We are grateful to Kiona Parker (UCI) for providing images of transgenic An. stephensi larvae, and to Fraser Colman (LSTM) and Beth Poulton (LSTM) for providing transgenic An. gambiae larvae. Beth Poulton (LSTM) also provided precious assistance during the imaging of An. gambiae larvae. This work was funded by the Tata Institute for Genetics and Society (TIGS) and the LSTM&#39;s Director Catalyst Fund awarded to A.A. (DCF2014AA). A.A.J. is a Donald Bren Professor at the University of California,Irvine.

NOTE: A schematic workflow of the illustrated protocol is shown in Figure 2.
1. Design of &#966;C31 attB -tagged plasmids (Figure 3)
<ol>
	<li>Create attB donor plasmids carrying the following essential components
	<ol>
		<li>Dominant fluorescent marker
		<ol>
			<li>Choose a promoter to drive the expression of the fluorescent marker. 
			&#8203;NOTE: For Anopheles transgenesis, fluorescent markers are usually under the regulation of the 3xP3 promoter39, which drives expression in the eyes and nerve cord. Alternatively, the PUBc promoter5 can be used when expression in multiple tissues is desired. Donor plasmids and docking lines used as examples in this protocol are marked using the 3xP3 promoter.</li>
			<li>Choose a fluorescent protein (FP) that is compatible with that of the receiving docking line so that they are readily distinguishable. 
			&#8203;NOTE: Do not use the same marker that is already present in the docking line and avoid the simultaneous use of GFP (green)/YFP (yellow) and GFP (green)/CFP (cyan) as they are very difficult to reliably differentiate. Donor plasmids used as examples in this protocol are marked with either DsRed or YFP as they are to be integrated in a docking line marked with CFP.</li>
		</ol>
		</li>
		<li>attB recombination site(s)
		<ol>
			<li>Use a single attB site for integration of a transgenic cassette (single-attB design) (Figure 3A).</li>
			<li>Use two inverted attB sites for RMCE (double-attB design) where the sites lay inverted in respect to one another and enclose the donor DNA template (Figure 3B). 
			NOTE: The orientation of the attB site(s) must be compatible with that of the attP site(s) present in the docking line.</li>
		</ol>
		</li>
		<li>Desired transgene cargo
		<ol>
			<li>Use any other desired features to be integrated in the mosquito genome based on the specific purpose of the experiment. Here, we describe the integration of an antimalarial effector molecule into the genome of An. stephensi and the integration of the components of the GAL4/UAS system into An. gambiae mosquitoes.</li>
		</ol>
		</li>
		<li>Plasmid backbone components
		<ol>
			<li>Include, amongst other essential components for plasmid replication in bacteria, a marker for plasmid selection in vitro (i.e., an antibiotic resistance gene). 
			NOTE: The plasmid backbone will be integrated in the mosquito genome in the single-attB design for integration (Figure 3A), while it will not be inserted in the double-attB design for RMCE (Figure 3B).</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Preparation of plasmids for the microinjection mix
NOTE: The protocol illustrated here involves the use of two plasmids: an attB-tagged donor plasmid carrying the transgene of interest, and a helper plasmid that expresses the &#966;C31 integrase under the regulation of the Drosophila Hsp70 promoter40.
<ol>
	<li>Purify donor and helper plasmids using an endotoxin-free plasmid purification kit. 
	NOTE: Sequence the final plasmid preparation used for injection to verify the integrity of all components.</li>
	<li>Combine appropriate amounts of the two plasmids to obtain a mix with a final concentration of 350 ng/&#181;L of the donor plasmid and 150 ng/&#181;L of the helper plasmid when resuspended in injection buffer. 
	NOTE: When calculating the necessary volume of mix, consider that 10-15 &#181;L are sufficient for each day of planned injections and DNA can be prepared in advance and stored at -20 &#176;C. Integrase helper plasmid concentrations of 60-500 ng/&#181;L and donor plasmid concentrations of 85-200 ng/&#181;L have also been reported21,22,26,41.</li>
	<li>Precipitate the DNA by adding 0.1 volumes of 3 M sodium acetate (pH 5.2) and 2.5 volumes of ice-cold 100% EtOH and vortex. A white precipitate should be immediately visible. Having highly concentrated initial plasmid preparations (i.e., ~1 &#181;g/&#181;L) improves precipitation efficiency. 
