We are grateful to Drusilla Stillinger, Kiona Parker, Parrish Powell and Madeline Nottoli for mosquito husbandry. Funding was provided by the University of California Irvine Malaria Initiative. AAJ is a Donald Bren Professor at the University of California, Irvine.

Animal ethics statement 
This study was carried out in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. Protocols were approved by the Institutional Animal Care and Use Committees of the University of California (Animal Welfare Assurance Numbers A3416.01).
1. Inundative release trials on non-gene drive mosquitoes (Figure 1)
<ol>
	<li>Cage setup and maintenance
	<ol>
		<li>Set up three sets of triplicate 0.216 m3&#160;cages by adding 60 second-instar wild-type (WT) larvae in each cage over three successive weeks. 
		NOTE: It is not possible to determine the sex of second-instar larvae by light microscopy, so the samples added to each cage will consist of both males and females.</li>
		<li>At every week, provide adult females in each cage with anesthetized mice as bloodmeal source (Figure 2A) and an oviposition container 3 days after the bloodmeal. 
		NOTE: While an alternative artificial feeding apparatus can be used, providing live anesthetized mice for bloodmeals results in better mosquito feeding performance in these large (0.216 m3) cage formats. This requires an approved animal use protocol and relevant (e.g., Institutional Animal Care and Use Committee, IACUC) approval for using mice. 
		NOTE: Individual mice are anesthetized with a mixture of 4 mg/mouse ketamine HCl and&#160;0.4 mg/mouse xylazine. Animals are injected with between 0.1 - 0.5 mL of this mixture.</li>
		<li>Hatch eggs from each cage weekly and select 60 second-instar (L2) larvae at random to be returned to their respective cages to offset mortality (weeks 4-8). 
		NOTE: Steps 1.1.1 to 1.1.3 are necessary to establish a stable and distributed age-structured population in the cages - referred to as &#39;Initial Phase&#39;.</li>
		<li>At week 9, assign cages assembled in step 1.1.1 randomly in triplicates for releases of the desired male release ratio.
		<ol>
			<li>Designate one set of triplicate cages as controls to assess consistency throughout the experiment.</li>
			<li>Designate one set of triplicates for each desired release ratio (for example, 1:1 or 1:0.1 transgenic:WT males). 
			NOTE: This point on is referred to as &#39;Experimental Phase&#39;.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Replicates and release ratios
	<ol>
		<li>Add 60 WT pupae (30 males and 30 females) to the control cages weekly.</li>
		<li>To maintain a 1:1 ratio, add weekly 30 transgenic male pupae along with 60 (30 male and 30 female) WT pupae into each respective cage.</li>
		<li>To maintain a 1:0.1 ratio, add weekly 300 transgenic male pupae along with 60 (30 male and 30 female) WT pupae into each respective cage. 
		NOTE: Continued addition of wild mosquitoes to the cages maintains the cage density, which is expected to diminish weekly due to age-related adult mortality.</li>
	</ol>
	</li>
	<li>Screening of phenotypes
	<ol>
		<li>Select a total of 300 larvae from each cage at random. With the use of a stereo microscope equipped with fluorescence filters, screen for the expression of the fluorescent dominant marker at the larval and pupal stages and score the sex of the resulting adults (Figure 3). 
		NOTE: The phenotypical screening will depend on the dominant marker included in the transgene construct integrated into the mosquitoes (for example, Discosoma sp. red fluorescent protein [DsRed], cyan fluorescent protein [CFP], green fluorescent protein [GFP]), and on the promoter driving its expression (the most used in mosquito transgenesis is the 3xP3 promoter driving expression in the eyes and nerve cord).</li>
		<li>Follow this protocol for as many generations as required by the outcome parameters defined in the experimental design. 
		&#8203;NOTE: The trial is usually terminated when all mosquitoes have at least one copy of the transgene (determined by the presence of the dominant fluorescent marker) or the ratio of transgenic-to-WT mosquitoes in a cage is stabilized and does not fluctuate greatly after a few (3-5) generations.</li>
	</ol>
	</li>
</ol>
2. Overlapping generation trials of gene-drive mosquitoes (Figure 4)
NOTE: Mosquitoes carrying gene-drive systems require written and reviewed protocols and should be approved by an Institutional Biosafety Committee (IBC) or equivalent, and others where required. Mosquito containment (ACL 2+ level) should follow recommended procedures5,6,7. Specifically, the gene drive experiments should employ two stringent confinement strategies. The first is usually physical barriers (Barrier Strategy) between organisms and the environment. This requires having a secure insectary and standard operating procedures (including monitoring) for ensuring that mosquitoes cannot escape. The second confinement strategy can be Molecular, Ecological or Reproductive5.
<ol>
	<li>Cage setup
	<ol>
		<li>Set up two sets of triplicate 0.216 m3 cages for each desired transgenic:WT male release ratio.
		<ol>
			<li>To achieve a male release ratio of 1:1, add 120 transgenic males, 120 WT males and 120 WT females at the pupa stage to each replicate cage.</li>
			<li>To achieve a release ratio of 1:10, add 12 transgenic males, 120 WT males and 120 WT females at the pupa stage to each replicate cage. 
			&#8203;NOTE: Different release ratios can be tested (1:1, 1:3, 1:10, etc.) and the number of mosquitoes used to initiate the experiments varies accordingly. However, it is important to consider the effects of low numbers on the statistical evaluation of the data.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Population maintenance and screening
	<ol>
		<li>Provide 4-7 days old females in each cage with a blood meal using anesthetized mice (Figure 2A).</li>
		<li>Three days after the blood meal, insert an oviposition container in each cage.</li>
		<li>Hatch eggs in a larval tray, select ~240 first instar (L1) larvae at random from each cage, rear them to adulthood, and return them to their respective cages.</li>
		<li>Provide additional (2-3) blood meals every 3-4 days for the newly emerged adults as described in step 2.2.1. 
		NOTE: No additional transgenic males are added during any of the subsequent generations.</li>
		<li>Select a total of 300 larvae from each cage at random and screen them for the presence of the dominant marker phenotype at the larval and pupal stages using a fluorescence stereo microscope and score emerging adults for sex (Figure 3). 
		NOTE: As before, the phenotypical screening will depend on the dominant marker and promoter included in the gene-drive system and integrated into the transgenic mosquitoes (e.g., DsRed, CFP or GFP). If homozygous or heteroallelic disruptions of the targeted genes result in a visible phenotype (for example, genes related to eye pigmentation), screening of this trait will depend on which stage it is easiest to visualize the altered phenotype.</li>
