We thank Texas A&#38;M Agrilife Research Insect Vectored Diseases Grant Program for funding. We also thank the Adelman lab members for help on developing this method and suggestions when drafting the manuscript, as well as Kevin Myles lab members. We also thank the reviewers and editors for their help to make this manuscript better.

1. Plate preparation
<ol>
	<li>Drill holes in 24 well tissue culture plate lids (4&#8722;6 holes per well) using a household drill with a ~1.6 mm (1/16 in) bit (Figure 2). 
	NOTE: These holes prevent water condensation from agarose to accumulate on the lid, where mosquitoes may lay eggs. The standard size of female Ae. aegypti (&#34;Liverpool&#34; strain) is ~3.11 mm wingspan. Reducing the size of the holes is recommended when using smaller mosquitoes to avoid escape from the plate.</li>
	<li>On the day prior to the oviposition experiment, wash and rinse the plates thoroughly and soak them in 1&#8722;5% bleach for 30&#8722;60 min at room temperature. Rinse thoroughly under running deionized water and dry them. 
	NOTE: This process reduces the chance of fungi and bacteria growing on agarose.</li>
	<li>Melt agarose at 2% in deionized water and immediately add 500 &#956;L of the molten agarose to each well of the 24 well plates using a 1,000 &#956;L pipette (Figure 3A,B). When the agarose begins to cool and clog the pipette tip, reheat the agarose and use a new pipette tip. 
	NOTE: Avoid touching the walls of the well with the pipette tip because it may leave a piece of agarose on the wall where a female may lay eggs, complicating the imaging and counting process.</li>
	<li>Before use, dry out any condensation on the well walls overnight on a lab bench (Figure 3C,D). 
	NOTE: The timeline of the EAgaL plate assay from blood feeding to larval imaging is depicted in Figure 4, which is for colony mosquitoes (Ae. aegypti &#8220;Liverpool&#8221;&#160;reared under 14 h light:10 h dark light cycle, 27 &#176;C, 80% relative humidity, 500 larvae in a 49.5 cm x 29.2 cm x 9.5 cm tray with 2 L of water), under insectary conditions. It is recommended that each laboratory test the EAgaL plate with the mosquitoes being used, especially when using different strains, different mosquito species, as well as different rearing protocols.</li>
</ol>
2. Mosquito feeding
NOTE: It is critical to use mosquitoes reared under uniform conditions for all treatment groups and control groups, because larval nutrition has an impact on mosquito fitness parameters10,11. Rear larvae under uncrowded conditions with sufficient food. Let female mosquitoes eclose in the presence of males so that mating is ensured, and mature for at least 3 days.
<ol>
	<li>Remove any source of water and/or sugar from the female mosquitoes at least 16 h prior to blood feeding in order to enhance blood feeding.</li>
	<li>Heat a water circulator for artificial feeding at 37 &#176;C and feed female mosquitoes using vertebrate blood placed in artificial feeders for 15&#8722;30 min (Figure 5A).</li>
	<li>Anesthetize mosquitoes with CO2 or on ice, transfer them to a glass dish on ice, and select ones that are engorged with blood (Figure 5B) into a container provided with 30% sucrose water for more than 72 h, when females finish excretion and egg development. 
	NOTE: If the females are provided with a lower concentration of sucrose water, remove the sucrose water and any wet surfaces after 48 h to prevent the females from ovipositing.</li>
</ol>
3. Oviposition
<ol>
	<li>About 1 h prior to transferring mosquitoes to the plates, add 2&#8722;3 drops (~80&#8722;120 &#956;L) of water into each plate well using a transfer pipette.</li>
	<li>At least 72 h after blood feeding, knockdown mosquitoes with CO2 or on ice, transfer them to a glass dish on ice, and individually place each mosquito on an inverted lid of the 24 well plate on ice (Figure 6A). 
	NOTE: This procedure has been applied from Ioshino et al.9.</li>
	<li>Once all 24 mosquitoes have been placed, cover the lid with an inverted plate bottom (Figure 6B), secure the lid and plate with a fresh, new rubber band and place in an environmental chamber (or rearing room) (Figure 6C) until mosquitoes recover (~10&#8722;15 min) and turn the plate right side up (Figure 6D).</li>
	<li>Allow the female mosquitoes to oviposit for 24&#8722;48 h and remove females by releasing them from the plates into a large cage. 
	NOTE: If it is important to keep track on individual females, anesthetize them by chilling the plates and remove them individually on ice. Delayed oviposition and dark excretions that interfere with egg counting were observed when females were transferred to the plates earlier than 72 h post blood meal (PBM) (Figure 7D).</li>
</ol>
4. Egg counting
<ol>
