The authors would like to thank Drs. Bill Reid and Alexander Franz from the University of Missouri for their support with this protocol. The authors would also like to thank Dr. Benjamin Krajacich from the NIH/NIAID for his support with the R analysis. This study was funded by the NIH, grant number R01 AI130085 (KEO), and the NIH/NIAID Division of Intramural Research Program AI001246 (EC).

NOTE: This protocol is written for transgenic and wild-type Ae. aegypti lines that have been previously validated and established. For more information on generating transgenic Ae. aegypti, see&#160;Kistler et al.22&#160;and Coates&#160;et al.23.&#160;All experiments outlined below were performed under standard Ae.&#160;aegypti rearing procedures. Mosquitoes were maintained at 28 &#176;C with 75%-80% relative humidity and a 12 h light/12 h dark cycle. A minimum of 100 individual egg papers per mosquito stain is highly recommended for statistical purposes. Comprehensive fitness studies take approximately 3 months to complete (Figure 1).
1. Measuring fecundity in female mosquitoes
<ol>
	<li>Hatch transgenic and WT mosquitoes (of the same genetic background but lacking the transgene) by placing egg papers into sterile water overnight in the insectary, following standard rearing conditions. Allow the larvae to feed ad libitum by providing several drops (each about 50 &#181;L) of ground fish food (Tetramin) slurry (roughly 30% w/v). The first instar larvae are recognizable 1 day post-hatching. Place larvae into larger containers so there is no crowding, generally 100-150 larvae can be reared in a 1 L volume of water with a surface area of ca. 250 cm2. Continue to provide Tetramin slurry as a food source, allowing the larvae to feed ad&#160;libitum. Add the food slurry in discrete droplets so&#160;that sufficient food is always available to the larvae, and so that the water does not appear cloudy. The larvae will increase in size with each instar and will ultimately enter pupation in the larval pans.</li>
	<li>Once the pupae begin emerging, ~5-6 days post-hatching, separate them by sex to ensure the adults are virgins once they emerge (for subsequent mating), as described below.
	<ol>
		<li>To sort the pupae by sex, begin by picking the pupae using 3 mL plastic pipettors into small plastic cups containing clean tap water. Male pupae tend to be smaller and tend to emerge first compared to females. First, sort the pupae by size, and then confirm this initial screen by morphology.</li>
		<li>Move the pupa to a glass microscope slide and remove excess water with a tissue. This immobilizes the pupa. Under a stereoscope at 2x-4x magnification, position the pupa ventral side up using a finely tipped paintbrush.</li>
		<li>Visually analyze the tail for differences in genital lobe shape (at the end of the pupal abdominal segments behind the paddles; Figure 2). Females have pointed fins that extend past the genital lobes, roughly resembling a crown or vampire teeth, whereas male genital lobes are rounded and do not have these pointed structures. See Carvalho et al.24 for additional figures.</li>
	</ol>
	</li>
	<li>Place male and female pupae in separate water cups and place them in proper containment, for example, 1.9 L cartons (1/2 gallon or 64 oz) with 0.2 m x 0.2 m (8 in x 8 in) white organdy fabric, each containing ~150 pupae.</li>
	<li>Provide a sugar and water source on top of the cages (e.g., raisins and a water-filled cup covered with gauze, secured with two rubber bands). Cut a pen-sized hole into the cartons and cover it with a rubber stopper so that additional pupae can be added into the cages as they emerge.</li>
	<li>At 3-5 days post-adult emergence, anesthetize the mosquitoes by placing the cartons at 4 &#176;C for ~2-5 min. Although they may still twitch, the mosquitoes are sufficiently anesthetized once the majority fall to the bottom of the carton. Place the mosquitoes on a glass Petri dish on ice for ~30 min-1 h. 
	NOTE: Leaving the mosquitoes at 4 &#176;C for too long may significantly impact their health, although we have not found any impacts if they are left on ice for up to 1 h.</li>
	<li>Cross WT or transgenic GD1 or GD29&#160;(CRISPR/Cas-9-based-gene drive) males en masse with virgin WT (control) females by adding anesthetized mosquitoes in ratios of five WT or transgenic males to one WT female into mosquito cartons, each containing ~150 mosquitoes. Be sure to label the cages by male type. 
	NOTE: This crossing scheme is to determine the fitness for GD1 or GD29 hemizygotes (i.e.,&#160;mosquitoes harboring one copy of the transgene). To determine the fitness for homozygous transgenic lines, the crossing scheme can be adapted so that transgenic males and females are allowed to mate in comparison with crosses of WT males and females.</li>
	<li>After 2-3 days, blood feed the females following standard rearing practices25 by providing a blood source on top of the cage. Remove the sugar source 24 h pre-blood feeding and the water source ~2-3 h pre-blood feeding to increase feeding efficiency.</li>
	<li>After blood feeding, anesthetize the mosquitoes by placing them at 4 &#176;C, as in step 1.5.</li>
	<li>Sort the blood fed (fully engorged) females from unfed females by visually screening their abdomens for engorgement. Discard the non-fed, partially fed, and males.</li>
	<li>Place the engorged females back into their respective cages and provide a sugar and water source on top of the cages.</li>
	<li>After 2-3 days, place individual females in 50 mL conical tubes lined with filter paper that has been pre-labeled in pencil (Figure 3). Cover the conical tubes with mesh and seal with two rubber bands.</li>
	<li>Once females are awake, fill all the tubes with ~5 mL of tap water. Place a small piece of raisin or sugar-soaked cotton ball on top of each conical tube. Leave the females for 1-2 days in the insectary and allow them to oviposit.</li>
	<li>After 1-2 days, drain each conical tube and anesthetize the mosquitoes by placing them at 4 &#176;C. For quick processing, use the tip of a 3 mL plastic pipettor pressed against the mesh to drain the water from each conical tube. Rotate the tubes upside down and place them on top of a paper towel to drain. Place the 50 mL conical tubes in a tube holder and place the rack at 4 &#176;C.</li>
	<li>Collect the egg paper and count the number of eggs on each paper. Count the females that do not lay eggs as 0 and include them in the analysis. Calculate the average fecundity as the total number of eggs counted on each paper divided by total number of females screened. 
	NOTE: Digital pictures of the individual egg papers may serve as a visual record for counting confirmation (Figure 4).</li>
	<li>Dry the egg papers and place them back into the insectary. It is recommended that the egg papers dry for at least 3-5 days for optimum hatching25.</li>
</ol>
2. Measuring wing length, area, and centroid size
<ol>
	<li>Freeze at least 25 WT or transgenic age-matched blood fed females from the completed fecundity study by placing the mosquito cages at -20 &#176;C for approximately 15 min. 
	NOTE: Do not freeze for a longer period of time and do not shake or drop the container, as this may cause the wings to fall off the mosquitoes.</li>
	<li>Dissect the left wing of each mosquito, excising at the joint of the wing and the thorax, removing the entirety of the wing, following the below protocol.
	<ol>
		<li>In one hand, hold the mosquito with forceps, grasping the thorax region beneath the wings.</li>
		<li>Using another pair of forceps in the other hand, dissect the left wing of each mosquito, excising at the joint of the wing and the thorax using gentle pressure to pull the wing off, removing the entirety of the wing without ripping the wing veins.</li>
	</ol>
	</li>
	<li>Using forceps, place the wing on a glass slide, lying flat. Using a transfer pipette, add one drop from a 15 mL stock solution, comprised&#160;of 70% ethanol, and one drop of dish soap to the wing on the slide.</li>
	<li>Add one drop of 80% glycerol to the coverslip and place on top of the wing in the ethanol-soap solution.</li>
	<li>Photograph the wings mounted on the slides using a camera with a 65 mm lens (7D-65 mm-1x; zoom = 1, 200, 6.3, ISO = 100). Note the pixel/mm scale and ensure this remains the same for all images. If multiple wings are imaged together, crop the photos to individual wings and add scale bars to each photo. Save the images in TIFF format with LZW compression, with each image clearly labelled with all information for the individual wing. 
	NOTE: The wings may be photographed using a different camera and magnification or with a camera attached to a microscope for other studies, if the settings remain the same and the pixel/mm scale remains the same for each photographed wing in the study. The pixel/mm ratio will be accounted for when determining measurements.</li>
	<li>Download tpsUtil and tpsDig, which are free, morphometric image-processing softwares. See Rohlf26,27 for more detailed information about the software.</li>
	<li>Digitize the wings using tpsUtil to create a TPS file of the wing photographs for the next step of digitization.
	<ol>
		<li>To do this, open tpsUtil, and under operation, select Build tps file from images. Click input, select the file containing the individual wing images, and then select any image to choose this folder. Click output and name the file that will be created in .tps format. Click save.</li>
