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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Artificial feeders</td>
			<td>Hemotek</td>
			<td>SP6W1-3</td>
			<td>Starter pack &#8211; 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&#34; - 0.216 m3 (60 cm3)</td>
		</tr>
		<tr>
			<td>Cage tub (popcorn)</td>
			<td>Amazon.com</td>
			<td>VP170-0006</td>
			<td>0.005 m3 (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,&#160; 0.4X4&#8221;</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&#34; Wide x 50 Yds - ~35.6 cm2 (14 in2)</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&#34;</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&#8221;) cut from bolt ~15 cm (6&#8221;) X ~23 m (25y)</td>
		</tr>
		<tr>
			<td>Zip ties</td>
			<td>Home Depot</td>
			<td>295715</td>
			<td>Pk of 100, 14&#8221; 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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2.5K 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

Flow-sorting and Exome Sequencing of the Reed-Sternberg Cells of Classical Hodgkin Lymphoma

The Hodgkin Reed-Sternberg cells of classical Hodgkin lymphoma are sparsely distributed within a background of inflammatory lymphocytes and typically comprise less than 1% of the tumor mass. Material derived from bulk tumor contains tumor content at a concentration insufficient for characterization. Therefore, fluorescence activated cell sorting using eight antibodies, as well as side- and forward-scatter, is described here as a method of rapidly separating and concentrating with high purity thousands of HRS cells from the tumor for subsequent study. At the same time, because standard protocols for exome sequencing typically require 100-1,000 ng of input DNA, which is often too high, even with flow sorting, we also provide an optimized, low-input library construction protocol capable of producing high-quality data from as little as 10 ng of input DNA. This combination is capable of producing next-generation libraries suitable for hybridization capture of whole-exome baits or more focused targeted panels, as desired. Exome sequencing of the HRS cells, when compared against healthy intratumor T or B cells, can identify somatic alterations, including mutations, insertions and deletions, and copy number alterations. These findings elucidate the molecular biology of HRS cells and may reveal avenues for targeted drug treatments.

Here, we describe a combined flow cytometric cell sorting and low-input, next-generation library construction protocol designed to produce high-quality, whole-exome data from the Hodgkin Reed-Sternberg (HRS) cells of classical Hodgkin lymphoma (CHL).

The Hodgkin Reed-Sternberg cells of classical Hodgkin lymphoma are sparsely distributed within a background of inflammatory lymphocytes and typically ...

Procedure for Adaptive Laboratory Evolution of Microorganisms Using a Chemostat

Natural evolution involves genetic diversity such as environmental change and a selection between small populations. Adaptive laboratory evolution (ALE) refers to the experimental situation in which evolution is observed using living organisms under controlled conditions and stressors; organisms are thereby artificially forced to make evolutionary changes. Microorganisms are subject to a variety of stressors in the environment and are capable of regulating certain stress-inducible proteins to increase their chances of survival. Naturally occurring spontaneous mutations bring about changes in a microorganism's genome that affect its chances of survival. Long-term exposure to chemostat culture provokes an accumulation of spontaneous mutations and renders the most adaptable strain dominant. Compared to the colony transfer and serial transfer methods, chemostat culture entails the highest number of cell divisions and, therefore, the highest number of diverse populations. Although chemostat culture for ALE requires more complicated culture devices, it is less labor intensive once the operation begins. Comparative genomic and transcriptome analyses of the adapted strain provide evolutionary clues as to how the stressors contribute to mutations that overcome the stress. The goal of the current paper is to bring about accelerated evolution of microorganisms under controlled laboratory conditions.

Here, we present a protocol to obtain adaptive laboratory evolution of microorganisms under conditions using chemostat culture. Also, genomic analysis of the evolved strain is discussed.

Natural evolution involves genetic diversity such as environmental change and a selection between small populations. Adaptive laboratory evolution (ALE) ...