	<li>Check each well of the 24 well plate, as sometimes mosquitoes lay eggs on the wall of wells and at the margin of the agarose/plastic surface, where they are difficult to resolve in photographs. Using a wet paint brush, move the eggs laid on the wall and edge of agarose to the flat surface of the agarose so that all eggs are in a uniform plane and do not overlap with each other. 
	NOTE: The edge of the agarose is typically out of the camera&#8217;s focal plane due to the surface tension of the agarose solution.</li>
	<li>Using forceps, remove any broken legs, wings, and other particles in the wells that may interfere with imaging eggs (Figure 7C).</li>
	<li>Insert white paper underneath the plate to increase the contrast with dark mosquito eggs prior to imaging using a stereomicroscope illuminator (Figure 7A).</li>
	<li>Take an image of each well using a compact digital camera in microscope mode, which allows the user to focus on objects as close as 1 cm. This capability allows the camera to be placed directly on the plate to image eggs without a tripod or a stand (Figure 7A). To distinguish individual eggs laid in clumps, use the fine or super-fine mode to capture high-resolution images so that close-up details can be seen. 
	NOTE: Clear the memory card of the camera before use to prevent confusion between new and old images.</li>
	<li>After photographing each well of a plate, take an image of the entire plate with an imaging order label to distinguish each plate later (Figure 7E).</li>
	<li>Add a thin layer ~5 drops of water (~200 &#956;L) with a transfer pipette to each well to prevent its agarose plug and the eggs from drying and to induce embryo development and hatching. Pay attention to the water levels for the first few hours and check every day, because there may be water loss due to absorption by the agarose or evaporation. Add water when its level is too low for larvae to hatch. 
	NOTE: Evaporation of water occurs non-uniformly both within a plate and within a stack of plates. It has been observed that drying occurs in the following order: 1) corner wells (A1, A6, D1, D6) dry the fastest; followed by 2) wells at the outer edge of the plate (A2-A5, B1, B6, C1, C6, D2-D5); and last 3) wells inside. When plates are stacked, the top plate dries the fastest.</li>
	<li>Transfer the images to a computer with ImageJ (Fiji) and rename the files for easier organization such as &#8220;[plate ID]_[well ID].jpg&#8221; (Figure 8A,B).</li>
	<li>Create a spreadsheet file to record egg numbers (i.e., fecundity) and larval numbers and calculate hatch rate (i.e., fertility) (Figure 9E).</li>
	<li>Open the images with ImageJ (Fiji)12 and use the &#8220;multi-point&#8221; tool to mark each egg (Figure 9A&#8722;C); zoom-in or zoom-out using &#8220;+&#8221; or &#8220;&#8211;&#8221; key to count the eggs in&#160;clumps. After marking all the eggs, double-click the multi-point icon to bring up the number of marks (Figure 9D). Record the results in a spreadsheet.</li>
</ol>
5. Fertility assessment
NOTE: In 2 days, first instar larvae may begin to eclose in the wells. Wait for 3&#8722;5 more days before imaging/counting to ensure that all viable eggs hatch.
<ol>
	<li>Prepare larval food by mixing 1/16 tablespoon (~168 mg) of ground fish food (i.e., normal mosquito larval diet) in 20 mL of water and waiting for large solid particles to settle (Figure 10A&#8722;D). Start adding food (the supernatant) to the wells that contain hatched larvae as soon as they appear, because they do not survive for long without food. 
	NOTE: Excess addition of food particles may interfere with imaging and counting of hatched larvae.</li>
	<li>Approximately 5&#8211;8 days after addition of water to the wells, cool the plate by covering with crushed ice for 15&#8722;20 min to anesthetize larvae.</li>
	<li>Take images of each well while keeping the plates on ice as was done with the eggs. For larval images, provide a dark background by inserting a black material underneath the plate to help enhance the contrast. After photographing each well of a plate, take an image of the entire plate with an imaging order label to distinguish each plate later on. 
	NOTE: Images are taken while the plates are on ice because larval movement may compromise the counting. The plates are reusable; freeze and remove the mosquitoes and agarose to clean for the next use. The EAgaL plate is not suitable for rearing larvae to advanced stages.</li>
	<li>Open images with ImageJ (Fiji) and use the &#8220;multi-point&#8221; tool to count by clicking on each larva. Over the 3&#8722;5 days period some larvae may have molted, especially in wells with low numbers of larvae. Therefore, when counting, exclude the shed cuticles (i.e., exuviae), which look like head-only larvae with a little bit of body, or shrunken larvae (Figure 10E). Record the results in the spreadsheet.</li>
</ol>
6. Perform analysis
<ol>
	<li>Having collected all necessary data for the fecundity and fertility analysis, perform appropriate statistical analysis and create graphs using preferred software.</li>
</ol>