		<li>Then, under actions, click setup. Select include all, click create and close the pop-up box. Under operation, now select Randomly order specimens. Click input and select the .tps file that was just created. Click output, rename the .tps file to indicate that this is the file version with the samples randomized, and click save. Then, click Create. Close the tpsUtil program. 
		NOTE: To reduce measurement error, the wing photos may be duplicated and utilized twice for a technical replicate.</li>
	</ol>
	</li>
	<li>Use the tpsDig software to identify two-dimensional Cartesian coordinates for the landmarks. Under input, select the .tps file created in step 2.7. Set the scale to the scale determined by step 2.5; under options, select image tools, and then click measure.</li>
	<li>Add landmarks to the image by clicking on the blue crosshair icon representing digitize landmarks, and then click on the wing joints indicated in Figure 5, in order from Landmark 1 to Landmark 14. 
	NOTE: These landmarks were chosen because they have previously been used in wing landmark digitization and analysis19, are easy to locate at the joints of wing veins across many samples, and provide enough wing landmarks to capture the size and shape of the wing.</li>
	<li>When marking landmarks, mark any missing or obscured landmarks (due to wing folding or damage) as missing in tpsDig by clicking the M icon to exclude them from analysis and retain the correct order of the landmarks. Click the red, right-facing arrow to repeat the landmark digitization for the next images until all images in the file have been completed. Save and close the file. 
	NOTE: The landmarks must be added to the wing image in the correct order for each image for the calculations of wing length, area, and size to be correct.</li>
	<li>Calculate the wing length and area using the R script provided in Supplementary File 1, modifying for the .tps file location and image scale as per the experiment.</li>
	<li>Calculate the wing centroid size, a measure of size statistically independent from shape, defined as the square root of the sum of squared distances of each of the landmarks from the center point of the wing28,29.</li>
	<li>Use a generalized procrustes alignment to remove non-shape variation and plot mean landmark coordinates, using wings that have all the landmarks intact. For both wing centroid size and mean landmark determination, use the R script provided in Supplementary File 2, modifying for the .tps file location and image scale as per the experiment.</li>
</ol>
3. Assessing egg fertility
<ol>
	<li>Hatch individual F1 eggs from the papers collected from single females (collected in step 1) into polypropylene clear deli containers containing ~100 mL of freshly sterilized deionized water. Add two or three drops of ground fish food slurry for ad libitum feeding. Line the tops of the containers with mesh or organdy, and secure with a rubber band.</li>
	<li>At 2-3 days post hatching, remove the egg papers from the water and allow them to dry overnight by placing them into a new container that is covered with a lid or netting and leaving them in the insectary.</li>
	<li>Re-hatch the egg papers in the same containers in which they were initially hatched by placing them back into the water container. 
	NOTE: Drying and re-hatching the egg papers greatly improves the hatch efficiency. This may not be necessary for other mosquito species.</li>
	<li>At 3-5 days post-initial hatching, visually inspect the larvae (2nd to 4th instar) for a transgenic marker (if applicable; Figure 6). Inspect several larvae under a fluorescent microscope at the same time by placing them on a piece of filter paper placed on top of an organdy or mesh lined with paper towels. The paper towels absorb excess water, so that the filter paper can be used to screen and manipulate the immobilized larvae.</li>
	<li>Record the number of positive or negative transgenic larva. Discard the negative larvae.</li>
	<li>Clean the larval water to ensure optimal development times. To do this, pipette the larvae into a small cup, discard the dirty water, and replace it with fresh tap water Add the positive larvae and ensure enough ground fish food slurry is available.</li>
	<li>Calculate fertility as the number of transgenic or control larvae divided by the total number of eggs.</li>
</ol>
4. Determination of sex ratio in the pupae
<ol>
	<li>Up to 14 days post-hatching, continue collecting and recording the number of male and female pupae (as described in step 1.2.3). Females or males can be pooled, by sex, into small cups housed in 1.9 L containers.</li>
	<li>To determine sex ratio, add the number of F1 females or the number of F1 males collected across the study timeline per individual adult female mosquito. Divide this number by the total number of pupae collected for the same mosquito line generated from a single female.</li>
</ol>
5. Determination of larvae viability
<ol>
	<li>To determine larvae viability, add the total number of pupae collected across the study timeline per individual adult female mosquito.</li>
	<li>Divide this number by the total number of larvae counted per individual adult female mosquito for that same line counted in step 3.</li>
</ol>
6. Determination of larvae-to-pupae development time
<ol>
	<li>To determine larvae-to-pupae development, track the number of larvae and the number of pupae per line daily.</li>
	<li>Calculate larvae-to-pupae development as the average time to pupae development post-hatching.</li>
</ol>
7. Determination of male contribution
<ol>
	<li>Using age-matched males, anesthetize adult virgin transgenic male mosquitoes on ice, as well as adult virgin male and female control mosquitoes.</li>
	<li>Cross either the transgenic or control male adults with control female adults by adding 50 transgenic or 50 control males (age-matched) with 100 control females into 64 oz cartons, so that there are a total of 50 males with 100 females (two control females:one control or transgenic male). Be sure to label which carton contains transgenic males and which carton contains control males.</li>
	<li>Allow mating to occur for 2-3 days. After this time, offer mosquitoes a bloodmeal following standard rearing practices.</li>
	<li>Three days after blood feeding, separate individual fully engorged females into 50 mL conical tubes lined with pre-labeled white filter paper, as described in step 1 (fecundity assay). Discard the non-fed females or offer them another bloodmeal the next day. Fill all the tubes with ~5 mL of tap water. Place a small piece of raisin on the mesh placed on top of each conical tube.</li>
	<li>Leave females for 1-2 days in the insectary and allow them to oviposit. Visually inspect the egg papers for 4-5&#160;days post-bloodmeal for eggs. Allow the females an additional day to oviposit to increase the egg-laying rates.</li>
	<li>To collect the egg papers, drain the water and anesthetize the females by placing conical tubes at 4 &#176;C, as previously described. Discard female mosquitoes by placing in 70% ethanol.</li>
	<li>Leave the papers in tubes in the insectary for at least 5 days to dry. Use forceps to remove the papers and place them in polypropylene clear deli containers containing ~100 mL of freshly sterilized deionized water for hatching. Add 2-3 drops of ground fish food slurry for ad libitum feeding. Line the tops of the containers with mesh or organdy, secured with a rubber band.</li>
	<li>At 2-3 days post-hatching, remove the egg papers from the water and allow them to dry overnight by placing them into a new container that is covered with a lid or netting and placing them back into the insectary.</li>
	<li>Re-hatch the egg papers in the same containers by placing them back into the water container.</li>
	<li>At 3-5 days post-initial hatching, visually inspect the larvae (2nd to 4th instar) for a transgenic marker (if applicable). Calculate the male contribution as the number of F2 transgenic larvae divided by the number of total larvae per mosquito line.</li>
</ol>
8. Determination of mosquito longevity
<ol>
	<li>Transfer 50 male and 50 female WT or transgenic age-matched non-blood fed mosquitoes into a proper enclosure. Use 64 oz cartons, with the top opening lined with panty hose, twisted and knotted shut, for easy access and manipulation to the mosquitoes as the experiment progresses (Figure 7). 
	NOTE: It is recommended to do this in triplicate for statistical purposes.</li>
	<li>Provide a sugar and water source. Here, raisins and a water-filled cup lined with gauze, secured with two rubber bands were used.</li>
	<li>Track the number of dead male and female mosquitoes each day. Calculate longevity as the average number of days passed before mosquitoes die across the cages, omitting mosquitoes that die of non-informative causes (causes not due to mosquito lifespan, such as being squished or drowning in sugar) and omitting mosquitoes that are still alive at the end of the study. 
	NOTE: Survival is defined as the probability of being alive up to time t, or the function S(t) = Pr(T&#62;t). Median survival is the time at which the probability of survival is 0.5, or T_0.5 = S-1 x 0.5.</li>
</ol>