		<li>Follow this protocol for as many generations as required by the outcome parameters defined in the experimental design. 
		&#8203;NOTE: Each generation (delimited by the blood meal) takes ~three weeks. The trial is usually terminated when all mosquitoes are deemed homozygous for the gene-drive construct or the populations stabilize at a maximum percentage of mosquitoes carrying at least one copy of the gene-drive construct.</li>
	</ol>
	</li>
</ol>
3. Non-overlapping generation trials of gene-drive mosquitoes (Figure 5).
<ol>
	<li>Cage setup
	<ol>
		<li>Set up triplicate 0.005 m3 cage populations for each specific release ratio of transgenics to WT males to be investigated (for example, three sets of triplicate cages each set up with 1:1, 1:3, 1:10 release ratios). Set up all cages with an equal total number of males and females. 
		NOTE: The Supplementary File is a video demonstrating the construction of the 0.005 m3 colony cage.
		<ol>
			<li>Add 50 transgenic males, 50 WT males, and 100 WT females to each of three replicate cages to achieve a 1:1 male release ratio.</li>
			<li>Add 25 transgenic males, 75 WT males, and 100 WT females to each of three replicate cages to achieve a 1:3 male release ratio.</li>
			<li>Add 9&#160;transgenic males, 90 WT males, and 100 WT females to each of three replicate cages to achieve a 1:10 male release ratio. 
			NOTE: Different release ratios can be tested and the number of mosquitoes used to initiate the experiments can vary accordingly. However, it is important to consider the impact of low numbers of mosquitoes on the statistical analyses. These are single releases; no additional transgenic males are added at any subsequent generation.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Population maintenance and screening
	<ol>
		<li>Provide the 4-7 days old females in each cage with blood meals using an artificial feeding apparatus (Figure 2B) on two consecutive days. 
		NOTE: Routine blood meals for females consist of a commercially available source of blood (e.g., calf&#39;s blood) provided from a feeding apparatus. Live anesthetized mice are used only to provide bloodmeals in larger (0.216 m3) cage formats for better feeding performance.</li>
		<li>Add an oviposition container 3 days after the second bloodmeal. After three days, remove the oviposition containers. 
		NOTE: At this step, 5-10 females can be selected at random from each cage and placed individually in vials to assess additional fitness parameters, such as fertility and fecundity, if needed.</li>
		<li>Score by sex all adults (dead and alive) remaining in the cage and store them at -80&#176;C for molecular analysis.</li>
		<li>Hatch eggs and randomly select 200 L1 larvae from the 1:1 and 1:3 ratio cages to populate new cages for the next generation. 
		NOTE: Due to the low frequency of starting transgenic individual in the 1:10 ratio cages, random sampling may lead to excessive loss of transgenic progeny in the next generation to carry on the population.</li>
		<li>To ensure an accurate sampling for the 1:10 cages and sufficient numbers of transgenic mosquitoes, screen all larvae for the dominant marker and select 200 larvae reflecting the observed transgene frequency to populate the new cages. 
		NOTE: The 1:10 cages can be maintained identically to 1:1 and 1:3 cages when they reach a transgene frequency of &#8805;80%.</li>
		<li>Select 500 larvae from each cage at random for an in-depth analysis. Screen under a fluorescence stereo microscope for the expected marker phenotypes at the larval and pupal stages and score sex of&#160;adults (Figure 3). 
		NOTE: &#39;Exceptional&#39; phenotypes can be selected to be further crossed and analyzed molecularly to monitor resistant allele formation.</li>
		<li>This protocol can be followed for as many generations as required by the outcome parameters defined in the experimental design. 
		NOTE: Each generation is delimited by the bloodmeal and takes ~three weeks. The trial is usually terminated when all mosquitoes are deemed homozygous for the gene-drive construct or populations stabilize at a maximum prevalence of transgenic mosquitoes. And as before, screening for phenotypes will depend on the dominant markers and promoter integrated in the transgenic mosquitoes (for example, DsRed, CFP, GFP) or in the targeted genes if they present a visible phenotype (for example, genes related to eye pigmentation).</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Carballar-Lejaraz&#250;, R., James, A. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Population+modification+of+Anopheline+species+to+control+malaria+transmission.">Population modification of Anopheline species to control malaria transmission.</a> Pathogens and Global Health. 111 (8), 424-435 (2017).</li><li>Pham, T. B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Experimental+population+modification+of+the+malaria+vector+mosquito,+Anopheles+stephensi.">Experimental population modification of the malaria vector mosquito, Anopheles stephensi.</a> PLOS Genetics. 15 (12), 1008440 (2019).</li><li>Carballar-Lejaraz&#250;, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Next-generation+gene+drive+for+population+modification+of+the+malaria+vector+mosquito,+Anopheles+gambiae.">Next-generation gene drive for population modification of the malaria vector mosquito, Anopheles gambiae.</a> Proceedings of the National Academy of Sciences. 117 (37), 22805-22814 (2020).</li><li>Adolfi, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficient+population+modification+gene-drive+rescue+system+in+the+malaria+mosquito+Anopheles+stephensi.">Efficient population modification gene-drive rescue system in the malaria mosquito Anopheles stephensi.</a> Nature Communications. 11 (1), 5553 (2020).</li><li>Akbari, O. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Safeguarding+gene+drive+experiments+in+the+laboratory.">Safeguarding gene drive experiments in the laboratory.</a> Science. 349 (6251), 927-929 (2015).</li><li>Benedict, M. Q., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Recommendations+for+Laboratory+Containment+and+Management+of+Gene+Drive+Systems+in+Arthropods.">Recommendations for Laboratory Containment and Management of Gene Drive Systems in Arthropods.</a> Vector-Borne and Zoonotic Diseases. 18 (1), 2-13 (2018).</li><li>Adelman, Z., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rules+of+the+road+for+insect+gene+drive+research+and+testing.">Rules of the road for insect gene drive research and testing.</a> Nature Biotechnology. 35 (8), 716-718 (2017).</li><li>Long, K. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Core+commitments+for+field+trials+of+gene+drive+organisms.">Core commitments for field trials of gene drive organisms.</a> Science. 370 (6523), 1417-1419 (2021).</li><li>Facchinelli, L., North, A. R., Collins, C. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Large-cage+assessment+of+a+transgenic+sex-ratio+distortion+strain+on+populations+of+an+African+malaria+vector.">Large-cage assessment of a transgenic sex-ratio distortion strain on populations of an African malaria vector.</a> Parasites Vectors. 12, 70 (2019).</li></ol>

