demonstration.So this protocol was done because we wanted to prove that UBA is capable of crossing tissues. So UBA is one of the most successful intracellular paras on the face of the earth. Infects from 20 to 70%of insects and many particles.And also is, is very important in terms of human diseases because also infect nematodes that transmit diseases like river blindness or s and ssis. And so, and CCA is our obligatory endo in, in these worms. And there's a great, a very big interest in using CCA also as means to control parasitic diseases transmitter by mosquito and Danny and this and by mosquito such as malaria and dengue.So this, this, this protocol basically was developed to show that once CCA inside the abdominal cavity of a fly, they can cross tissues and reach the germ. So basically what you can see here is the oph of the drosophila ovary and go back once present in the he leaf it, it can cross a pink peronial. She that is around the ovary.And then they cross a muscle layer. And actually one of the first places that they, they go inside the ovary is the somatic stem cell image. And then they can reach the germline.And this bacteria is transmitted the same way that mitochondria is transmitted is through, is through the mother transmitted, through the mother cyop plants present in the egg. So of course they needs to be present into the germline. So that's why this protocol is developed.And this protocol is, can be very useful to basically traditionally the way that people would try to trans infect wbaa basically take wbaa and introduce a new insect species. It would be through doing micro injections into the, into the embryo. But those are much more complicated.And basically here we are just taking advantage of the o biology since they are capable to cross tissues and moving into the stem cell tissues from, they can pass to the, through to the germline. So that's make easier taking UBA from certain, from, from certain species of insects that are infected and they can be transmitted to uninfected species. Prob I think it's the injection itself.Basically you need to be careful to mobilize the fly as well as you can. We are using double sided stick tape and you need to penetrate the needle basically as close to the, as parallel to the fly as possible so you don't go to deep. And it's less likely that we're gonna kill the fly after they're performing the surgery.

Microinjection of A. aegypti Embryos to Obtain Transgenic Mosquitoes

In this video, Nijole Jasinskiene demonstrates the methodology employed to generate transgenic Aedes aegypti mosquitoes, which are vectors for dengue fever.  The techniques for correctly preparing microinjection needles, dessicating embryos, and performing microinjection are demonstrated.

In this video, Nijole Jasinskiene demonstrates the methodology employed to generate transgenic Aedes aegypti mosquitoes, which are vectors for dengue fever. The techniques for correctly preparing microinjection needles, dessicating embryos, and performing microinjection are demonstrated.

In this video, Nijole Jasinskiene demonstrates the methodology employed to generate transgenic Aedes aegypti mosquitoes, which are vectors for dengue ...

Protocol for Dengue Infections in Mosquitoes (A. aegypti) and Infection Phenotype Determination

The purpose of this procedure is to infect the Aedes mosquito with dengue virus in a laboratory condition and examine the infection level and dynamic of the virus in the mosquito tissues. This protocol is routinely used for studying mosquito-virus interactions, especially for identification of novel host factors that are able to determine vector competence. The entire experiment must be conducted in a BSL2 laboratory. Similar to Plasmodium falciparum infections, proper attire including gloves and lab coat must be worn at all times. After the experiment, all the materials that came in contact with the virus need to be treated with 75% ethanol and bleached before proceeding with normal washing. All other materials need to be autoclaved before discarding them.

Protocol for Dengue Infections in Mosquitoes A. aegypti and Infection Phenotype Determination

Once a gene is identified as potentially refractory for the dengue virus, it must be evaluated for it's role in preventing viral infections within the mosquito. This protocol illustrates how the extent of dengue infections of mosquitoes can be assayed. The techniques for growing up the virus in culture, membrane feeding mosquitoes human blood, and assaying viral titers in the mosquito midgut are demonstrated.

The purpose of this procedure is to infect the Aedes mosquito with dengue virus in a laboratory condition and examine the infection level and dynamic of ...