The authors have nothing to disclose.

Ae. aegypti fitness studies are often performed in the laboratory to assess fitness costs associated with transgenic cargo (e.g.,&#160;gene drive elements) or gene knock outs, as discussed in this manuscript; however, these studies may be performed for a variety of purposes&#8212;any that aim to evaluate the health of an Ae. aegypti group, such as Wolbachia-infected30,31, insecticide resistant32,33, or arbovirus-infected34. Therefore, because these experiments can be used for a wide array of purposes, the protocols were intended to be simple, efficient, and reproducible.
Investigators should use caution when extrapolating fitness data collected in the laboratory to mosquitoes in the field. In this protocol, the control group should be mosquitoes of the same genetic background but lacking the transgene (or treatment condition). However, common wild-type Ae. aegypti strains used in the laboratory have been inbred over many generations and thus may not be an accurate representation of those that are field-derived. For example, the common laboratory strain, Rockefeller (ROCK), is thought to have been established from Ae. aegypti collected in Cuba as early as 188135. Studies have shown that Rockefeller females survive significantly longer in the laboratory than field mosquitoes36. Another commonly used laboratory strain, particularly in transgenic studies, is Higgs&#39; White Eye (HWE), so named for Dr. Stephen Higgs, who selected these for their white eye mutant strain37. Because these mosquitoes lack eye pigmentation, transgenic expression of fluorescent markers is easier to detect than in wild-type Ae. aegypti,&#160;that have black eyes. However, these mosquitoes are significantly less fit than those from the field38. Laboratory fitness studies are also not an absolute representation of the conditions that mosquitoes endure in the field. Nonetheless, controlled laboratory fitness studies are a good indication of life parameters that may not otherwise be measurable. After performing fitness studies in the laboratory, larger cage studies may be necessary to evaluate fitness in semi-field conditions.
Although this manuscript is specific for Ae. aegypti, many of the protocols may be adapted for other species; however, we urge investigators to take into account the biology of their diverse species, which may differ in such a way that the protocols need slight modifications. For example, we found that hatching dried, individual egg papers twice&#8212;that is, hatching the egg papers once in water, drying the paper overnight, and re-hatching the same line again in water&#8212;was more efficient and a better representation of the hatching rates compared to those after a single hatch. However, it is important to note that Ae. aegypti eggs are resistant to desiccation39,40, whereas other mosquito species may not be tolerant to drying, including most species of Anopheles.
Perhaps the most critical, and time consuming, aspect of these protocols is accurately sexing male and female pupae. Adults mate 36-48 h post-emergence41, so it is critical that pupae are sexed correctly to ensure they are virgins for subsequent crosses. Females are monogamous, but males are polygamous and can effectively inseminate at least five females41,42,43. This means that a single male of an incorrect genotype can inseminate at least five females in a cage&#8212;rendering the cage relatively worthless for subsequent experiments&#8212;but a single female will not significantly impact a cage of males.
In summary, we have outlined basic protocols for measuring common life history traits in Ae. aegypti mosquitoes. Such studies are of paramount significance when assessing fitness costs associated with transgenes in genetically modified mosquitoes, as this data can be used in downstream applications such as modeling. For studies evaluating novel genetic control strategies, larger cage studies in semi-field conditions would then be necessary before deployment.

