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,</s...

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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&#160;</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&#160;</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&#160;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&#160;</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&#160;</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&#34; 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 &#215; 15 mm</td>
			<td>VWR</td>
			<td>75845-546</td>
			<td>for anesthesizing/manipulating mosquitoes on ice</td>
		</tr>
		<tr>
			<td>Hogs&#39; gut</td>
			<td>Any Deli</td>
			<td></td>
			<td>we buy in bulk, split, wash and store in small aliquots of ~4X12&#34; 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&#160;</td>
			<td>size 10-0</td>
			<td>for manipulating larvae/pupae (Amazon)</td>
		</tr>
		<tr>
			<td>Panty hose</td>
			<td>Walmart</td>
			<td>L&#39;eggs Everyday</td>
			<td>&#160;Women&#39;s Nylon Plus 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&#176; 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&#160;</td>
			<td>WebstaurantStore</td>
			<td>&#160;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>&#160;#100736/#909606 /#3777415</td>
			<td>12&#34;, #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&#8221; &#215; 8&#8221;</td>
			<td>FabricWholesale.com&#160;</td>
			<td>4491676</td>
			<td>Joann Casa Collection Organza Fabric by Casa Collection&#160;</td>
		</tr>
		<tr>
			<td>Whitman Grade 1 Qualitative Filter paper&#160;</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</em...

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

Research

JoVE Journal

Biology

714 Views.  Colorado State University. 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.

Video: Quantifying Fitness Costs in Transgenic Aedes aegypti Mosquitoes

Injection of dsRNA into Female A. aegypti Mosquitos

Reverse genetic approaches have proven extremely useful for determining  which genes underly resistance to vector pathogens in mosquitoes.  This video protocol illustrates a method used by the James lab to inject dsRNA into female A. aegypti mosquitoes, which harbor the dengue virus.  The technique for calibrating injection needles, manipulating the injection setup, and injecting dsRNA into the thorax is illustrated.

Reverse genetic approaches have proven extremely useful for determining which genes underly resistance to vector pathogens in mosquitoes. This video protocol illustrates a method used by the James lab to inject dsRNA into female A. aegypti mosquitoes, which harbor the dengue virus. The technique for calibrating injection needles, manipulating the injection setup, and injecting dsRNA into the thorax is illustrated.

Reverse genetic approaches have proven extremely useful for determining  which genes underly resistance to vector pathogens in mosquitoes.  This video ...

Microinjection of A. aegypti Embryos to Obtain Transgenic Mosquitoes

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

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

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

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

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

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

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

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

Dissection of Midgut and Salivary Glands from Ae. aegypti Mosquitoes

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


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

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

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

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

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

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

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Human cancer and response to therapy is better represented in orthotopic animal models. This paper describes the development of an orthotopic mouse model of ovarian cancer, treatment of cancer via oral delivery of drugs, and monitoring of tumor cell behavior in response to drug treatment in real time using in vivo imaging system. In this orthotopic model, ovarian tumor cells expressing luciferase are applied topically by injecting them directly into the mouse bursa where each ovary is enclosed. Upon injection of D-luciferin, a substrate of firefly luciferase, luciferase-expressing cells generate bioluminescence signals. This signal is detected by the in vivo imaging system and allows for a non-invasive means of monitoring tumor growth, distribution, and regression in individual animals. Drug administration via oral gavage allows for a maximum dosing volume of 10 mL/kg body weight to be delivered directly to the stomach and closely resembles delivery of drugs in clinical treatments. Therefore, techniques described here, development of an orthotopic mouse model of ovarian cancer, oral delivery of drugs, and in vivo imaging, are useful for better understanding of human ovarian cancer and treatment and will improve targeting this disease.

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Orthotopic animal models of ovarian cancer replicate better human disease and therefore enhance our understanding of cancer progression and tumor response to therapy. A mouse model receives an intrabursal injection of luciferase-expressing ovarian tumor cells. Treatment is administered via oral gavage. Tumor growth is monitored by in vivo imaging system.