	NOTE: Stopping point - The precipitate can be stored at -20 &#176;C overnight.</li>
	<li>Centrifuge at 15,000 x g for 20 min at 4 &#176;C, discard the supernatant, and wash the pellet with 1 mL of ice-cold 70% EtOH.</li>
	<li>Wash the pellet with 1 mL of ice-cold 70% EtOH and centrifuge at 15,000 x g for 5 min at room temperature.</li>
	<li>Discard the supernatant without disturbing the pellet and air dry.</li>
	<li>Resuspend the pellet in 1x injection buffer (0.1 mM Na3PO4, 5 mM KCl, pH 7.2, 0.22 &#181;m filter sterilized) to reach a total final concentration of 500 ng/&#181;L. 
	NOTE: Assume that some DNA will be lost during the precipitation process; therefore, add a smaller volume of injection buffer first, check the concentration on a spectrophotometer (e.g., Nanodrop), and then add an appropriate remaining volume to reach 500 ng/&#181;L.</li>
	<li>Ensure that the DNA is thoroughly resuspended, prepare aliquots of 10-15 &#181;L each and store them at -20 &#176;C.</li>
	<li>On the day of injection, thaw one aliquot and centrifuge at 15,000 x g for 5 min to remove any particulate residues. 
	&#8203;NOTE: An alternative method for particulate removal is to filter the solution through a 0.22 &#181;m filter. Avoid the presence of particulate residues in the injection mix as they lead to needle blockage during embryo microinjection.</li>
</ol>
3. Microinjection of embryos from an Anopheles docking line
<ol>
	<li>Blood feed 4-7-day-old mosquitoes from the desired docking line 72 h prior to microinjection (i.e., for injection on Monday and Tuesday feed females on the previous Friday; for injection on Thursday and Friday feed females on Monday of the same week).</li>
	<li>Blood feed wild-type (WT) mosquitoes (i.e., mosquitoes with the same genomic background of the docking line) on the same day; these will be needed for outcrossing. 
	NOTE: The size and quality of the blood meal affect egg quality, so it is recommended to always use fresh blood (i.e., blood drawn within the previous 7 days). Arm feeding or feeding on mice may increase the quality and quantity of eggs, however these methods are not encouraged. Specific approved protocols will be necessary for human and animal use.</li>
	<li>Perform embryo microinjections
	<ol>
		<li>Perform An. gambiae embryo microinjections in 25 mM NaCl42 by targeting the posterior pole of the embryo at a 45-degree angle. For a detailed protocol for embryo collection, alignment, and microinjection refer to Pondeville et al.43 and Lobo et al.44.</li>
		<li>Perform An. stephensi embryo microinjections in halocarbon oil 700:27 (2:1) by targeting the posterior pole of the embryo at a 30-degree angle. A detailed protocol for embryo collection, alignment, and microinjection can be found in Terenius et al.45 and Lobo et al.44.</li>
		<li>Transfer eggs immediately after injection in a Petri dish filled with sterile distilled water (pH 7.2) and return them to insectary conditions.</li>
	</ol>
	</li>
	<li>Upon hatching, transfer G0 larvae into a tray with salted distilled water (0.1% tonic salt) daily and rear to pupae.</li>
	<li>Record hatching rate (i.e., number of larvae hatched/number of embryos injected). 
	&#8203;NOTE: Embryo movement aids hatching, so gentle swirling is desirable. Hatching should start ~48 h after injection. Since injection may cause a slight developmental delay it is advisable to keep monitoring for late-hatching larvae for 3-4 days.</li>
</ol>
4. Crossing and screening of transformed individuals
<ol>
	<li>[OPTIONAL STEP] Screen G0 (injected) 1st or 2nd instar larvae (L1-L2) for transient expression of the fluorescent marker.
	<ol>
		<li>Use a fine-tip glass pipette to transfer G0 L1-L2 larvae to a microscope slides with wells. Place one larva in each well.</li>
		<li>Use a fluorescence stereoscope with the appropriate filter to screen for the presence of transient expression of the fluorescent marker. 
		NOTE: The pattern of transient expression is dictated by the promoter used. When using the 3xP3 promoter, transient expression of the fluorescent marker is visible in the anal papillae (see Figure 6 in Pondeville et al.43)</li>
		<li>Rear G0 positive individuals separately.</li>
	</ol>
	</li>
	<li>Sort G0 pupae by sex under a stereoscope52.</li>
	<li>Let males emerge in separate cages in groups of 3-5 (founder families) and add a 10-fold excess of age-matched WT females. 