The authors have no disclosures.

Genetically-engineered mosquitoes that have pathogen blocking ability or bear sterility genes constitute new tools to control vector-borne diseases. Given the multiplicity of parameters that comprise these alternative approaches, a critical step in their research consists of laboratory-confined experimental evaluations that allow a fast and safe prediction of the potential outcomes of a synthetic transgene release for control purposes1.
Because the monitoring of the transgene dynamics in caged populations can extend for several months, one of the central aspects of the protocols is the consistency in experimental design between replicates (including mosquito rearing, cage size, age-structured populations, fixed release ratios, stable blood meal sources and minimally invasive screening procedures).
Male-only releases are considered ideal because male mosquitoes neither transmit pathogens nor feed on humans, therefore they can safely introduce heritable characteristics into wild populations. In laboratory cage experiments, it is possible to detect transgenic strains with reduced male mating competitiveness and other fitness loads associated with transgene integration. However, direct and specific experiments, such as those conducted in large cages10, can be conducted to properly analyze male competitiveness, as well as female fecundity in more natural mosquito densities2. Furthermore, empirical data from the cage trials can be used to parameterize models of cage population dynamics, including resistant allele formation, and provide useful information on effectiveness and possible adjustments in the proposed technology.
The protocols described here can be easily adapted to other experimental designs as required, with minimal requirements regarding regular insectary infrastructure and conditions. In addition, except for the commercial cages and microscopes, most of the materials are inexpensive and allow low-cost multiple replicates and iterations of the trials. Notably, this also allows multiple transgenic strains to be pre-screened in small cage trials in order to prioritize best-performing candidates to be moved forward in the phased testing pathway and to suspend testing on those showing sub-optimal performances.
Finally, concern regarding the use of genetically modified organisms motivates the elaboration of frameworks for the development, evaluation, and application of genetic strategies for prevention of mosquito-borne diseases5,8,9. The relevance and execution of the protocols defined here are consistent with these guidelines.