Survival of the fittest is the Darwinian idea that individuals who harbor genes best adapted to their environment will pass those genes to subsequent generations1. This means that it is fitness that determines whether their genes will survive. This more than 150-year-old concept is perhaps the most significant determinant of engineering a successful gene drive in transgenic mosquitoes. Gene drives, or the super-Mendelian inheritance pattern of a selfish genetic element that allows it to spread through populations2, are being explored for genetic pest management3. In the context of vector control, this strategy aims to either replace wild-type (WT) arthropods with those resistant to pathogens (population modification) or eliminate them all together (population suppression)4. However, transgenic mosquitoes often exhibit fitness costs (also called genetic load) in comparison to their WT counterparts, which means that the transgene will be lost in populations that can outcompete them. Coupling transgenes with a gene drive system is therefore necessary to offset any fitness costs and push the transgene through the population at levels greater than expected from typical Mendelian inheritance4.
Laboratory studies across mosquito vector species have shown that transgenes often display fitness costs5,6,7. For example, Irvin et al. measured various life parameters in Aedes (Ae.) aegypti engineered to express enhanced green fluorescent protein (eGFP) or transposase genes under Drosophila actin 5C or synthetic 3XP3 promoters, and compared them to the wild-type Orlando strain (the same genetic background from which they were derived)5. Most notably, they found that all transgenic strains had significantly reduced reproductive fitness5. Dependent on the transgene, some Anopheles (An.) mosquitoes expressing transgenes that inhibited Plasmodium parasite development also exhibited fitness costs6. Specifically, An. stephensi expressing a bee venom phospholipase (PLA2) under constitutive or bloodmeal-inducible promoters laid significantly fewer eggs compared to controls6. Those authors also found that transgenic Anopheles expressing a different transgene, an SM1 dodecapeptide tetramer did not exhibit fitness costs, leading them to conclude that transgene-conferred fitness costs are dependent on, at least likely in part, the effect of the transgenic protein produced6. Indeed, fitness costs may be attributed to transgene products, positional effects, off-target effects, insertional mutagenesis, or inbreeding effects in laboratory-reared strains7. Gene drives must therefore be robust enough to offset these transgene-induced fitness costs while also avoiding the development of insertions and deletions (indels) that block the drive itself.
Developmental or reproductive fitness costs can be measured in laboratory or cage studies7, with a caveat being that unknown factors in the field may also have an impact. Nonetheless, controlled fitness studies are an important first step when planning or evaluating genetically modified mosquito releases, such as a gene drive program, to determine whether the transgenic line will persist over subsequent generations. For example, Hammond et al. evaluated An gambiae CRISPR/Cas9-based-gene drives meant to disrupt the genes necessary for female fertility8. Through cage studies maintained for at least 25 generations, the authors found that the mosquitoes incurred gene-drive resistant alleles that blocked CRISPR-targeted cleavage and restored female fertility8. Their modeling efforts suggested that fertility in females heterozygous for the gene drive had the most dramatic impacts on gene drive fixation in simulated conditions8. Along these same lines, Ae. aegypti (Higgs&#39; White Eye strain, HWE) engineered to express autonomous gene drive cassettes under different promoters or at different intergenic loci exhibited different rates of gene drive fixation in simulated populations9. Using MGDrivE modeling10, combined with measured rates of indel formation, maternal effects, and laboratory-collected life parameter data, the authors found that mosquito fitness most strongly influenced the persistence of the gene drive in simulated conditions9. Fitness costs that are most likely to impair gene drive efficiency are attributable to somatic Cas9 (over) expression or from the gene that is targeted, particularly in heterozygotes, rather than the intrinsic drive itself11,12,13,14,15,16,17.
Given its significance, fitness is an important factor for the ability of a transgenic line to persist over subsequent generations, and it can be used as an indicator for any physiological effects associated with a transgene. For example, off-target effects may be associated with fitness costs. In this case, backcrossing a transgenic line for several generations is recommended. Furthermore, crossing the transgenic line with one that is reflective of a field population may also be necessary to investigate how well the transgenic population can compete in the real world. To quantify fitness costs so that they can be comparable, this manuscript provides simple protocols for measuring common life history traits in Ae. aegypti mosquitoes, with a particular emphasis on fitness costs associated with transgenes, so that such studies can be more easily reproduced. Protocols include fecundity, fertility, sex ratio, viability, development times, male contribution, and adult longevity. Measuring wing length and area was also chosen as a fitness measurement as it correlates with thorax length18,19 and body size measurements, which is directly linked to bloodmeal size, fecundity, and immunity20,21. Although there are many ways to assess fitness, these parameters were chosen because they reflect reproductive success, are simple to measure, and are commonly reported in the literature.