	NOTE: Since males mate multiple times, it is important to provide an excess of WT females to maximize the mating chances of each male.</li>
	<li>Let females emerge in separate cages in groups of 10-15 (founder families) and add an equal number of age-matched WT males. 
	NOTE: If there is limited space in the insectary, females can emerge all together in a single cage. The female to male ratio can be as low as 1 male to 3 females.</li>
	<li>Allow adults to mate for 4-5 days and provide females with a blood meal. 
	&#8203;NOTE: Blood feed and collect eggs from G0 females multiple times to maximize the chances of getting transformants from multiple gonotrophic cycles.</li>
	<li>Blood feed WT individuals at the same time for outcrossing.</li>
	<li>Collect eggs and rear emerging next generation G1s.</li>
	<li>Screen G1 L3-L4 larvae for appropriate fluorescence to identify transformants.
	<ol>
		<li>Collect larvae in a Petri dish lined with filter paper or on a microscope slide and screen using a fluorescent stereoscope with appropriate filters for the presence of the marker introduced with the attB-tagged cargo. 
		&#8203;NOTE: Fluorescence driven by the 3xP3 promoter is visible in all postembryonic stages and the screening may be performed on younger larvae, however these are more fragile and must be handled relatively carefully. Pupae can also be screened.
		<ol>
			<li>For single-attB designs for integration screen for the presence of the new and pre-existing marker; they should both be present since the new cassette is inserted next to the original one (Figure 3A,&#160;Figure 4). 
			&#8203;NOTE: Screening exception for single attB designs: When using marker-less docking lines22, screen for the presence of the new marker only. When using docking lines where integration results in the inactivation of the pre-existing marker21, screen for the presence of the new marker and the loss of the pre-existing one.</li>
			<li>For double-attB designs for RMCE, screen for the presence of the new marker and the loss of the pre-existing one, only the newly introduced marker should be present since the new cassette replaces the original one (Figure 3B, Figure 5). 
			&#8203;NOTE: Occasional integration events can be recovered in RMCE experiments where only a single attP recombined and thus both markers will be present. The screening of G1 individuals can be carried out also at the pupa stage following the same procedure52.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Transfer transformed G1 individuals into a larval tray and rear to pupae. Discard non-fluorescent individuals and individuals with an unexpected marker expression pattern.</li>
	<li>Sort transformed G1 pupae by sex and cross them en masse with opposite-sex age-matched WT individuals.</li>
	<li>Allow adults to mate for 4-5 days, provide a blood meal, collect the eggs, and rear the next generation G2 progeny.
	<ol>
		<li>For single integration experiments, collect eggs directly from the en masse cross as the integration site is identical in all individuals.</li>
		<li>For RMCE experiments, collect eggs from single females and maintain progeny separate until molecular assessment is complete due to the potential presence of two alternative cassette orientations (Figure 3B).</li>
	</ol>
	</li>
	<li>Screen the G2 progeny (at either the larva or pupa stage) for the presence of the fluorescent marker (50% of the individuals are expected to be positive), discard non-fluorescent progeny.</li>
	<li>Set aside a subset of G2 positive individuals for molecular analysis, rear the rest to adulthood. 
	NOTE: If all G2 individuals must be kept alive, molecular analysis can be conducted on single adult&#39;s legs46 or pupal case DNA extractions (L. Grigoraki personal communication). Alternatively, molecular analysis can be performed after all the G2 individuals have oviposited and eggs have hatched.</li>
	<li>Allow adult males and females to intercross in the same cage to establish the new transgenic line. 
	&#8203;NOTE: For RMCE experiments, adult intercross must occur between siblings deriving from a single female until orientation of insertion is determined via molecular analysis.</li>
</ol>
5. Molecular validation of the insertion site by DNA amplification (PCR)
<ol>
	<li>Prepare a map of the predicted insertion site in the genome of the docking line after transformation.
	<ol>
		<li>Single integration: Ensure that the predicted insertion site carries the original docking construct plus the whole sequence of the donor plasmid between the two hybrid sites attL and attR (Figure 3A).</li>
		<li>RMCE: Ensure that the predicted insertion site is identical to that of the docking line where hybrid inverted attL sites replace the original inverted attP sites and the exchange template replaces the cassette originally present between them (Figure 3B).</li>
	</ol>
	</li>
	<li>Design oligonucleotide primers to amplify the insertional junction at either side of the integration locus.