Maintaining Wolbachia in Cell-free Medium

In this video protocol, procedures are demonstrated to (1) purify Wolbachia symbionts out of cultured mosquito cells, (2) use a fluorescent assay to ascertain the viability of the purified Wolbachia and (3) maintain the now extracellular Wolbachia in cell-free medium.  Purified Wolbachia remain alive in the extracellular phase but do not replicate until re-inoculated into eukaryotic cells.  Extracellular Wolbachia purified in this manner will remain viable for at least a week at room temperature, and possibly longer.  Purified Wolbachia are suitable for micro-injection, DNA extraction and other applications. 


This video describes a method for purifying Wolbachia pipientis from an Anopheles gambiae cell line and then culturing the endosymbiont in cell-free medium. An assay for viability of the bacterium is demonstrated.

In this video protocol, procedures are demonstrated to (1) purify Wolbachia symbionts out of cultured mosquito cells, (2) use a fluorescent assay to ...

Dissection of Midgut and Salivary Glands from Ae. aegypti Mosquitoes

The mosquito midgut and salivary glands are key entry and exit points for pathogens such as Plasmodium parasites and Dengue viruses. This video protocol demonstrates dissection techniques for removal of the midgut and salivary glands from Aedes aegypti mosquitoes. 



The mosquito midgut and salivary glands are key entry and exit points for vector pathogens like Plasmodium falciparum and the dengue virus. This video demonstrates the dissection techniques for removing the midgut and salivary glands from Aedes aegypti mosquitoes.

The mosquito midgut and salivary glands are key entry and exit points for pathogens such as Plasmodium parasites and Dengue viruses. This video protocol ...

Building a Better Mosquito: Identifying the Genes Enabling Malaria and Dengue Fever Resistance in A. gambiae and A. aegypti Mosquitoes

In this interview, George Dimopoulos focuses on the physiological mechanisms used by mosquitoes to combat Plasmodium falciparum and dengue virus infections.   Explanation is given for how key refractory genes, those genes conferring resistance to vector pathogens, are identified in the mosquito and how this knowledge can be used to generate transgenic mosquitoes that are unable to carry the malaria parasite or dengue virus.

In this interview, George Dimopoulos focuses on the physiological mechanisms used by mosquitoes to combat Plasmodium falciparum and dengue virus infections. Explanation is given for how key refractory genes, those genes conferring resistance to vector pathogens, are identified in the mosquito and how this knowledge can be used to generate transgenic mosquitoes that are unable to carry the malaria parasite or dengue virus.

In this interview, George Dimopoulos focuses on the physiological mechanisms used by mosquitoes to combat Plasmodium falciparum and dengue virus ...

RNAi-mediated Gene Knockdown and In Vivo Diuresis Assay in Adult Female Aedes aegypti Mosquitoes

This video protocol demonstrates an effective technique to knockdown a particular gene in an insect and conduct a novel bioassay to measure excretion rate. This method can be used to obtain a better understanding of the process of diuresis in insects and is especially useful in the study of diuresis in blood-feeding arthropods that are able to take up huge amounts of liquid in a single blood meal. 
This RNAi-mediated gene knockdown combined with an in vivo diuresis assay was developed by the Hansen lab to study the effects of RNAi-mediated knockdown of aquaporin genes on Aedes aegypti mosquito diuresis1. 
The protocol is setup in two parts: the first demonstration illustrates how to construct a simple mosquito injection device and how to prepare and inject dsRNA into the thorax of mosquitoes for RNAi-mediated gene knockdown. The second demonstration illustrates how to determine excretion rates in mosquitoes using an in vivo bioassay.

RNAi-mediated Gene Knockdown and In Vivo Diuresis Assay in Adult Female Aedes aegypti Mosquitoes

In this protocol we combine RNAi-mediated gene silencing with an in vivo diuresis assay to study the effects knockdown of genes of interest has on mosquito fluid excretion.

This video protocol demonstrates an effective technique to knockdown a particular gene in an insect and conduct a novel bioassay to measure excretion ...