High-Throughput Robotically Assisted Isolation of Temperature-sensitive Lethal Mutants in Chlamydomonas reinhardtii

Systematic identification and characterization of genetic perturbations have proven useful to decipher gene function and cellular pathways. However, the conventional approaches of permanent gene deletion cannot be applied to essential genes. We have pioneered a unique collection of ~70 temperature-sensitive (ts) lethal mutants for studying cell cycle regulation in the unicellular green algae Chlamydomonas reinhardtii1. These mutations identify essential genes, and the ts alleles can be conditionally inactivated by temperature shift, providing valuable tools to identify and analyze essential functions. Mutant collections are much more valuable if they are close to comprehensive, since scattershot collections can miss important components. However, this requires the efficient collection of a large number of mutants, especially in a wide-target screen. Here, we describe a robotics-based pipeline for generating ts lethal mutants and analyzing their phenotype in Chlamydomonas. This technique can be applied to any microorganism that grows on agar. We have collected over 3000 ts mutants, probably including mutations in most or all cell-essential pathways, including about 200 new candidate cell cycle mutations. Subsequent molecular and cellular characterization of these mutants should provide new insights in plant cell biology; a comprehensive mutant collection is an essential prerequisite to ensure coverage of a broad range of biological pathways. These methods are integrated with downstream genetics and bioinformatics procedures for efficient mapping and identification of the causative mutations that are beyond the scope of this manuscript.

High-Throughput Robotically Assisted Isolation of Temperature-sensitive Lethal Mutants in Chlamydomonas reinhardtii

Temperature-sensitive (ts) lethal mutants are valuable tools to identify and analyze essential functions. Here we describe methods to generate and classify ts lethal mutants in high throughput.

Systematic identification and characterization of genetic perturbations have proven useful to decipher gene function and cellular pathways. However, the ...

An Array-based Comparative Genomic Hybridization Platform for Efficient Detection of Copy Number Variations in Fast Neutron-induced Medicago truncatula Mutants

Mutants are invaluable genetic resources for gene function studies. To generate mutant collections, three types of mutagens can be utilized, including biological such as T-DNA or transposon, chemical such as ethyl methanesulfonate (EMS), or physical such as ionization radiation. The type of mutation observed varies depending on the mutagen used. For ionization radiation induced mutants, mutations include deletion, duplication, or rearrangement. While T-DNA or transposon-based mutagenesis is limited to species that are susceptible to transformation, chemical or physical mutagenesis can be applied to a broad range of species. However, the characterization of mutations derived from chemical or physical mutagenesis traditionally relies on a map-based cloning approach, which is labor intensive and time consuming. Here, we show that a high-density genome array-based comparative genomic hybridization (aCGH) platform can be applied to efficiently detect and characterize copy number variations (CNVs) in mutants derived from fast neutron bombardment (FNB) mutagenesis in Medicago truncatula, a legume species. Whole genome sequence analysis shows that there are more than 50,000 genes or gene models in M. truncatula. At present, FNB-induced mutants in M. truncatula are derived from more than 150,000 M1 lines, representing invaluable genetic resources for functional studies of genes in the genome. The aCGH platform described here is an efficient tool for characterizing FNB-induced mutants in M. truncatula.

An Array-based Comparative Genomic Hybridization Platform for Efficient Detection of Copy Number Variations in Fast Neutron-induced Medicago truncatula Mutants

This protocol provides experimental steps and information about reagents, equipment, and analysis tools for researchers who are interested in carrying out whole genome array-based comparative genomic hybridization (CGH) analysis of copy number variations in plants.

Mutants are invaluable genetic resources for gene function studies. To generate mutant collections, three types of mutagens can be utilized, including ...

Intraventricular Transplantation of Engineered Neuronal Precursors for In Vivo Neuroarchitecture Studies

Gene control of neuronal cytoarchitecture is currently the subject of intensive investigation. Described here is a simple method developed to study in vivo gene control of neocortical projection neuron morphology. This method is based on (1) in vitro lentiviral engineering of neuronal precursors as "test" and "control" cells, (2) their co-transplantation into wild-type brains, and (3) paired morphometric evaluation of their neuronal derivatives. Specifically, E12.5 pallial precursors from panneuronal, genetically labeled donors, are employed for this purpose. They are engineered to take advantage of selected promoters and tetON/OFF technology, and they are free-hand transplanted into neonatal lateral ventricles. Later, upon immunofluorescence profiling of recipient brains, silhouettes of transplanted neurons are fed into NeurphologyJ open source software, their morphometric parameters are extracted, and average length and branching index are calculated. Compared to other methods, this one offers three main advantages: it permits achieving of fine control of transgene expression at affordable costs, it only requires basic surgical skills, and it provides statistically reliable results upon analysis of a limited number of animals. Because of its design, however, it is not adequate to address non cell-autonomous control of neuroarchitecture. Moreover, it should be preferably used to investigate neurite morphology control after completion of neuronal migration. In its present formulation, this method is exquisitely tuned to investigate gene control of glutamatergic neocortical neuron architecture. Taking advantage of transgenic lines expressing EGFP in other specific neural cell types, it can be re-purposed to address gene control of their architecture.