	<ol>
		<li>Single integration: Design oligonucleotide primer pairs that span across the attR and/or attL sites. One primer must bind to the previously integrated docking construct and the other to the newly integrated transgene (Figure 3A).</li>
		<li>RMCE: Cassette replacement can occur in two different orientations with respect to the chromosome (designated A and B). Design alternative combinations of 4 oligonucleotide primers to give a discrete product in only one of the orientations, with one pair being diagnostic for orientation A, and the other for orientation B (Figure 3B, Figure 6).</li>
	</ol>
	</li>
	<li>Extract genomic DNA from G2 positive individuals and perform the diagnostic PCR and gel electrophoresis to visualize the presence of expected diagnostic amplicons from the predicted integration site maps. 
	NOTE: DNA may alternatively be extracted from single adult&#39;s legs46 or pupal cases (L. Grigoraki personal communication).</li>
	<li>Sequence PCR products to confirm expected sequences.</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62146/62146fig02.jpg" /> 
Figure 2. Workflow diagram for site-directed &#966;C31 genome modification in Anopheles mosquitoes. <a href="https://www.jove.com/files/ftp_upload/62146/62146fig02large.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/62146/62146fig03.jpg" /> 
Figure 3. Molecular basis of &#966;C31-mediated single integration (A) and RMCE (B). A) Schematic maps of the genomic insertion in an An. stephensi docking line (80.9, Table 1) carrying a single attP site and marked with CFP (top), a single-attB design donor plasmid marked with DsRed (middle), and the expected insertion site resulting after successful integration (bottom). B) Schematic maps of the genomic insertion in an An. gambiae docking line (A11, Table 1) carrying two inverted attP sites and marked with CFP (top), a double-attB design donor plasmid marked with YFP (middle), and the expected insertion site resulting after successful RMCE (bottom). Wavy line: mosquito genome; Striped arrows: piggyBac transposon arms; 3xP3: promoter of the fluorescent marker; SV40: viral terminator; Ori: origin of replication; AmpR: ampicillin resistance gene. Crossing lines represent the site(s) of recombination between attP and attB sites. Numbered black arrows represent primer binding sites for the molecular validation of the insertion locus (step 5 of the protocol). Fully annotated single and double attB-tagged plasmids are available from the authors upon request. <a href="https://www.jove.com/files/ftp_upload/62146/62146fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

The authors have nothing to disclose.

The accurate design of attB-tagged plasmids that are compatible with the docking line of choice is paramount for the success of the experiment. Careful consideration must be given to the choice of the marker used for the screening of transformants, including the fluorescence color and its pattern of expression, which will be subject to the pattern already present in the docking line. It is necessary to use fluorescent markers that are easily distinguishable: good marker combinations include RFP (red)/CFP (cyan), RFP (red)/GFP (green), RFP (red)/YFP (yellow), and YFP (yellow)/CFP (cyan), while combinations to avoid are YFP (yellow)/GFP (green) and CFP (cyan)/GFP (green). The 3xP3 promoter39, specific to the eyes and nerve cord, is the most frequently used to drive the expression of fluorescent markers for mosquito transgenesis. Indeed, all the Anopheles docking lines currently available utilize this promoter. Alternative regulatory regions are that of the An. gambiae polyubiquitin gene (PUBc)5 or the viral promoter IE120, which drive expression in multiple tissues. When used along with 3xP3, these promoters would expand the possible color combinations and even the use of the same fluorophore. The indicated promoters are active throughout the mosquito life cycle allowing screening and fluorescence monitoring at all life stages. An additional consideration during plasmid design is the size of the cargo to be integrated or exchanged. While the &#966;C31 system has remarkable carrying capacities18, it should be considered that the size of the donor plasmid generally correlates negatively with transformation efficiency22.
In the described protocol the source of integrase is a helper plasmid expressing the enzyme ubiquitously40. The ubiquitous presence of the integrase may lead to the transformation of somatic cells if microinjections are not precisely directed to the area where the germline forms. While such transformation events will be lost as they are not heritable, somatic effects can decrease the fitness of injected individuals. To avoid this and increase transformation efficiency, integrase expression can be restricted to the germline, for example by using the vasa promoter22,26. Other protocols describe the use of in vitro transcribed messenger RNA (mRNA) as source of &#966;C31 integrase19,24,43. However, this involves the laborious preparation of mRNA and requires careful handling of the injection mix and the use of RNase free reagents to avoid degradation. Plasmid sources of integrase have been demonstrated in both An. gambiae9,21,22,26,33,37 and An. stephensi (A.A. personal communication) to be reliable and lead to efficient transformation, and are thus our preferred option. A further option for integrase delivery is its in vivo production in self-docking helper lines. Such lines were created in An. gambiae that express the &#966;C31 integrase under the regulation of the germline-specific promoter nanos and were found to lead to an improved survival and transformation efficiency20. However, potential fitness loads imposed by the in vivo production of the integrase enzyme on the helper line must be considered.