<table><tbody><tr><td>1 oz. translucent plastic souffle cups</td><td>WebstaurantStore</td><td>301100PC</td><td></td></tr><tr><td>2 oz. translucent polystyrene souffle cups</td><td>WebstaurantStore</td><td>760P200N</td><td></td></tr><tr><td>3 mL plastic pipettors&amp;nbsp;</td><td>Cornin</td><td>357524</td><td></td></tr><tr><td>50 mL conical tubes</td><td>Any brand</td><td></td><td></td></tr><tr><td>64 oz. white double poly-coated paper food cup</td><td>WebstaurantStore</td><td>999SOUP64WB</td><td>for mosquito enclosement</td></tr><tr><td>65 mm lens&amp;nbsp;</td><td>Canon</td><td>MP-E 65mm f/2.8 1-5x Macro Photo</td><td>Canon Macro Photo MP-E 65mm, 7D-65mm-1X; zoom=1, 200, 6.3, ISO=100; for photographing wings or egg papers, although other cameras are likely sufficient</td></tr><tr><td>Aedes aegypti&amp;nbsp;mosquitoes</td><td>BEI</td><td>multiple strains as eggs are available</td><td></td></tr><tr><td>Artifical membrane feeders</td><td>https://lillieglassblowers.com/</td><td>Meduim membrane feeder, Custom made, 33mm</td><td>Chemglass also offers, but sizes are wrong for us. Ours are about 3 cm?</td></tr><tr><td>ATP</td><td>MP Biomedical</td><td>ICN15026605</td><td>Any good quality ATP, 10mM filter sterile aliquots at -20</td></tr><tr><td>Autoclave</td><td></td><td></td><td>for sterilizing water for hatching</td></tr><tr><td>Canon EOS 7D camera&amp;nbsp;</td><td>Canon</td><td>3814B004</td><td>for photographing wings or egg papers, although other cameras are likely sufficient</td></tr><tr><td>defibrinated sheep blood&amp;nbsp;</td><td>Colorado Serum Co.</td><td>60 ml, every 2 weeks</td><td>https://colorado-serum-com.3dcartstores.com/sheep-defibrinated</td></tr><tr><td>Dual Gooseneck Microscope Illuminator</td><td>Dolan Jenner Fiber-Lite 180</td><td>181-1 System</td><td></td></tr><tr><td>Ethanol</td><td></td><td></td><td></td></tr><tr><td>Forceps</td><td>Dumont</td><td>5SF</td><td></td></tr><tr><td>Gauze</td><td></td><td>omnisorb ii</td><td>4&quot; non-woven sponges</td></tr><tr><td>glass microscope slide</td><td>Fisher Scientific</td><td>12-544-2</td><td></td></tr><tr><td>Glass Petri dishes, 100 &amp;times; 15 mm</td><td>VWR</td><td>75845-546</td><td>for anesthesizing/manipulating mosquitoes on ice</td></tr><tr><td>Hogs&#039; gut</td><td>Any Deli</td><td></td><td>we buy in bulk, split, wash and store in small aliquots of ~4X12&quot; at -20 in 50 ml conical</td></tr><tr><td>Ice</td><td></td><td></td><td></td></tr><tr><td>Ice bucket</td><td></td><td></td><td></td></tr><tr><td>Kimwipe</td><td>Fisher Scientific</td><td>06-666A</td><td></td></tr><tr><td>Leica GZ4 StereoZoom microscope</td><td></td><td></td><td>for screening transgenic mosquitoes (if applicable)</td></tr><tr><td>Paintbrush</td><td>AIT synthetic brush&amp;nbsp;</td><td>size 10-0</td><td>for manipulating larvae/pupae (Amazon)</td></tr><tr><td>Panty hose</td><td>Walmart</td><td>L&#039;eggs Everyday</td><td>&amp;nbsp;Women&#039;s Nylon &lt;strong&gt;Plus &lt;/strong&gt;Knee Highs Sheer Toe, 16 pairs (plus fits the carton)</td></tr><tr><td>Pencils</td><td>Any brand</td><td></td><td></td></tr><tr><td>Plastic containers for 2&amp;deg; storage of cartons</td><td>Walmart</td><td></td><td>Sterilite 58 Qt Storage Box Clear Base White Lid Set of 8</td></tr><tr><td>Plastic containers for growing larvae</td><td>Walmart</td><td></td><td>Sterilite 28 Qt. Storage Box Plastic, White, Set of 10</td></tr><tr><td>Plastic containers for hatching larvae</td><td>Walmart</td><td></td><td>Sterilite 6 Qt. Storage Box Plastic, White</td></tr><tr><td>polypropylene clear deli containers&amp;nbsp;</td><td>WebstaurantStore</td><td>&amp;nbsp;127DM12BULK</td><td>12 oz, or 16 oz if needed for bigger (127RD16BULK)</td></tr><tr><td>Rubber bands</td><td>Office Max</td><td>&amp;nbsp;#100736/#909606 /#3777415</td><td>12&quot;, #64 and #10</td></tr><tr><td>Rubber stopper</td><td>VWR</td><td>217-0515</td><td>for mosquito enclosement</td></tr><tr><td>Sugar source, such as sugar cubes or raisins</td><td></td><td></td><td></td></tr><tr><td>Tetramin flake food</td><td>Tetramin</td><td>16106</td><td></td></tr><tr><td>tpsDig</td><td>Stony Brook Morphometrics</td><td></td><td>A free morphometric image-processing software distributed online available at https://www.sbmorphometrics.org/</td></tr><tr><td>tpsUtil</td><td>Stony Brook Morphometrics</td><td></td><td>A free morphometric image-processing software distributed online available at https://www.sbmorphometrics.org/</td></tr><tr><td>White organza fabric 8&amp;rdquo; &amp;times; 8&amp;rdquo;</td><td>FabricWholesale.com&amp;nbsp;</td><td>4491676</td><td>Joann Casa Collection Organza Fabric by Casa Collection&amp;nbsp;</td></tr><tr><td>Whitman Grade 1 Qualitative Filter paper&amp;nbsp;</td><td>Whitman</td><td>1001-824</td><td>for egg papers. The white color makes it easier to see the black eggs.</td></tr></tbody></table>

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.