Strategies based on genetically-engineered mosquitoes are being pursued to control transmission of vector borne pathogens such as those that cause malaria1. These include technologies 1) aimed at decreasing the numbers and densities of Anopheles mosquitoes (population suppression), or 2) aimed at impairing the ability of vectors to transmit parasites responsible for human disease (population modification, replacement, or alteration) wherein strains of vectors are engineered to express effector genes that prevent pathogen transmission. These genetic approaches have been bolstered by the advent of CRISPR/Cas9-based gene drives, with proofs-of-concept in parasite-transmitting mosquitoes of effective spread of payload traits as well as anti-parasitic effector molecules in caged populations.
Small laboratory cage trials represent a first step for evaluating the characteristic of transgenic strains as part of a phased approach to their further development towards field applications2. Specific outcome considerations include heritability of the introduced DNA in a competitive environment, penetrance and expressivity of the phenotype, and stability. Relevant experimental design features include the size of the cages, mosquito densities, number of replicates, overlapping or non-overlapping generations, age-structured target populations, single or multiple releases of engineered strains, male-only, female-only or mixed-sex releases, release ratios, blood meal sources (artificial or live animal), and screening procedures.
We describe here protocols used to evaluate strains of anopheline mosquitoes for inundative releases (no gene-drive system) and those that carry autonomous gene-drive systems mediated by Cas9 endonucleases and guide RNAs (gRNA). Applications of these protocols appear in Pham et al. (2019)2, Carballar-Lejaraz&#250; et al. (2020)3, and Adolfi et al. (2020)4.
Inundative release trials evaluate the spreading rate of a designed transgene under Mendelian inheritance following multiple releases of a large number of transgenic mosquitoes into a wild population. Without the attachment of the transgene to a drive system, data from inundative release trials provide&#160;information regarding the fitness and dynamic of the transgene of interest in a stabilized population.
When mosquito populations contain an autonomous gene-drive system, small cage trials are designed to assess the dynamics of the spread of the desired transgene by determining the rate of dominant marker increase following a single introduction of transgenic males. Autonomous gene-drive elements carry the genes encoding the Cas9 nuclease, gRNA and dominant marker linked in such a fashion as to be active in subsequent generations.
&#39;Overlapping&#39; generations refer to the simultaneous presence of multiple generations in the same cage to create an age-structured continuous population, while &#39;non-overlapping&#39; refers to single discrete generations in each consecutive caged population2. Gene-drive cage experiments can be terminated once the initial dynamics of the drive (conversion) rate can be determined (8-10 generations depending on the construct), and while they provide information on the short-term stability of the transgene within the mosquito population, they may not reveal what happens when and if the dominant marker frequencies reach or are close to full introduction (every mosquito carrying at least one copy of the gene-drive system).

<table><tbody><tr><td>Artificial feeders</td><td>Hemotek</td><td>SP6W1-3</td><td>Starter pack &amp;ndash; 6 feeders with 3ml reservoirs</td></tr><tr><td>Cage, commercial</td><td>BioQuip</td><td>1450D</td><td>Collapsible Cage, 24 X 24 X 24&quot; - 0.216 m&lt;sup&gt;3&lt;/sup&gt; (60 cm&lt;sup&gt;3&lt;/sup&gt;)</td></tr><tr><td>Cage tub (popcorn)</td><td>Amazon.com</td><td>VP170-0006</td><td>0.005 m&lt;sup&gt;3&lt;/sup&gt; (170 fl oz)</td></tr><tr><td>Dissecting microscope with fluorescence light and filters</td><td>Leica</td><td>M165FC</td><td></td></tr><tr><td>Glue sticks</td><td>Michaels</td><td>88646598807</td><td>Gluesticks 40 pk,&amp;nbsp; 0.4X4&amp;rdquo;</td></tr><tr><td>Hot glue gun</td><td>Woodwards Ace</td><td>2382513</td><td>Stanley, 40 watt, GR20</td></tr><tr><td>Nylon screen (netting)</td><td>Joann.com</td><td>1102912</td><td>Tulle 108&quot; Wide x 50 Yds - ~35.6 cm&lt;sup&gt;2&lt;/sup&gt; (14 in&lt;sup&gt;2&lt;/sup&gt;)</td></tr><tr><td>Oviposition cups</td><td>Fisher</td><td>259126</td><td>Beaker PP grad 50 mL</td></tr><tr><td>Razor cutting tool</td><td>Office Depot</td><td>487899</td><td>Box cutters</td></tr><tr><td>Scissors</td><td>Office Depot</td><td>978561</td><td>Scotch Precision Ultra Edge Titanium Non-Stick Scissors, 8&quot;</td></tr><tr><td>Stapler</td><td>Office Depot</td><td>908194</td><td>Swingline Commercial Desk Stapler</td></tr><tr><td>Surgical sleeve (stockinette)</td><td>VWR</td><td>56612-664</td><td>~48 cm (19&amp;rdquo;) cut from bolt ~15 cm (6&amp;rdquo;) X ~23 m (25y)</td></tr><tr><td>Zip ties</td><td>Home Depot</td><td>295715</td><td>Pk of 100, 14&amp;rdquo; cable ties - 35.6 cm (14 in)</td></tr></tbody></table>