As with other transgenic techniques, special care must be reserved to the rearing and crossing of individuals deriving from injected embryos to maximize the chances to recover transformants. Individuals that have stably inherited the transgene can be firstly recovered at the G1 progeny. However, early signs of potential transformation can be evaluated by the presence of transient episomal expression of the fluorescent marker in the anal papillae and/or nerve cord of G0 first and second instar larvae when using the 3xP3 promoter43. While the presence of transient fluorescence suggests successful plasmid delivery, it does not guarantee heritable germline transformation. Similarly, the lack of transient expression does not exclude successful transformation. Nevertheless, it has been observed that transiently positive individuals are more likely to yield transgenic progeny compared to transiently negative ones43,48. In expert hands, rearing and crossing of only positive individuals may be an option to reduce mosquito numbers. However, given the importance and fragility of small G0 larvae, the least amount of manipulation is still advisable and the rearing of all G0 individuals is always recommended.
The mating scheme reported in this protocol is designed to maximize the chance of mating and to isolate independent transformation events. However, if insectary space or personnel availability is an issue, G0 adults can be pooled by sex in single cages if enough opposite-sex individuals are provided. Such a setup will not allow discrimination between multiple transformation events occurring in individuals from the same cage. Depending on the experimental setup, the presence of a double (single integration) or single (RMCE) marker is expected during the screening process. In single integration experiments it is important to verify the presence of the original marker from the docking line, while in RMCE is important to verify the loss of the previously integrated marker. Indeed, it is not uncommon in RMCE designs to recover transformants in which single integration instead of exchange occurred due to the recombination of a single attP site9,33. In such individuals both fluorescent markers are present as well as the whole donor plasmid backbone highlighting the importance of conducting a thorough screening for both fluorescent markers.
While the presence of expected fluorescence patterns indicates successful transformation, molecular characterization of the insertion site must be undertaken. To do so, the preparation of accurate maps of the predicted insertion locus, including the flanking genomic regions of the docking line, is crucial for the design of adequate diagnostic oligonucleotide primers for gene amplification analyses. Single integration events result in the formation of attR and attL hybrid sites at the junction between the newly integrated DNA and the previously inserted cassette. These sites can be targeted for insertion site validation. In RMCE designs, the insertion of the donor cassette can occur in two alternative orientations in respect to the genomic locus, thus four primers can be used in alternative PCR combinations to detect which orientation the line carries. As the orientation of cassette insertion may affect transgene expression, in comparative gene expression analysis it is important to use lines carrying the same orientation of insertion.
When working with low numbers of transformants it may not be desirable to sacrifice whole individuals for molecular analysis. An option to this is conducting molecular analysis on DNA extracted from single adult&#39;s legs46 as leg loss does not affect an adult female ability to mate and oviposit49. However, there is a risk of damaging the individual in the process of leg removal. Success has been obtained using discarded pupal cases (L. Grigoraki personal communication), however the safest approach is to perform molecular analysis on G2 parents after obtaining viable G3 progeny.
In recent years, CRISPR/Cas9 has revolutionized the way of performing site-specific genome editing26,41,50,51. Unlike site-directed RMCE, CRISPR/Cas9-mediated gene integrations (knock-ins) are independent of the presence of pre-inserted recombination sites with only a one-step transformation event needed. Nevertheless, the CRISPR/Cas9 system relies on the presence of large known genomic sequences flanking the desired insertion site for successful homology directed repair as well as on the efficient site recognition mediated by guide RNAs. These conditions cannot always be met or may be laborious to troubleshoot and, given the availability of multiple docking lines in An. gambiae and An. stephensi and lines derived from them, the &#966;C31 system remains a very valuable tool to perform direct phenotypic comparisons between transgenes at the same genomic locations.