Following the above protocol, the fitness of two mosquito lines were evaluated: (1) CRISPR/Cas9-mediated knock out of the Ae. aegypti D7L1 (AAEL006424) salivary protein and (2) Ae. aegypti lines expressing autonomous CRISPR/Cas9-mediated gene drives9. In the case of the former, a homozygous D7L1 knock-out line was established by exploiting the non-homologous end-joining pathway (NHEJ) to generate the disruption after microinjecting embryos with sgRNAs specific to the D7L1 gene. Heterozygous D7L1-KO mosquitoes were backcrossed for four generations to WT before generating a stable homozygous line to remove potential off-target effects. In the case of the latter, a gene drive cassette expressing a Cas9 ribonucleoprotein under either the nanos (AAEL012107) or the innexin-4 (AAEL006726) promoters (GD1 and GD2 in this protocol, respectively) was knocked into the Carb109 locus (Chr3:409699138) by microinjecting embryos with a CRISPR/Cas9 injection mix, as described previously9. Positive G1 larvae were outcrossed to WT at least twice to establish the transgenic lines, and integration of the transgene was validated by PCR and Sanger sequencing of mosquito genomic DNA9.
Once the transgenic/mutant mosquito lines were established, fitness studies were performed following the protocol outlined above. The fitness of homozygous D7L1-KO Ae. aegypti was compared to WT Ae. aegypti Liverpool&#160;(LVP strain), while the fitness of hemizygote Ae. aegypti harboring a single copy of a gene drive cassette, generated by backcrossing the homozygous line to WT, was compared to the HWE&#160;strain. These comparison groups were chosen because they were of the same genetic background as their transgenic counterpart but lacking the transgene. At least 150-200 females per line were selected from the fecundity study (section 1) and allowed to oviposit for the subsequent sections to ensure at least 100 individual F1 egg papers would be obtained for statistical purposes. The life parameter data described below are defined as the following: (1) fecundity: the number of eggs one female oviposits in the first gonotrophic cycle; (2) wing length, area, and centroid size: distance from wing end-to-end, area of the wing, and a measurement of wing size independent of shape variation; (3) fertility: total number of larvae that develop from eggs oviposited after one gonotrophic cycle divided by the total number of eggs oviposited after one gonotrophic cycle; (4) sex ratio: total number of female or male pupae divided by total number of pupae; (5) larvae viability: total number of pupae divided by total number of larvae; (6) larvae-to-pupae development: time to pupa development post-egg hatching; (7) male contribution: number of transgenic-positive F2 larvae divided by total number of larvae; and (8) longevity: number of days adult mosquitoes survive while maintained on sugar.
Male mosquitoes lacking the D7L1 protein exhibited significantly delayed larvae-to-pupae and pupae-to-adult development times (Figure 8) and decreased survival times (Figure 9) as compared to LVP WT Ae. aegypti. Additionally, Ae. aegypti D7L1 knock-out mosquitoes had a significantly smaller wing area compared to their WT counterparts, but there were no differences in wing size or centroid size (Figure 10). This analysis suggests that the genetic modification did not alter mosquito body size but may have induced developmental stress that led to modified wing shape. On the other hand, mosquitoes harboring one copy of a gene drive cassette under expression of the nanos (GD1) or zpg (GD2) promoters did not have significantly different fitness when compared to the HWE line, with the notable exception of larva viability (Table 1). Ae. aegypti expressing a gene drive cassette under the nanos promoter exhibited a significantly lower percentage of larval survival (p = 0.035).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65136/65136fig01.jpg" /> 
Figure 1: Fitness study timeline. Estimated time required to perform all fitness study protocols described in this manuscript. Each life history parameter is highlighted in red. Abbreviations: TG = transgenic; WT = wild-type; O/N = overnight. Figure made with Biorender.com. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65136/65136fig01large.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/65136/65136fig02.jpg" /> 
Figure 2: Distinguishing female versus male pupae by genital lobes. Female pupae have extended fins past the genital lobes (black arrows) not seen in males. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65136/65136fig02large.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/65136/65136fig03.jpg" /> 
Figure 3: The 50 mL conical tubes serve as individual egg chambers. The&#160;50 mL conical tubes, lined with white filter paper (indicated by red arrow) that has been cut into rectangles, rolled, and pre-labeled in pencil, may serve as individual egg laying chambers. Mesh is applied to the top of the tube, secured with two rubber bands, where a sugar source may be applied. Nearly 5 mL of water is added into each tube, which serves as a water source. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65136/65136fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/65136/65136fig04.jpg" /> 
Figure 4: White filter paper serves as egg laying paper for individual female. White filter paper, cut into a rectangle, was removed from individual egg chambers and visually inspected for eggs, as seen here. Labeling the paper in pencil in the top left facilitates identifying the parental female. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65136/65136fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/65136/65136fig05.jpg" /> 
Figure 5: Aedes aegypti wing landmarks. A total of&#160;14 landmarks (shown in red) are used for wing digitization in the tpsDig software to generate two-dimensional Cartesian coordinates. The order of the wing coordinates must be maintained throughout the process of digitizing coordinates for all images. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65136/65136fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/65136/65136fig06.jpg" /> 
Figure 6: Transgenic larvae screening. Transgenic larvae engineered to express mCherry under a photoreceptor-specific promoter (3XP3) can be screened for positive fluorescence in their eyes (left), not seen in non-transgenic larvae (right). A stereoscope was used with a magnification between 0.5x-4x. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65136/65136fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/65136/65136fig07v2.jpg" /> 
Figure 7: Mosquito containment for longevity assay. A 64 oz carton, with the top opening lined with panty hose, twisted, and knotted shut, allowing for easy access and manipulation to the mosquitoes as the experiment progresses, serves as convenient mosquito containment for the longevity assay. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65136/65136fig07largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/65136/65136fig08.jpg" /> 
Figure 8:&#160;Male AeD7L1-knock out Ae. aegypti have delayed larvae-to-pupae and pupae-to-adult development times. Larvae-to-pupae and pupae-to-adult development times were assessed with a Kruskal-Wallis ANOVA and Dunn&#39;s post-hoc comparisons. WT male mosquitoes have a shorter time to pupation than WT females (Dunn&#39;s test, p &#60; 0.05), while D7L1 males do not significantly differ in their pupation time relative to D7L1 females (Dunn&#39;s test, p &#62; 0.05). D7L1 females take longer to reach pupation than WT females (Dunn&#39;s test, p &#60; 0.05), as do D7L1 males relative to WT males (Dunn&#39;s test, p &#60; 0.05). In terms of pupae-to-adult development time, WT and D7L1 females did not significantly differ. WT males had a faster time to pupation than WT females (p &#60; 0.05) and a faster time to pupation than D7L1 males (p &#60; 0.05). D7L1 males and females did not significantly differ (p &#62; 0.05). However, D7L1 females had a slower time than WT females (p &#60; 0.05). Abbreviations: WT = wild-type Liverpool strain; F = female; M = male; D7L1 KO = D7L1-knock out Ae. aegypti. N = 25 age-matched males and 25 females per mosquito line. * = p &#60; 0.05, Dunn&#39;s post-hoc test, Kruskal-Wallis ANOVA. ns = not significant. Bars represent mean + standard error. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65136/65136fig08large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 9" class="xfigimg" src="/files/ftp_upload/65136/65136fig09.jpg" /> 
Figure 9: WT (Liverpool strain) and D7L1-knock out Aedes aegypti males and females differed significantly in their survival. WT female mosquitoes never reached their median survival time during the study: p(survival) = 0.83 at 40 days. All other groups reached their median survival before 40 days, with a distinct temporal separation of the median survival times (WT males &#8776; 33 days, D7L1 females &#8776; 39 days, D7L1 males &#8776; 22 days). Kaplan-Meier survival curves were generated to model the survival of each line over time and estimate the median survival times, while accounting for censoring and non-normal survival distributions. A Log-Rank test was performed to evaluate between-group differences. The Log-Rank test assumes non-informative censoring and proportional hazards. The Cox Proportional Hazards Model was then used to evaluate the hazard ratio, or instantaneous risk of death at time t (given it has not already occurred) for each group relative to the other. The models&#39; proportional hazard assumption was met, verified by Schoenfeld&#39;s residuals (verified by the cox.zph function; p &#62; 0.05 for sex and line). N = 50 male and 50 female age-matched, non-bloodfed mosquitoes per Ae. aegypti line. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65136/65136fig09large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 10" class="xfigimg" src="/files/ftp_upload/65136/65136fig10.jpg" /> 
Figure 10:&#160;WT (Liverpool strain) and D7L1-knock out Ae. aegypti females differed significantly in wing area but not in wing length or centroid size. Wing length, area, and centroid size were measured after imaging the wings and digitizing the landmarks to assess for changes in size and shape. Wing area differed between WT and D7L1-knock outs, suggesting that the gene knock-out did not affect body size but may have induced developmental stress that altered wing shape. (A) Length: WT mean = 2.4 mm, &#963; = 0.21. D7L1 mean = 2.498 mm, &#963; = 0.35. Not significant (Mann-Whitney T-test, p = 0.6370). (B) Area: WT mean = 529,740 pixels/mm, &#963; = 58,852. D7L1 mean = 457,879 pixels/mm, &#963; = 143,633. Statistically significant difference (Mann-Whitney T-test, p=0.0158). C) Centroid Size: WT mean = 2321 pixels, &#963; = 61.08. D7L1 mean = 2,296 pixels, &#963; = 76.77, two outliers excluded. Not significant (Mann-Whitney T-test, p = 0.2993). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65136/65136fig10large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Line</td>
			<td>WT (HWE)</td>
			<td>GD1</td>
			<td>GD2</td>
		</tr>
		<tr>
			<td>larva-to-pupa development (days)</td>
			<td>9.8 &#177; 4.3</td>
			<td>9.5 &#177; 5.6</td>
			<td>9.5 &#177; 7.0</td>
		</tr>
		<tr>
			<td>larva viability (% survival)</td>
			<td>44.2 &#177; 7.3</td>
			<td>30.9 &#177; 2.9*</td>
			<td>44.8 &#177; 2.5</td>
		</tr>
		<tr>
			<td>% female adults</td>
			<td>45.4 &#177; 1.6</td>
			<td>47.8 &#177; 1.5</td>
			<td>48.2 &#177; 1.1</td>
		</tr>
		<tr>
			<td>% male adults</td>
			<td>54.6 &#177; 1.7</td>
			<td>52.2 &#177; 1.5</td>
			<td>51.8 &#177; 1.1</td>
		</tr>
		<tr>
			<td>50% female survival (days)</td>
			<td>34 &#177; 6.2</td>
			<td>44.5</td>
			<td>41.5</td>
		</tr>
		<tr>
			<td>50% male survival (days)</td>
			<td>20.7 &#177; 3.3</td>
			<td>16 &#177; 1.6</td>
			<td>18 &#177; 4.5</td>
		</tr>
		<tr>
			<td>fecundity (# eggs)</td>
			<td>62.6 &#177; 2.8</td>
			<td>57.6 &#177; 2.6</td>
			<td>64.6 &#177; 2.0</td>
		</tr>
		<tr>
			<td>fertility (egg hatchability) (%)</td>
			<td>49.2 &#177; 8.2</td>
			<td>44.0 &#177; 3.6</td>
			<td>58.3 &#177; 3.4</td>
		</tr>
		<tr>
			<td colspan="2">male contribution</td>
			<td>35.3 &#177; 1.02 
			(1931/5447)</td>
			<td>25.6 
			(1035/4049)</td>
		</tr>
	</tbody>
</table>
Table 1: Life parameter data for two Ae. aegypti lines (GD1 and GD2) hemizygous for gene drive cassettes compared to Higgs&#39; White Eye (HWE) control line. Data derived from hemizygous males or HWE males that were allowed to mate with female HWE. * = 0.01 &#60; p &#60; 0.05 compared to HWE by one-tailed t-test. No stars indicate no significant difference by one-way ANOVA. Means + standard errors are provided. This table has been modified from9.
Supplementary File 1: R code to calculate wing length and area. Code is provided under the name:&#160;Supplemental_File_R_Winglengtharea.R. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65136/Supplemental_File_R_Winglengtharea.zip" target="_blank">Please click here to download this File.</a>
Supplementary File 2: R code to determine wing centroid size and mean landmark. Code is provided under the name: Supplemental_File_R_Wingcentroidsize_meanlandmarks.R. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65136/Supplemental_File_R_Wingcentroidsize_meanlandmarks.zip" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Quantifying Fitness Costs in Transgenic Aedes aegypti Mosquitoes at JoVE.com