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.

Transgenic anopheline&#160;mosquitoes generated to bear non-gene drive or autonomous gene-drive modifications are set up for cage trials as described in the Protocols section. The representative results shown here depict the phenotype dynamics of the best-performing replicates of each of the cage trials experiments performed by Pham et al. (2019)2 for Anopheles stephensi mosquitoes. The three trials (1 - 3, respectively: inundative non-gene drive, overlapping gene-drive and non-overlapping gene-drive) varied in different parameters, such as the size of the cage (0.216 m3 vs 0.005 m3), whether or not the target population was age-structured, source of blood meal (mice or artificial feeder) and release ratios. As a means of representation, Figure 6 displays the observed data selected from the same release ratio (1:1) for all three protocols used, on the course of seven generations.
The 1:1 non-drive release reaches &#62;80% transgene introduction within 6-7 generations. For gene-drive transgenic cage trials, the 1:1 releases in both the non-overlapping and overlapping protocols reach this level within 3-4 generations, thus, validating the expectation that a single release of a gene drive system can be more efficient than non-drive inundative releases for transgene introduction. The faster trajectory can also be confirmed by the slope of the trendlines. Both gene-drive protocols, despite different set ups, present similar angles and slope trends. At the end of observation, non-drive cages achieve ~80% of individuals bearing the transgene, while cages with gene-drive individuals reach complete (or near complete) introduction. Complete data and processing details on individual experiment results using the protocols described here can be found in Figures 1-3 of Pham et al. (2019)2, Figures 2-3 of Carballar-Lejaraz&#250; et al. (2020)3 and Figure 3&#160;of Adolfi et al. (2020)4.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62588/62588fig01.jpg" /> 
Figure 1. Non-drive inundative release trial schematic.&#160; Nine 0.216 m3 cages are set up with 60 wild-type second-instar (mixed-sexes) larvae added to each. Beginning week 3, females are provided a bloodmeal weekly and eggs are collected and hatched. Until week 8, 60 larvae are randomly selected and returned to their respective cages weekly to create an age-structured population in the cages (initial phase). Beginning week 9, the nine cages are randomly assigned in triplicate according to their transgenic:wild-type male release ratios (experimental phase). Cages A (Control) have no transgenic pupae added. Females are provided a bloodmeal weekly and eggs are collected, hatched, and reared to pupae. 30 male and 30 female wild- type pupae are added back to their cages. Cages 1:1 have an additional 30 transgenic male pupae added. Cages 1:0.1 have an additional 300 transgenic male pupae added. 300 larvae from each of the 9 cages are selected randomly and screened for the fluorescent marker. This procedure was repeated&#160;weekly until transgene fixation (stabilized ratio of transgenic-wildtype mosquitoes after a few generations). Adapted from Pham et al. (2019)2.&#160;<a href="https://www.jove.com/files/ftp_upload/62588/62588fig01large.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/62588/62588fig02.jpg" /> 
Figure 2. Blood feeding of cage populations.&#160;&#160;(A) Anesthetized mice or (B) Hemotek blood feeders are offered for blood feeding female mosquitoes on the 0.216 m3 cages or the small 0.005 m3 cages, respectively.&#160;<a href="https://www.jove.com/files/ftp_upload/62588/62588fig02large.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/62588/62588fig03.jpg" /> 
Figure 3. Screening phenotypes for non-drive, overlapping gene-drive and non-overlapping gene-drive cage trials.&#160; Fluorescent images of a larva, pupa and adult of transgenic or wild-type phenotypes. In this example, An. stephensi individuals were screened for the DsRed marker driven by the 3xP3 promoter in the eyes (DsRed+ or DsRed-), visible in all three stages, and adults were screened for sex (&#9792; or &#9794;). Note the background fluorescence in wild-type larvae associated with the food bolus in the midgut.&#160;<a href="https://www.jove.com/files/ftp_upload/62588/62588fig03large.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/62588/62588fig04.jpg" /> 