<ol>
	<li>Bond, J. G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optimization+of+irradiation+dose+to+Aedes+aegypti+and+Ae.+albopictus+in+a+sterile+insect+technique+program.">Optimization of irradiation dose to Aedes aegypti and Ae. albopictus in a sterile insect technique program.</a> PLoS One. 14 (2), 0212520 (2019).</li><li>Fernandes, K. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Aedes+aegypti+larvae+treated+with+spinosad+produce+adults+with+damaged+midgut+and+reduced+fecundity.">Aedes aegypti larvae treated with spinosad produce adults with damaged midgut and reduced fecundity.</a> Chemosphere. 221, 464-470 (2019).</li><li>Inocente, E. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Insecticidal+and+Antifeedant+Activities+of+Malagasy+Medicinal+Plant+(Cinnamosma+sp.)+Extracts+and+Drimane-Type+Sesquiterpenes+against+Aedes+aegypti+Mosquitoes.">Insecticidal and Antifeedant Activities of Malagasy Medicinal Plant (Cinnamosma sp.) Extracts and Drimane-Type Sesquiterpenes against Aedes aegypti Mosquitoes.</a> Insects. 10 (11), 373 (2019).</li><li>Marrelli, M. T., Moreira, C. K., Kelly, D., Alphey, L., Jacobs-Lorena, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mosquito+transgenesis:+what+is+the+fitness+cost.">Mosquito transgenesis: what is the fitness cost.</a> Trends in Parasitology. 22 (5), 197-202 (2006).</li><li>da Silva Costa, G., Rodrigues, M. M. S., Silva, A. A. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Toward+a+blood-free+diet+for+Anopheles+darlingi+(Diptera:+Culicidae).">Toward a blood-free diet for Anopheles darlingi (Diptera: Culicidae).</a> Journal of Medical Entomology. , 217 (2019).</li><li>Gonzales, K. K., Tsujimoto, H., Hansen, I. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Blood+serum+and+BSA,+but+neither+red+blood+cells+nor+hemoglobin+can+support+vitellogenesis+and+egg+production+in+the+dengue+vector+Aedes+aegypti.">Blood serum and BSA, but neither red blood cells nor hemoglobin can support vitellogenesis and egg production in the dengue vector Aedes aegypti.</a> PeerJ. 3, 938 (2015).</li><li>Gonzales, K. K., et al. <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+SkitoSnack,+an+Artificial+Blood+Meal+Replacement,+on+Aedes+aegypti+Life+History+Traits+and+Gut+Microbiota.">The Effect of SkitoSnack, an Artificial Blood Meal Replacement, on Aedes aegypti Life History Traits and Gut Microbiota.</a> Scientific Reports. 8 (1), 11023 (2018).</li><li>Tsujimoto, H., Anderson, M. A. E., Myles, K. M., Adelman, Z. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+of+Candidate+Iron+Transporters+From+the+ZIP/ZnT+Gene+Families+in+the+Mosquito+Aedes+aegypti.">Identification of Candidate Iron Transporters From the ZIP/ZnT Gene Families in the Mosquito Aedes aegypti.</a> Frontiers in Physiology. 9, 380 (2018).</li><li>Ioshino, R. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Oviplate:+A+Convenient+and+Space-Saving+Method+to+Perform+Individual+Oviposition+Assays+in+Aedes+aegypti.">Oviplate: A Convenient and Space-Saving Method to Perform Individual Oviposition Assays in Aedes aegypti.</a> Insects. 9 (3), 103 (2018).</li><li>Price, D. P., Schilkey, F. D., Ulanov, A., Hansen, I. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Small+mosquitoes,+large+implications:+crowding+and+starvation+affects+gene+expression+and+nutrient+accumulation+in+Aedes+aegypti.">Small mosquitoes, large implications: crowding and starvation affects gene expression and nutrient accumulation in Aedes aegypti.</a> Parasites &amp; Vectors. 8, 252 (2015).</li><li>Valerio, L., Matilda Collins, C., Lees, R. S., Benedict, M. Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Benchmarking+vector+arthropod+culture:+an+example+using+the+African+malaria+mosquito,+Anopheles+gambiae+(Diptera:+Culicidae).">Benchmarking vector arthropod culture: an example using the African malaria mosquito, Anopheles gambiae (Diptera: Culicidae).</a> Malaria Journal. 15 (1), 262 (2016).</li><li>Rueden, C. T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ImageJ2:+ImageJ+for+the+next+generation+of+scientific+image+data.">ImageJ2: ImageJ for the next generation of scientific image data.</a> BMC Bioinformatics. 18 (1), 529 (2017).</li></ol>