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.

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 ...

quantifying-fitness-costs-in-transgenic-aedes-aegypti-mosquitoes

Nanyang Technological University

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JoVE Journal

Biology

748 Views.  Colorado State University. Dit protocol demonstreert standaardmetingen, die een weerspiegeling zijn van fitheid, bij aedes aegypti-muggen, deze fitnessparameters zijn gekozen omdat ze reproductief succes weerspiegelen, eenvoudig te meten zijn en vaak worden gerapporteerd in de literatuur. Leg om te beginnen eierpapier een nacht in steriel water in het insectium om transgene en wilde muggen uit te broeden. Laat de larven zich ad libitum voeden door enkele druppels gemalen visvoerbrij toe te dienen.Om er zeker van...

Video: Quantifying Fitness Costs in Transgenic Aedes aegypti Mosquitoes

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.

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Environment

Reproducible dsRNA Microinjection and Oviposition Bioassay in Mosquitoes and House Flies

Synthetic dsRNAs, used to induce RNA interference, may have dose dependent phenotypic effects. These effects are difficult to define if the dsRNAs are delivered using a non-quantitative method. Accurate delivery of known quantities of nucleic acids or other chemicals is critical to measure the efficacy of the compound being tested and to allow reliable comparison between compounds.
Here we provide a reproducible, quantitative microinjection protocol that ensures accurate delivery of specific doses of dsRNA, reducing the mortality typically induced by injection injury. These modifications include the addition of Rhodamine B, a graduated injection needle, and an improved recovery method borrowed from Isoe and Collins. This method allows calculation of dose responses and facilitates comparisons between compounds. Versions of this method have been successfully used on three genera of mosquitoes as well as house flies to assess the reduction in fecundity resulting from gene silencing of ribosomal RNA transcripts.
This protocol provides strategies to reduce several challenges of small insect microinjection. Together, mechanical delivery of dsRNAs accompanied by visual verification, identification of effective locations for delivery, and inclusion of a post-injection recovery period ensure accurate dosing and low injury mortality. This protocol also describes an oviposition bioassay for uniform determination of effects on fecundity.