Figure 4. Overlapping gene-drive cage trial schematic.&#160;&#160;Six 0.216 m3 cages are set up in triplicate according to their gene-drive:wild-type male release ratios. 120 wild-type males and 120 wild-type females were added to each cage. Cages with a 1:1 gene-drive male release ratio had an additional 120 transgenic males added. Cages with a 1:10 male release ratio had an additional 12 transgenic males added. Until full introduction of the transgene, every 3 weeks, adult females are provided with bloodmeals and eggs are collected and hatched. A total of 240 larvae were selected randomly and returned to their respective cages. Three-hundred (300) larvae are selected randomly and screened for the dominant marker. They are later screened as pupae and adults for eye-color and sex. No additional transgenic males are added to the original cages. Adapted from Pham et al. (2019)2.&#160;<a href="https://www.jove.com/files/ftp_upload/62588/62588fig04large.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/62588/62588fig05.jpg" /> 
Figure 5. Non-overlapping gene-drive cage trial schematic.&#160; Nine small 0.005 m3 cages are set up in triplicate according to their gene-drive:wild-type male release ratios. Cages with a 1:1 male release ratio have 100 wild-type females, 50 wild-type males, and 50 gene-drive males added. Cages with a 1:3 male release ratio have 100 wild-type females, 75 wild-type males, and 25 gene-drive males added. Cages 1:10 male release ratio have 100 wild-type females, 90 wild-type males, and 9 gene-drive males added. Females are provided a blood meal and eggs collected and hatched. For 1:1 and 1:3 cages, 200 larvae are selected randomly and used to populate new cages, separate from that of their parents, for the next generation. An additional 500 larvae are selected randomly and reared to pupae, when they are screened for the dominant marker gene. The 500 pupae are then reared to adults and scored by sex. All remaining larvae are screened for the marker. For the 1:10 cages, all larvae are scored in generations 1-12 and 200 larvae reflecting the existing transgene frequency are used to populate new cages. Beginning at generation 13, these cages are set up identically to the 1:1 and 1:3 cages. Adapted from Pham et al. (2019)2 and Carballar-Lejaraz&#250; et al. (2020)3.&#160;<a href="https://www.jove.com/files/ftp_upload/62588/62588fig05large.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/62588/62588fig06.jpg" /> 
Figure 6. Predicted transgene fixation dynamics for the different population replacement cage trials.&#160;&#160;Representation of the expected phenotype dynamics of the best-performing replicates for each of the cage trials experiments performed by Pham et al. (2019)2, monitored over 7 generations. Experiments set ups are described in the Protocols. The predictions are based on data from all 9 experiments on the 1:1 release models (triplicate replicates for each of the three different cage trial protocols). The X-axis is the generation number after initial introduction and the Y-axis is the proportion of larvae showing the DsRed marker phenotype (DsRed+) over time. Dashed lines represent linear trendlines of the data. The DsRed+ phenotype results from having at least one copy of the modified allele. Hence the results reflect the spread of the transgene, expedited in the gene drive system, reaching (near) full introduction at the end of the observation. For the variability between replicates and full detailed data on the experiments, please refer to Pham et al. (2019)2, Carballar-Lejaraz&#250; R et al. (2020)3 and Adolfi A et al. (2020)4. Images adapted from Pham TB et al. (2019) Experimental population modification of the malaria vector mosquito, Anopheles stephensi. PLOS Genet 15(12): e1008440. doi: 10.1371/journal.pgen.1008440, Adolfi A et al. (2020) Efficient population modification gene-drive rescue system in the malaria mosquito Anopheles stephensi. Nat Commun 11(1): 5553. doi: 10.1038/s41467-020-19426-0 and Carballar-Lejaraz&#250; R et al. (2020) Next-generation gene drive for population modification of the malaria vector mosquito, Anopheles gambiae. Proc Natl Acad Sci USA 117(37):22805-22814. doi: 10.1073/pnas.2010214117.&#160;<a href="https://www.jove.com/files/ftp_upload/62588/62588fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplemental File: The construction of the 0.005 m3 colony cage.&#160;<a href="https://www.jove.com/files/ftp_upload/62588/62588 Supplementalvideo-1.m4v" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Small-Cage Laboratory Trials of Genetically-Engineered Anopheline Mosquitoes at JoVE.com

Small-Cage Laboratory Trials of Genetically-Engineered Anopheline Mosquitoes

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 ...

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Research

JoVE Journal

Genetics

2.6K Views.  University of California, Irvine. 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.

Video: Small-Cage Laboratory Trials of Genetically-Engineered Anopheline 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.

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

Environment

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 ...