The authors declare no conflicts of interest for this study.

The EAgaL plate drastically reduces labor, time, and space to conduct individual fecundity and fertility assays in Aedes aegypti when compared to the FT method. Preliminary comparison between the FT method and the EAgaL plate resulted in shorter times for all steps (imaging technique was applied to the FT method) (Table 1). As a reference, an estimate of startup and&#160;per assay (one 24 well EAgaL plate versus 24 FTs) costs are provided in Table 2.
<p class="jove_...

The control of mosquitoes to protect humans from vector-borne pathogens is an important public health goal, mainly due to the lack of effective vaccines for most of the pathogens carried by mosquitoes. Many studies aim to reduce mosquito fitness in conjunction with a field-applicable population reduction strategy1,2,3. This includes extensive studies to create transgenic mosquitoes and/or CRISPR/Cas9 knockout lines. Such population modification approaches require a detailed assessment of individual fitness parameters4. Conventional laboratory techniques to assess the fitness of female mosquitoes includes the individual containment of mated, blood-fed female mosquitoes in 100 mL containers5, modified 50 mL conical tubes, or tubes for Drosophila rearing modified by providing moist surfaces using damp cotton and filter paper discs for oviposition (i.e., egg papers)1,2,6,7. Such methods require a relatively large space (e.g., 30 cm x 30 cm x 10 cm: W x L x H for up to 100 Drosophila tubes) (Figure 1), and the manipulation of individual egg papers for counting eggs and hatching larvae, which can be labor intensive. This manuscript presents a method to count mosquito eggs and determine hatch rates using 24 well plates and agarose as an oviposition surface to circumvent these issues8.
Concurrently, Ioshino et al.9 described a detailed method using 12 and 24 well plates to perform egg counting obtained from individual females. Their protocol represented a significant improvement from conventional methods in saving time and space9. However, the protocol they described continues to use wet filter paper as a surface for oviposition, which requires unfolding each individual paper to get counts, as eggs are often found underneath or in folds. Their protocol also did not include the use of imaging technologies or a method for larval counting.
Presented is an improved method to perform fitness assays for egg number (i.e., fecundity) and hatch rate (i.e., fertility) using agarose as an oviposition surface in a 24 well tissue culture plate format for Ae. aegypti that oviposit on moist surfaces. These plates were named &#8220;EAgaL&#8221; plates, from Egg, Agarose, and Larva. These 24 well plates provide individual mosquitoes with a minimal surface to lay eggs, thus simplifying and drastically reducing the time and effort needed to count and maintain eggs and hatched larvae for a few days. The EAgaL plate uses translucent agarose for the oviposition surface, which eliminates the need for handling egg papers and finding the eggs and larvae when hatched; photographing each well establishes a long-term archived record of the results and separates the counting process in both time and space from the rearing/handling process, where time is often limited.