This protocol describes a microinjection methodology that we have standardized and used for several years to deliver specific quantities of nucleic acids directly to the hemolymph of mosquitoes and house flies. This protocol results in minimal injection mortality and allows dose correlated measurements of fecundity.

Synthetic dsRNAs, used to induce RNA interference, may have dose dependent phenotypic effects. These effects are difficult to define if the dsRNAs are ...

Small-Cage Laboratory Trials of Genetically-Engineered Anopheline Mosquitoes

Control of mosquito-borne pathogens using genetically-modified vectors has been proposed as a promising tool to complement conventional control strategies. CRISPR-based homing gene drive systems have made transgenic technologies more accessible within the scientific community. Evaluation of transgenic mosquito performance and comparisons with wild-type counterparts in small laboratory cage trials provide valuable data for the design of subsequent field cage experiments and experimental assessments to refine the strategies for disease prevention. Here, we present three different protocols used in laboratory settings to evaluate transgene spread in anopheline mosquito vectors of malaria. These include inundative releases (no gene-drive system), and gene-drive overlapping and non-overlapping generation trials. The three trials vary in a number of parameters and can be adapted to desired experimental settings. Moreover, insectary studies in small cages are part of the progressive transition of engineered insects from the laboratory to open field releases. Therefore, the protocols described here represent invaluable tools to provide empirical values that will ultimately aid field implementation of new technologies for malaria elimination.

The protocols reported here illustrate three alternative ways to assess the performance of genetically-engineered mosquitoes destined for vector control in laboratory-contained small cage trials. Each protocol is tailored to the specific modification the mosquito strain bears (gene drive or non-gene drive) and the types of parameters measured.

Control of mosquito-borne pathogens using genetically-modified vectors has been proposed as a promising tool to complement conventional control ...

Genetics

Genome Editing in the Yellow Fever Mosquito Aedes aegypti using CRISPR-Cas9

The emergence of the clustered, regularly interspersed, short palindromic repeats (CRISPR)-Cas9 technology has revolutionized the genetic engineering field and opened the doors for precise genome editing in multiple species, including non-model organisms. In the mosquito Aedes aegypti, loss-of-function mutations and DNA insertions have been accomplished with this technology. Here, we describe a detailed protocol for genome editing through embryonic microinjection in the mosquito A. aegypti using the CRISPR-Cas9 technology, focusing on both the generation of gene knockout and knockin lines. In this protocol, quartz needles are filled with a mixture of guide RNA, recombinant Cas9, and a plasmid containing a DNA cassette encoding a gene for a fluorescent marker, if gene knockin is desired. Embryos at the preblastoderm stage are lined up onto a strip of double-sided sticky tape placed onto a coverslip, which is subsequently mounted onto a glass slide. With the help of a microinjector, the needles are inserted gently into the posterior end of the embryos and a small volume of the CRISPR mixture is dispensed. When the embryos are hatched, the larvae are checked under the fluorescent scope, and the pupae are sex-sorted and separated in different cages. Once the adults emerge, these are reciprocally crossed with wild-type individuals, blood-fed, and placed for egg laying. Once these eggs are hatched, the fluorescent larvae collected represent individuals with stable insertion of the DNA cassette into their genome. These larvae are then grown to the adult stage, outcrossed to wild-type individuals, and then further assessed through molecular techniques to confirm that the exact sequence of the DNA cassette is present at the desired site of the mosquito genome. Homozygous lines can also be obtained by following the provided pipeline of crossing schema and molecular screening of the mutations.

Here, we describe a detailed protocol for genome editing through embryonic microinjection in the mosquito A. aegypti using the CRISPR-Cas9 technology.

The emergence of the clustered, regularly interspersed, short palindromic repeats (CRISPR)-Cas9 technology has revolutionized the genetic engineering ...

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 ...

Feeding and Quantifying Animal-Derived Blood and Artificial Meals in Aedes aegypti Mosquitoes

Females of certain mosquito species can spread diseases while biting vertebrate hosts to obtain protein-rich blood meals required for egg development. In the laboratory, researchers can deliver animal-derived and artificial blood meals to mosquitoes via membrane feeders, which allow for manipulation of meal composition. Here, we present methods for feeding blood and artificial blood meals to Aedes aegypti mosquitoes and quantifying the volume consumed by individual females.
Targeted feeding and quantification of artificial/blood meals have broad uses, including testing the effects of meal components on mosquito behavior and physiology, delivering pharmacological compounds without injection, and infecting mosquitoes with specific pathogens. Adding fluorescein dye to the meal prior to feeding allows for subsequent meal size quantification. The meal volume consumed by mosquitoes can be measured either by weight, if the females are to be used later for behavioral experiments, or by homogenizing individual females in 96-well plates and measuring fluorescence levels using a plate reader as an endpoint assay. Meal size quantification can be used to determine whether changing the meal components alters the meal volume ingested or if meal consumption differs between mosquito strains. Precise meal size quantification is also critical for downstream assays, such as those measuring effects on host attraction or fecundity. The methods presented here can be further adapted to track meal digestion over the course of days or to include multiple distinguishable markers added to different meals (like nectar and blood) to quantify the consumption of each meal by a single mosquito.
These methods allow researchers to singlehandedly perform high-throughput measurements to compare the meal volume consumed by hundreds of individual mosquitoes. These tools will therefore be broadly useful to the community of mosquito researchers for answering diverse biological questions.

Feeding and Quantifying Animal-Derived Blood and Artificial Meals in Aedes aegypti Mosquitoes

The goal of this protocol is to deliver animal-derived and artificial blood meals to Aedes aegypti mosquitoes through an artificial membrane feeder and precisely quantify the volume of meal ingested.

Females of certain mosquito species can spread diseases while biting vertebrate hosts to obtain protein-rich blood meals required for egg development. In ...

Immunology and Infection

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, ...

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 ...

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 ...

Quantifying Fitness Costs in Transgenic Aedes aegypti Mosquitoes

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