Biology

Visualizing Efficacy of Pesticides Against Disease Vector Mosquitoes in the Field

Efficacy of public health pesticides targeting nuisance and disease-vector insects such as mosquitoes, sand flies, and filth-breeding flies is not uniform across ecological zones. To best protect public and veterinary health from these insects, the environmental limitations of pesticides need to be investigated to inform effective use of the most appropriate pesticide formulations and techniques. We have developed a research program to evaluate combinations of pesticides, pesticide application equipment, and application techniques in hot-arid desert, hot-humid tropical, warm and cool temperate, and urban locations to derive pesticide use guidelines specific to target insect and environment. To these ends we designed a system of protocols to support efficient, cost-effective, portable, and standardized evaluation of a diverse range of pesticides and equipment across multiple environments. At the core of these protocols is the use of an array of small cages with colony-reared sentinel mosquitoes (adults and immatures) and sand flies (adults), strategically arranged in natural habitats and exposed to pesticide spray. Spatial and temporal patterns of pesticide efficacy are derived from percent mortality in sentinel cages, then mapped and visualized in a geographic information system. Maps of sentinel mortality data may be statistically compared to evaluate relative efficacy of a pesticide across multiple environments, or to study multiple pesticides in a single environment. Protocols may be modified to accommodate a variety of scenarios, including, for example, the vertical orientation of sentinels in canopy habitats or simultaneous testing of ground and aerial application methods.

The efficacy of public health pesticides targeting nuisance and disease-vector insects is not uniform across different ecological zones. Here we present a system of techniques using captive vector insects as sentinels for pesticide efficacy to derive electronic maps supporting the standard evaluation of pesticides across multiple environments.

Efficacy of public health pesticides targeting nuisance and disease-vector insects such as mosquitoes, sand flies, and filth-breeding flies is not uniform ...

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 ...

Inoculating Anopheles gambiae Mosquitoes with Beads to Induce and Measure the Melanization Immune Response

The stimulation of immune responses is a common tool in invertebrate studies to examine the efficacy and the mechanisms of immunity. This stimulation is based on the injection of non-pathogenic particles into insects, as the particles will be detected by the immune system and will induce the production of immune effectors. We focus here on the stimulation of the melanization response in the mosquito Anopheles gambiae. The melanization response results in the encapsulation of foreign particles and parasites with a dark layer of melanin. To stimulate this response, mosquitoes are inoculated with beads in the thoracic cavity using microcapillary glass tubes. Then, after 24 hr, the mosquitoes are dissected to retrieve the beads. The degree of melanization of the bead is measured using image analysis software. Beads do not have the pathogenic effects of parasites, or their capacity to evade or suppress the immune response. These injections are a way to measure immune efficacy and the impact of immune stimulations on other life history traits, such as fecundity or longevity. It is not exactly the same as directly studying host-parasite interactions, but it is an interesting tool to study immunity and its evolutionary ecology.

Inoculating Anopheles gambiae Mosquitoes with Beads to Induce and Measure the Melanization Immune Response

Through inoculation with beads, the described technique enables the stimulation of the mosquito melanization response in the hemolymph circulating system. The amount of melanin covering the beads can be measured after dissection as a measure of the immune response.

The stimulation of immune responses is a common tool in invertebrate studies to examine the efficacy and the mechanisms of immunity. This stimulation is ...

Immunology and Infection

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, ...

Protocols for Testing the Toxicity of Novel Insecticidal Chemistries to Mosquitoes

New classes of insecticides with novel modes of action are needed to control insecticide resistant populations of mosquitoes that transmit diseases such as Zika, dengue and malaria. Assays for rapid, high-throughput analyses of unformulated novel chemistries against mosquito larvae and adults are presented. We describe protocols for single point-dose and dose response assays to evaluate the toxicity of small molecule chemistries to the Aedes aegypti vector of Zika, dengue and yellow fever,&#160;the malaria vector, Anopheles gambiae and the northern house mosquito, Culex quinquefasciatus, on contact and via ingestion. As an example, we evaluated the toxicity of amitriptyline, a small molecule antagonist of G protein-coupled receptors, via larval, adult topical and adult blood-feeding assay. The protocols provide a starting point to investigate insecticide potential. Results are discussed in the context of additional experiments to explore product applications and mechanisms for delivery.

Protocols are described for assessing the toxicity of chemistries to immature and adult mosquitoes for development as new classes of larvicides, adulticides and endectocides. The protocols enable high-throughput testing of multiple chemistries at single-point dose and subsequent evaluation via dose response assay to determine toxicity on contact or via ingestion.

New classes of insecticides with novel modes of action are needed to control insecticide resistant populations of mosquitoes that transmit diseases such ...

A Blood-Free Diet to Rear Anopheline Mosquitoes

Malaria research requires large-scale breeding and production conditions for mosquitoes (Anopheles spp.) in captivity. The sustainable and reliable production of mosquitoes is currently inhibited by the supply of fresh vertebrate blood. Alternatives to blood are required to promote efficient control strategies for malaria and other vector borne diseases that are transmitted by blood feeding insects. With this in mind, artificial liquid diets were formulated as substitutes for fresh vertebrate blood. Herein we report a blood-free artificial liquid diet that delivers feeding rates similar to blood and mimics the physiological effects of a fresh vertebrate blood meal. The diet induces ovarian and egg maturation of Anopheles mosquitoes and also produces good larval survival and development of functional adults. The formulated blood-free liquid diet is an important advance towards sustainable mosquito breeding in captivity and will reduce the maintenance costs of mosquito colonies and eliminate the need for fresh vertebrate blood.