<table><tbody><tr><td>1.6 mm &amp;Phi; drill bit</td><td></td><td></td><td>alternatively heated nails can be used</td></tr><tr><td>1000 &amp;mu;L pipette tips (long)</td><td>Olympus plastics</td><td>24-165RL</td><td></td></tr><tr><td>24-well tissue culture plate</td><td>Thermo Scientific</td><td>930186</td><td>clear, flat-bottom with ringed lid plates</td></tr><tr><td>Agarose</td><td>VWR</td><td>0710-500G</td><td></td></tr><tr><td>Compact digital camera</td><td>Olympus</td><td>TG-5/TG-6</td><td></td></tr><tr><td>Computer (Windows, Mac or Linux)</td><td></td><td></td><td></td></tr><tr><td>Deionized water</td><td></td><td></td><td></td></tr><tr><td>Fiji (imageJ) software</td><td></td><td></td><td>download from: https://fiji.sc/</td></tr><tr><td>Forceps</td><td>Dumont</td><td></td><td>sharp forceps may break mosquito&#039;s body</td></tr><tr><td>Glass Petri dishes</td><td>VWR</td><td></td><td></td></tr><tr><td>Household bleach</td><td></td><td></td><td></td></tr><tr><td>Household electric drill</td><td></td><td></td><td></td></tr><tr><td>illuminator for stereomicroscope (gooseneck)</td><td></td><td></td><td></td></tr><tr><td>P-1000 pipette</td><td>Gilson</td><td></td><td></td></tr><tr><td>paint brushes</td><td></td><td></td><td></td></tr><tr><td>Rubber bands</td><td></td><td></td><td></td></tr><tr><td>SD card</td><td></td><td></td><td>to record digital camera images (DSHC, SDXC should be better)</td></tr><tr><td>Spreadsheet software (Microsoft Excel)</td><td>Microsoft</td><td></td><td>Any spreadsheet software works</td></tr><tr><td>TetraMin fish food</td><td>Tetra</td><td></td><td>ground with coffee grinder, blender or morter &amp;amp; pestle</td></tr><tr><td>Transfer pipetts</td><td>VWR</td><td>16011-188</td><td></td></tr></tbody></table>

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.

Mosquitoes were injected with dsRNA targeting a candidate iron transporter (FeT) or control gene (EGFP), blood-fed, and measured for fecundity and fertility output using the EAgaL plate method, following the procedure described above.
Mosquitoes in which FeT expression was silenced following dsRNA injection exhibited a significant reduction in both egg number and hatch rate (Figure 11A&#8722;C). All control and treatment mosquitoes were placed in ...

Watch this Scientific Journal Video about Improved Fecundity and Fertility Assay for Aedes aegypti using 24 Well Tissue Culture Plates EAgaL Plates at JoVE.com

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.

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 ...

improved-fecundity-fertility-assay-for-aedes-aegypti-using-24-well

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Biology

5.0K Views.  Texas A&M University. 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.

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

Mass Production of Genetically Modified Aedes aegypti for Field Releases in Brazil

New techniques and methods are being sought to try to win the battle against mosquitoes. Recent advances in molecular techniques have led to the development of new and innovative methods of mosquito control based around the Sterile Insect Technique (SIT)1-3. A control method known as RIDL&#160;(Release of Insects carrying a Dominant Lethal)4, is based around SIT, but uses genetic methods to remove the need for radiation-sterilization5-8. A RIDL&#160;strain of Ae. aegypti was successfully tested in the field in Grand Cayman9,10; further field use is planned or in progress in other countries around the world.
Mass rearing of insects has been established in several insect species and to levels of billions a week. However, in mosquitoes, rearing has generally been performed on a much smaller scale, with most large scale rearing being performed in the 1970s and 80s. For a RIDL&#160;program it is desirable to release as few females as possible as they bite and transmit disease. In a mass rearing program&#160;there are several stages to produce the males to be released: egg production, rearing eggs until pupation, and then sorting males from females before release. These males are then used for a RIDL&#160;control program,&#160;released as either pupae or adults11,12.
To suppress a mosquito population using RIDL&#160;a large number of high quality male adults need to be reared13,14. The following describes the methods for the mass rearing of OX513A, a RIDL&#160;strain of Ae. aegypti 8, for release and covers the techniques required for the production of eggs and mass rearing RIDL&#160;males for a control program.