Physiological Recordings and RNA Sequencing of the Gustatory Appendages of the Yellow-fever Mosquito Aedes aegypti

Electrophysiological recording of action potentials from sensory neurons of mosquitoes provides investigators a glimpse into the chemical perception of these disease vectors. We have recently identified a bitter sensing neuron in the labellum of female Aedes aegypti that responds to DEET and other repellents, as well as bitter quinine, through direct electrophysiological investigation. These gustatory receptor neuron responses prompted our sequencing of total mRNA from both male and female labella and tarsi samples to elucidate the putative chemoreception genes expressed in these contact chemoreception tissues. Samples of tarsi were divided into pro-, meso- and metathoracic subtypes for both sexes. We then validated our dataset by conducting qRT-PCR on the same tissue samples and used statistical methods to compare results between the two methods. Studies addressing molecular function may now target specific genes to determine those involved in repellent perception by mosquitoes. These receptor pathways may be used to screen novel repellents towards disruption of host-seeking behavior to curb the spread of harmful viruses.

Physiological Recordings and RNA Sequencing of the Gustatory Appendages of the Yellow-fever Mosquito Aedes aegypti

Using two methods to estimate gene expression in the major gustatory appendages of Aedes aegypti, we have identified the set of genes putatively underlying the neuronal responses to bitter and repulsive compounds, as determined by electrophysiological examination.

Electrophysiological recording of action potentials from sensory neurons of mosquitoes provides investigators a glimpse into the chemical perception of ...

Fat Body Organ Culture System in Aedes Aegypti, a Vector of Zika Virus

The insect fat body plays a central role in insect metabolism and nutrient storage, mirroring functions of the liver and fat tissue in vertebrates. Insect fat body tissue is usually distributed throughout the insect body. However, it is often concentrated in the abdomen and attached to the abdominal body wall.
The mosquito fat body is the sole source of yolk proteins, which are critical for egg production. Therefore, the in vitro culture of mosquito fat body tissues represents an important system for the study of mosquito physiology, metabolism, and, ultimately, egg production. The fat body culture process begins with the preparation of solutions and reagents, including amino acid stock solutions, Aedes physiological saline salt stock solution (APS), calcium stock solution, and fat body culture medium. The process continues with fat body dissection, followed by an experimental treatment. After treatment, a variety of different analyses can be performed, including RNA sequencing (RNA-Seq), qPCR, Western blots, proteomics, and metabolomics.
In our example experiment, we demonstrate the protocol through the excision and culture of fat bodies from the yellow fever mosquito, Aedes aegypti, a principal vector of arboviruses including dengue, chikungunya, and Zika. RNA from fat bodies cultured under a physiological condition known to upregulate yolk proteins versus the control were subject to RNA-Seq analysis to demonstrate the potential utility of this procedure for investigations of gene expression.

Fat Body Organ Culture System in Aedes Aegypti, a Vector of Zika Virus

The fat body is the central metabolic organ in insects. We present a live organ culture system that enables the user to study the responses of isolated fat body tissue to various stimuli.

The insect fat body plays a central role in insect metabolism and nutrient storage, mirroring functions of the liver and fat tissue in vertebrates. Insect ...

Improved Fecundity and Fertility Assay for Aedes aegypti using 24 Well Tissue Culture Plates (EAgaL Plates)

Mosquitoes represent a significant public health problem as vectors of various pathogens. For those studies that require an assessment of mosquito fitness parameters, in particular egg production and hatch rates at the individual level, conventional methods have put a substantial burden on investigators due to high labor intensity and laboratory space requirements. Described is a simple method using 24 well tissue culture plate with agarose in each well and digital imaging of each well to determine egg numbers and hatch rates at an individual level with substantially reduced time and space requirements.

Improved Fecundity and Fertility Assay for Aedes aegypti using 24 Well Tissue Culture Plates EAgaL Plates

Described is a time and space-saving method to count eggs and determine hatch rates of individual mosquitoes using 24 well tissue culture plates, which can substantially increase the scale and speed of fecundity and fertility assays.

Mosquitoes represent a significant public health problem as vectors of various pathogens. For those studies that require an assessment of mosquito fitness ...