Based on in vitro lentiviral engineering of neuronal precursors, their co-transplantation into wild-type brains and paired morphometric evaluation of "test" and "control" derivatives, this method allows accurate modeling of in vivo gene control of neocortical neuron morphology in a simple and affordable way.

Gene control of neuronal cytoarchitecture is currently the subject of intensive investigation. Described here is a simple method developed to study in ...

Generation of Tumor Organoids from Genetically Engineered Mouse Models of Prostate Cancer

Methods based on homologous recombination to modify genes have significantly furthered biological research. Genetically engineered mouse models (GEMMs) are a rigorous method for studying mammalian development and disease. Our laboratory has developed several GEMMs of prostate cancer (PCa) that lack expression of one or multiple tumor suppressor genes using the site-specific Cre-loxP recombinase system and a prostate-specific promoter. In this article, we describe our method for necropsy of these PCa GEMMs, primarily focusing on dissection of mouse prostate tumors. New methods developed over the last decade have facilitated the culture of epithelial-derived cells to model organ systems in vitro in three dimensions. We also detail a 3D cell culture method to generate tumor organoids from mouse PCa GEMMs. Pre-clinical cancer research has been dominated by 2D cell culture and cell line-derived or patient-derived xenograft models. These methods lack tumor microenvironment, a limitation of using these techniques in pre-clinical studies. GEMMs are more physiologically-relevant for understanding tumorigenesis and cancer progression. Tumor organoid culture is an in vitro model system that recapitulates tumor architecture and cell lineage characteristics. In addition, 3D cell culture methods allow for growth of normal cells for comparison to tumor cell cultures, rarely possible using 2D cell culture techniques. In combination, use of GEMMs and 3D cell culture in pre-clinical studies has the potential to improve our understanding of cancer biology.&#160;

We show a method for necropsy and dissection of mouse prostate cancer models, focusing on prostate tumor dissection. A step-by-step protocol for generation of mouse prostate tumor organoids is also presented.

Methods based on homologous recombination to modify genes have significantly furthered biological research. Genetically engineered mouse models (GEMMs) ...

Modulation of Tau Subcellular Localization as a Tool to Investigate the Expression of Disease-related Genes

Tau is a microtubule binding protein expressed in neurons and its main known function is related to the maintenance of cytoskeletal stability. However, recent evidence indicated that Tau is present also in other subcellular compartments including the nucleus where it is implicated in DNA protection, in rRNA transcription, in the mobility of retrotransposons and in the structural organization of the nucleolus. We have recently demonstrated that nuclear Tau is involved in the expression of the VGluT1 gene, suggesting a molecular mechanism that could explain the pathological increase of glutamate release in the early stages of Alzheimer&#39;s disease. Until recently, the involvement of nuclear Tau in modulating the expression of target genes has been relatively uncertain and ambiguous due to technical limitations that prevented the exclusion of the contribution of cytoplasmic Tau or the effect of other downstream factors not related to nuclear Tau. To overcome this uncertainty, we developed a method to study the expression of target genes specifically modulated by the nuclear Tau protein. We employed a protocol that couples the use of localization signals and the subcellular fractionation, allowing the exclusion of the interference from the cytoplasmic Tau molecules. Most notably, the protocol is easy and is composed of classic and reliable methods that are broadly applicable to study the nuclear function of Tau in other cell types and cellular conditions.

Tau is a neuronal protein present both in the cytoplasm, where it binds microtubules, and in the nucleus, where it exerts unconventional functions including the modulation of Alzheimer&#39;s disease-related genes. Here, we describe a method to investigate the function of nuclear Tau while excluding any interferences coming from cytoplasmic Tau.

Tau is a microtubule binding protein expressed in neurons and its main known function is related to the maintenance of cytoskeletal stability. However, ...