Mass Production of Genetically Modified Aedes aegypti for Field Releases in Brazil

To achieve population suppression of Aedes aegypti using the RIDL&#174; (Release of Insects carrying a Dominant Lethal) system, large numbers of male mosquitoes need to be released. This requires the use of mass rearing techniques and technology to provide reliable systems to obtain the maximum number of high quality male mosquitoes.

New techniques and methods are being sought to try to win the battle against mosquitoes. Recent advances in molecular techniques have led to the ...

Environment

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 ...

A Calcium Bioluminescence Assay for Functional Analysis of Mosquito (Aedes aegypti) and Tick (Rhipicephalus microplus) G Protein-coupled Receptors

Arthropod hormone receptors are potential targets for novel pesticides as they regulate many essential physiological and behavioral processes. The majority of them belong to the superfamily of G protein-coupled receptors (GPCRs). We have focused on characterizing arthropod kinin receptors from the tick and mosquito. Arthropod kinins are multifunctional neuropeptides with myotropic, diuretic, and neurotransmitter function. Here, a method for systematic analyses of structure-activity relationships of insect kinins on two heterologous kinin receptor-expressing systems is described. We provide important information relevant to the development of biostable kinin analogs with the potential to disrupt the diuretic, myotropic, and/or digestive processes in ticks and mosquitoes.
The kinin receptors from the southern cattle tick, Boophilus microplus (Canestrini), and the mosquito Aedes aegypti (Linnaeus), were stably expressed in the mammalian cell line CHO-K1. Functional analyses of these receptors were completed using a calcium bioluminescence plate assay that measures intracellular bioluminescence to determine cytoplasmic calcium levels upon peptide application to these recombinant cells. This method takes advantage of the aequorin protein, a photoprotein isolated from luminescent jellyfish. We transiently transfected the aequorin plasmid (mtAEQ/pcDNA1) in cell lines that stably expressed the kinin receptors. These cells were then treated with the cofactor coelenterazine, which complexes with intracellular aequorin. This bond breaks in the presence of calcium, emitting luminescence levels indicative of the calcium concentration. As the kinin receptor signals through the release of intracellular calcium, the intensity of the signal is related to the potency of the peptide.
This protocol is a synthesis of several previously described protocols with modifications; it presents step-by-step instructions for the stable expression of GPCRs in a mammalian cell line through functional plate assays (Staubly et al., 2002 and Stables et al., 1997). Using this methodology, we were able to establish stable cell lines expressing the mosquito and the tick kinin receptors, compare the potency of three mosquito kinins, identify critical amino acid positions for the ligand-receptor interaction, and perform semi-throughput screening of a peptide library. Because insect kinins are susceptible to fast enzymatic degradation by endogenous peptidases, they are severely limited in use as tools for pest control or endocrinological studies. Therefore, we also tested kinin analogs containing amino isobutyric acid (Aib) to enhance their potency and biostability. This peptidase-resistant analog represents an important lead in the development of biostable insect kinin analogs and may aid in the development of neuropeptide-based arthropod control strategies.

A Calcium Bioluminescence Assay for Functional Analysis of Mosquito Aedes aegypti and Tick Rhipicephalus microplus G Protein-coupled Receptors

This protocol provides instructions for clonal-cell line selection and a calcium bioluminescence assay to analyze the structure-activity relationships of synthesized arthropod neuropeptides on their cognate GPCRs. This assay can be used for receptor deorphanization and structure-activity relationship studies for synthetic analog design and peptide/drug-lead discovery.

Arthropod hormone receptors are potential targets for novel pesticides as they regulate many essential physiological and behavioral processes.  The ...

Immunology and Infection

Propagation of the Microsporidian Parasite Edhazardia aedis in Aedes aegypti Mosquitoes

Edhazardia aedis is a microsporidian parasite of Aedes aegypti mosquitoes, a disease vector that transmits multiple arboviruses which cause millions of disease cases each year. E. aedis causes mortality and reduced reproductive fitness in the mosquito vector and has been explored for its potential as a biocontrol agent. The protocol we present for culturing E. aedis is based on its natural infection cycle, which involves both horizontal and vertical transmission at different life stages of the mosquito host. Ae. aegypti mosquitoes are exposed to spores in the larval stage. These infected larvae then mature into adults and transmit the parasite vertically to their offspring. Infected offspring are then used as a source of spores for future horizontal transmission. Culturing E. aedis can be challenging to the uninitiated given the complexities of the parasite&#8217;s life cycle, and this protocol provides detailed guidance and visual aids for clarification.