Human cancer and response to therapy is better represented in orthotopic animal models.  This paper describes the development of an orthotopic mouse model ...

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 ...

A Quantitative Fitness Analysis Workflow

Quantitative Fitness Analysis (QFA) is an experimental and computational workflow for comparing fitnesses of microbial cultures grown in parallel1,2,3,4. QFA can be applied to focused observations of single cultures but is most useful for genome-wide genetic interaction or drug screens investigating up to thousands of independent cultures. The central experimental method is the inoculation of independent, dilute liquid microbial cultures onto solid agar plates which are incubated and regularly photographed. Photographs from each time-point are analyzed, producing quantitative cell density estimates, which are used to construct growth curves, allowing quantitative fitness measures to be derived. Culture fitnesses can be compared to quantify and rank genetic interaction strengths or drug sensitivities. The effect on culture fitness of any treatments added into substrate agar (e.g. small molecules, antibiotics or nutrients) or applied to plates externally (e.g. UV irradiation, temperature) can be quantified by QFA.
The QFA workflow produces growth rate estimates analogous to those obtained by spectrophotometric measurement of parallel liquid cultures in 96-well or 200-well plate readers. Importantly, QFA has significantly higher throughput compared with such methods. QFA cultures grow on a solid agar surface and are therefore well aerated during growth without the need for stirring or shaking.
QFA throughput is not as high as that of some Synthetic Genetic Array (SGA) screening methods5,6. However, since QFA cultures are heavily diluted before being inoculated onto agar, QFA can capture more complete growth curves, including exponential and saturation phases3. For example, growth curve observations allow culture doubling times to be estimated directly with high precision, as discussed previously1.
Here we present a specific QFA protocol applied to thousands of S. cerevisiae cultures which are automatically handled by robots during inoculation, incubation and imaging. Any of these automated steps can be replaced by an equivalent, manual procedure, with an associated reduction in throughput, and we also present a lower throughput manual protocol. The same QFA software tools can be applied to images captured in either workflow.
We have extensive experience applying QFA to cultures of the budding yeast S. cerevisiae but we expect that QFA will prove equally useful for examining cultures of the fission yeast S. pombe and bacterial cultures.

Quantitative Fitness Analysis (QFA) is a complementary series of experimental and computational methods for estimating microbial culture fitnesses. QFA estimates the effect of genetic mutations, drugs or other applied treatments on microbe growth. Experiments scaling from focussed analysis of single cultures to thousands of parallel cultures can be designed.

Quantitative Fitness Analysis (QFA) is an experimental and computational workflow for comparing fitnesses of microbial cultures grown in parallel1,2,3,4. ...

Transgenic Rodent Assay for Quantifying Male Germ Cell Mutant Frequency

De novo mutations arise mostly in the male germline and may contribute to adverse health outcomes in subsequent generations. Traditional methods for assessing the induction of germ cell mutations require the use of large numbers of animals, making them impractical. As such, germ cell mutagenicity is rarely assessed during chemical testing and risk assessment. Herein, we describe an in vivo male germ cell mutation assay using a transgenic rodent model that is based on a recently approved Organisation for Economic Co-operation and Development (OECD) test guideline. This method uses an in vitro positive selection assay to measure in vivo mutations induced in a transgenic &#955;gt10 vector bearing a reporter gene directly in the germ cells of exposed males. We further describe how the detection of mutations in the transgene recovered from germ cells can be used to characterize the stage-specific sensitivity of the various spermatogenic cell types to mutagen exposure by controlling three experimental parameters: the duration of exposure (administration time), the time between exposure and sample collection (sampling time), and the cell population collected for analysis. Because a large number of germ cells can be assayed from a single male, this method has superior sensitivity compared with traditional methods, requires fewer animals and therefore much less time and resources.

De novo mutations in the male germline may contribute to adverse health outcomes in subsequent generations. Here we describe a protocol for the use of a transgenic rodent model for quantifying mutations in male germ cells induced by environmental agents.

De novo mutations arise mostly in the male germline and may contribute to adverse health outcomes in subsequent generations. Traditional methods for ...

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

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

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

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

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