Generation of Defined Genomic Modifications Using CRISPR-CAS9 in Human Pluripotent Stem Cells

Human pluripotent stem cells offer a powerful system to study gene function and model specific mutations relevant to disease. The generation of precise heterozygous genetic modifications is challenging due to CRISPR-CAS9 mediated indel formation in the second allele. Here, we demonstrate a protocol to help overcome this difficulty by using two repair templates in which only one expresses the desired sequence change, while both templates contain silent mutations to prevent re-cutting and indel formation. This methodology is most advantageous for gene editing coding regions of DNA to generate isogenic control and mutant human stem cell lines for studying human disease and biology. In addition, optimization of transfection and screening methodologies have been performed to reduce labor and cost of a gene editing experiment. Overall, this protocol is widely applicable to many genome editing projects utilizing the human pluripotent stem cell model.

This protocol provides a method to facilitate the generation of defined heterozygous or homozygous nucleotide changes using CRISPR-CAS9 in human pluripotent stem cells.

Human pluripotent stem cells offer a powerful system to study gene function and model specific mutations relevant to disease. The generation of precise ...

Investigation of the Transcriptional Role of a RUNX1 Intronic Silencer by CRISPR/Cas9 Ribonucleoprotein in Acute Myeloid Leukemia Cells

The bulk of the human genome (~98%) is comprised of non-coding sequences. Cis-regulatory elements (CREs) are non-coding DNA sequences that contain binding sites for transcriptional regulators to modulate gene expression. Alterations of CREs have been implicated in various diseases including cancer. While promoters and enhancers have been the primary CREs for studying gene regulation, very little is known about the role of silencer, which is another type of CRE that mediates gene repression. Originally identified as an adaptive immunity system in prokaryotes, CRISPR/Cas9 has been exploited to be a powerful tool for eukaryotic genome editing. Here, we present the use of this technique to delete an intronic silencer in the human RUNX1 gene and investigate the impacts on gene expression in OCI-AML3 leukemic cells. Our approach relies on electroporation-mediated delivery of two preassembled Cas9/guide RNA (gRNA) ribonucleoprotein (RNP) complexes to create two double-strand breaks (DSBs) that flank the silencer. Deletions can be readily screened by fragment analysis. Expression analyses of different mRNAs transcribed from alternative promoters help evaluate promoter-dependent effects. This strategy can be used to study other CREs and is particularly suitable for hematopoietic cells, which are often difficult to transfect with plasmid-based methods. The use of a plasmid- and virus-free strategy allows simple and fast assessments of gene regulatory functions.

Investigation of the Transcriptional Role of a RUNX1 Intronic Silencer by CRISPR/Cas9 Ribonucleoprotein in Acute Myeloid Leukemia Cells

Direct delivery of preassembled Cas9/guide RNA ribonucleoprotein complexes is a fast and efficient means for genome editing in hematopoietic cells. Here, we utilize this approach to delete a RUNX1 intronic silencer and examine the transcriptional responses in OCI-AML3 leukemic cells.

The bulk of the human genome (~98%) is comprised of non-coding sequences. Cis-regulatory elements (CREs) are non-coding DNA sequences that contain binding ...

Screening and Identification of RNA Silencing Suppressors from Secreted Effectors of Plant Pathogens

RNA silencing is an evolutionarily conserved, sequence-specific gene regulation mechanism in eukaryotes. Several plant pathogens have evolved proteins with the ability to inhibit the host plant RNA silencing pathway. Unlike virus effector proteins, only several secreted effector proteins have showed the ability to suppress RNA silencing in bacterial, oomycete, and fungal pathogens, and the molecular functions of most effectors remain largely unknown. Here, we describe in detail a slightly modified version of the co-infiltration assay that could serve as a general method for observing RNA silencing and for characterizing effector proteins secreted by plant pathogens. The key steps of the approach are choosing the healthy and fully developed leaves, adjusting the bacteria culture to the appropriate optical density (OD) at 600 nm, and observing green fluorescent protein (GFP) fluorescence at the optimum time on the infiltrated leaves in order to avoid omitting effectors with weak suppression activity. This improved protocol will contribute to rapid, accurate, and extensive screening of RNA silencing suppressors and serve as an excellent starting point for investigating the molecular functions of these proteins.

Here, we present a modified screening method that can be extensively used to quickly screen RNA silencing suppressors in plant pathogens.

RNA silencing is an evolutionarily conserved, sequence-specific gene regulation mechanism in eukaryotes. Several plant pathogens have evolved proteins ...