Propagation of the Microsporidian Parasite Edhazardia aedis in Aedes aegypti Mosquitoes

A protocol to culture the microsporidian parasite Edhazardia aedis. The parasite is passaged from one generation of Aedes aegypti mosquitoes to the next via horizontal transfer at the larval stage followed by vertical transmission at the adult stage. Live sporoplasms survive long-term in infected eggs.

Edhazardia aedis is a microsporidian parasite of Aedes aegypti mosquitoes, a disease vector that transmits multiple arboviruses which cause millions 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 ...

Maintaining Aedes aegypti Mosquitoes Infected with Wolbachia

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

Maintaining Aedes aegypti Mosquitoes Infected with Wolbachia

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

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

Preparing Irradiated and Marked Male Aedes aegypti Mosquitoes for Release in an Operational Sterile Insect Technique Program

The control of such human diseases as dengue, Zika, and chikungunya relies on the control of their vector, the Aedes aegypti mosquito, because there is no prevention. Control of mosquito vectors can rely on chemicals applied to the immature and adult stages, which can contribute to the mortality of non-targets and more importantly, lead to insecticide resistance in the vector. The sterile insect technique (SIT) is a method of controlling populations of pests through the release of sterilized adult males that mate with wild females to produce non-viable offspring. This paper describes the process of producing sterile males for use in an operational SIT program for the control of Aedes aegypti mosquitoes. Outlined here are the steps used in the program including rearing and maintaining a colony, separating male and female pupae, irradiating and marking adult males, and shipping Aedes aegypti males to the release site. Also discussed are procedural caveats, program limitations, and future objectives.

Preparing Irradiated and Marked Male Aedes aegypti Mosquitoes for Release in an Operational Sterile Insect Technique Program

The sterile insect technique (SIT) is used to control specific, medically important mosquito populations that may be resistant to chemical controls. Here, we describe a method of mass rearing and preparation of sterile male mosquitoes for release in an operational SIT program targeting the Aedes aegypti mosquito.

The control of such human diseases as dengue, Zika, and chikungunya relies on the control of their vector, the Aedes aegypti mosquito, because there is no ...

Using the Fluorescent Dye, Rhodamine B, to Study Mating Competitiveness in Male Aedes aegypti Mosquitoes

The success of sterile or incompatible insect technique-based population suppression programs depends on the ability of released males to compete for wild-type females and induce sterility in the target population. Hence, laboratory assessment of male mating competitiveness is essential for evaluating the release strain&#8217;s fitness before field release. Conventionally, such an assay is performed by determining the proportion of viable eggs produced by the females after being simultaneously exposed to two sets of males (wild-type and release strains) for copulation. However, this process is time-consuming and laborious due to the need to first blood-feed the females for egg production and then hatch and enumerate the hatched eggs to determine egg viability.&#160;
Moreover, this method cannot discern the degree of competitiveness between two sterile or Wolbachia-infected mosquito lines as wild-type female mosquitoes will only produce non-viable eggs upon mating with both. To circumvent these limitations, this paper describes a more direct method of measuring male mosquito mating competitiveness in laboratory settings using the fluorescent dye, rhodamine B (RhB), which can be used to mark males by feeding them in sucrose solution containing RhB. After the mating assay, the presence of fluorescing sperms in the spermathecae of a female can be used to determine her mating partner. This method is cost-effective, reduces the experimental time by 90% and allows comparison of mating fitness between two sterile or Wolbachia-infected lines.&#160;

Using the Fluorescent Dye, Rhodamine B, to Study Mating Competitiveness in Male Aedes aegypti Mosquitoes

Here, we present a protocol for studying mating competitiveness of male Aedes aegypti using fluorescent dye as a marker. Female mosquitoes are exposed to both marked and unmarked males for copulation. Post mating, their spermathecae are examined under a fluorescence microscope to determine their mating partner.