A protocol is presented for formulation of a blood-free artificial diet to feed Anopheles mosquitoes in captivity. This diet has a similar performance to vertebrate blood and triggers oogenesis and egg maturation and produces viable adult progeny.

Malaria research requires large-scale breeding and production conditions for mosquitoes (Anopheles spp.) in captivity. The sustainable and reliable ...

Preparing and Injecting Embryos of Culex Mosquitoes to Generate Null Mutations using CRISPR/Cas9

Culex mosquitoes are the major vectors of several diseases that negatively impact human and animal health including West Nile virus and diseases caused by filarial nematodes such as canine heartworm and elephantasis. Recently, CRISPR/Cas9 genome editing has been used to induce site-directed mutations by injecting a Cas9 protein that has been complexed with a guide RNA (gRNA) into freshly laid embryos of several insect species, including mosquitoes that belong to the genera Anopheles and Aedes. Manipulating and injecting Culex mosquitoes is slightly more difficult as these mosquitoes lay their eggs upright in rafts rather than individually like other species of mosquitoes. Here we describe how to design gRNAs, complex them with Cas9 protein, induce female mosquitoes of Culex pipiens to lay eggs, and how to prepare and inject newly laid embryos for microinjection with Cas9/gRNA. We also describe how to rear and screen injected mosquitoes for the desired mutation. The representative results demonstrate that this technique can be used to induce site-directed mutations in the genome of Culex mosquitoes and, with slight modifications, can be used to generate null-mutants in other mosquito species as well.&#160;

Preparing and Injecting Embryos of Culex Mosquitoes to Generate Null Mutations using CRISPR/Cas9

CRISPR/Cas9 is increasingly used to characterize gene function in non-model organisms. This protocol describes how to generate knock-out lines of Culex pipiens, from preparing injection mixes, to obtaining and injecting mosquito embryos, as well as how to rear, cross, and screen injected mosquitoes and their progeny for desired mutations.

Culex mosquitoes are the major vectors of several diseases that negatively impact human and animal health including West Nile virus and diseases caused by ...

Effective Oral RNA Interference (RNAi) Administration to Adult Anopheles gambiae Mosquitoes

RNA interference has been a heavily utilized tool for reverse genetic analysis for two decades. In adult mosquitoes, double-stranded RNA (dsRNA) administration has been accomplished primarily via injection, which requires significant time and is not suitable for field applications. To overcome these limitations, here we present a more efficient method for robust activation of RNAi by oral delivery of dsRNA to adult Anopheles gambiae. Long dsRNAs were produced in Escherichia coli strain HT115 (DE3), and a concentrated suspension of heat-killed dsRNA-containing bacteria in 10% sucrose was offered on cotton balls ad-libitum to adult mosquitoes. Cotton balls were replaced every 2 days for the duration of the treatment. Use of this method to target doublesex (a gene involved in sex differentiation) or fork head (which encodes a salivary gland transcription factor) resulted in reduced target gene expression and/or protein immunofluorescence signal, as measured by quantitative Real-Time PCR (qRT-PCR) or fluorescence confocal microscopy, respectively. Defects in salivary gland morphology were also observed. This highly flexible, user-friendly, low-cost, time-efficient method of dsRNA delivery could be broadly applicable to target genes important for insect vector physiology and beyond.

Effective Oral RNA Interference RNAi Administration to Adult Anopheles gambiae Mosquitoes

The oral administration of dsRNA produced by bacteria, a delivery method for RNA interference (RNAi) that is routinely used in Caenorhabditis elegans, was successfully applied here to adult mosquitoes. Our method allows for robust reverse genetics studies and transmission-blocking vector studies without the use of injection.

Small-Cage Laboratory Trials of Genetically-Engineered Anopheline Mosquitoes

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

Microinjection of A. aegypti Embryos to Obtain Transgenic Mosquitoes

Visualizing Efficacy of Pesticides Against Disease Vector Mosquitoes in the Field

Reproducible dsRNA Microinjection and Oviposition Bioassay in Mosquitoes and House Flies

Inoculating Anopheles gambiae Mosquitoes with Beads to Induce and Measure the Melanization Immune Response

Maintaining Aedes aegypti Mosquitoes Infected with Wolbachia

Protocols for Testing the Toxicity of Novel Insecticidal Chemistries to Mosquitoes

A Blood-Free Diet to Rear Anopheline Mosquitoes

Preparing and Injecting Embryos of Culex Mosquitoes to Generate Null Mutations using CRISPR/Cas9

Effective Oral RNA Interference (RNAi) Administration to Adult Anopheles gambiae Mosquitoes