The success of sterile or incompatible insect technique-based population suppression programs depends on the ability of released males to compete for ...

High-Throughput Screening of Microbial Isolates with Impact on Caenorhabditis elegans Health

With its small size, short lifespan, and easy genetics, Caenorhabditis elegans offers a convenient platform to study the impact of microbial isolates on host physiology. It also fluoresces in blue when dying, providing a convenient means of pinpointing death. This property has been exploited to develop high-throughput label-free C. elegans survival assays (LFASS). These involve time-lapse fluorescence recording of worm populations set in multiwell plates, from which population median time of death can be derived. The present study adopts the LFASS approach to screen multiple microbial isolates at once for the effects on C. elegans susceptibility to severe heat and oxidative stresses. Such microbial screening pipeline, which can notably be used to prescreen probiotics, using severe stress resistance as a proxy for host health is reported here. The protocol describes how to grow both C. elegans gut microbiota isolate collections and synchronous worm populations in multiwell arrays before combining them for the assays. The example provided covers the testing of 47 bacterial isolates and one control strain on two worm strains, in two stress assays in parallel. However, the approach pipeline is readily scalable and applicable to the screening of many other modalities. Thus, it provides a versatile setup to rapidly survey a multiparametric landscape of biological and biochemical conditions that impact C. elegans health.

High-Throughput Screening of Microbial Isolates with Impact on Caenorhabditis elegans Health

Gut microbes may positively or negatively impact the health of their host via specific or conserved mechanisms. Caenorhabditis elegans is a convenient platform to screen for such microbes. The present protocol describes high-throughput screening of 48 bacterial isolates for impact on nematode stress resistance, used as a proxy for worm health.

With its small size, short lifespan, and easy genetics, Caenorhabditis elegans offers a convenient platform to study the impact of microbial isolates on ...

Quantifying Fitness Costs in Transgenic Aedes aegypti Mosquitoes

Transgenic mosquitoes often display fitness costs compared to their wild-type counterparts. In this regard, fitness cost studies involve collecting life parameter data from genetically modified mosquitoes and comparing them to mosquitoes lacking transgenes from the same genetic background. This manuscript illustrates how to measure common life history traits in the mosquito Aedes aegypti, including fecundity, wing size and shape, fertility, sex ratio, viability, development times, male contribution, and adult longevity. These parameters were chosen because they reflect reproductive success, are simple to measure, and are commonly reported in the literature. The representative results quantify fitness costs associated with either a gene knock-out or a single insertion of a gene drive element. Standardizing how life parameter data are collected is important because such data may be used to compare the health of transgenic mosquitoes generated across studies or to model the transgene fixation rate in a simulated wild-type mosquito population. Although this protocol is specific for transgenic Aedes aegypti, the protocol may also be used for other mosquito species or other experimental treatment conditions, with the caveat that certain biological contexts may require special adaptations.

Quantifying Fitness Costs in Transgenic Aedes aegypti Mosquitoes

The present protocol describes how to measure common life parameter data in Aedes aegypti mosquitoes, including fecundity, wing size, fertility, sex ratio, viability, development times, male contribution, and adult longevity. These measurements can be used to assess the fitness of transgenic mosquitoes.

Transgenic mosquitoes often display fitness costs compared to their wild-type counterparts. In this regard, fitness cost studies involve collecting life ...

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

Summary

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Chapters in this video

Mass Production of Genetically Modified Aedes aegypti for Field Releases in Brazil

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

A Calcium Bioluminescence Assay for Functional Analysis of Mosquito (Aedes aegypti) and Tick (Rhipicephalus microplus) G Protein-coupled Receptors

Propagation of the Microsporidian Parasite Edhazardia aedis in Aedes aegypti Mosquitoes

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

Maintaining Aedes aegypti Mosquitoes Infected with Wolbachia

Preparing Irradiated and Marked Male Aedes aegypti Mosquitoes for Release in an Operational Sterile Insect Technique Program

Using the Fluorescent Dye, Rhodamine B, to Study Mating Competitiveness in Male Aedes aegypti Mosquitoes

High-Throughput Screening of Microbial Isolates with Impact on Caenorhabditis elegans Health

Quantifying Fitness Costs in Transgenic Aedes aegypti Mosquitoes