The ability to modify the genome of mosquito vectors of diseases reliably and reproducibly has bolstered in vivo functional validation of genes and opened the doors to realizable genetic vector control strategies, such as those targeting Anopheles mosquitoes that transmit malaria1.
Early mosquito genome editing relied solely on transposable element (TE)-mediated transformation, with piggyBac being the most commonly used transposon in Anopheles2,3,4. However, the random nature of TE integration can lead to undesirable modifications such as gene knockouts (insertional mutagenesis) and significant position effects on transgene expression5,6,7,8. Multiple insertions are also a common occurrence when using piggyBac5,9, which makes the validation and the isolation of transgenic lines with single insertions laborious. Other drawbacks include their potential remobilization, as observed in the germline of Anopheles stephensi when providing a source of piggyBac transposase10,11,12, and their limited size of DNA cargo (10-15 kb in length) with transformation efficiency declining with increasing size of the donor plasmid13,14.
Site-directed integration approaches were introduced to circumvent these issues. The most common site-directed genome modification in mosquitoes is that mediated by the &#966;C31 system (Figure 1a). This is driven by a viral integrase that catalyzes the recombination between two heterospecific attachment (att) sites occurring naturally in the genome of the bacteriophage &#966;C31 (attP) and in the Streptomyces bacterium host (attB)15. Recombination of the two sites is unidirectional and results in the formation of hybrid sites (attL and attR). The recombination of such hybrid sites (leading to DNA excision) would require not only the presence of an active viral integrase but also another phage-encoded recombination factor16,17. A stable integration site is thus generated that relieves the issue of potential undesired remobilization15. Moreover, the system allows the integration of large cargoes (e.g., integration of &#62;100 kb constructs was reported in D. melanogaster18), significantly increasing carrying capacities. Integration occurs in a single predefined genomic locus which greatly simplifies the validation of insertion and the mating scheme to obtain a stable transgenic line. Finally, the site-directed nature of the integration allows normalization of expression as alternative transgenes are located in the same locus and therefore are regulated within the same neighboring genomic context. Indeed, one of the main applications of the technique is the direct comparison of phenotypes conferred by different transgenes following insertion into an identical locus.
Achieving &#966;C31-mediated integration involves two phases: phase I is the creation of transgenic docking lines carrying attP site(s), and phase II is the site-directed integration of an attB-flanked cargo in the genome of the docking line19. The creation of phase I docking lines has relied on the TE-mediated random integration of attP-tagged constructs and thus involved an initial laborious process (including southern blot and inverse PCR analyses on single-female progeny) to isolate and validate transgenic lines carrying a single integration event in unique, transcriptionally active, and fitness neutral genomic locations. Nevertheless, several docking lines for &#966;C31-mediated single integration have been developed and validated in An. gambiae19,20,21,22 and in An. stephensi23,24,25&#160;(Table 1). Each of these lines varies in terms of the genomic location of the docking site and the strain-specific genetic background and from them a great variety of new transgenic lines can be created. The complex validation of TE-mediated integrations for producing docking lines can now be circumvented by the CRISPR/Cas9 technology26; however this relies on the a priori knowledge of neutral loci to be targeted and their surrounding sequences.
&#966;C31-mediated integration has been applied extensively to insect genome editing from the model organism D. melanogaster27, to the mosquitoes Aedes aegypti13,28, Ae. albopictus29, An. gambiae19, and An. stephensi24, as well as other insects including Ceratitis capitata30 and Bombyx mori31.
A limitation of &#966;C31-mediated integration, especially in view of potential field releases for vector control, is the integration in the mosquito genome of the entire attB-bearing donor plasmid, including undesirable sequences such as antibiotic-resistance gene markers and plasmid backbone components of bacterial origin. To address this, a modification of the standard system, recombinase-mediated cassette exchange (RMCE), was implemented that allows the precise replacement of a previously integrated transgenic cassette with a new donor DNA (Figure 1b). This is achieved by using two inverted att sites flanking the donor and recipient cassettes at each end, which drives two independent recombination events to take place simultaneously resulting in cassette exchange without integration of the plasmid backbone. This improved design circumvents the integration of undesired sequences and expands the application of &#966;C31 systems to include for example the integration of unmarked DNA cargos by screening for the loss of a previously integrated fluorescent marker32.
RMCE was achieved first with D. melanogaster32 and later applied successfully to non-model insects including An. gambiae9,26,33, Ae. aegypti34, Plutella xylostella34, and B. mori35. Several docking lines for RMCE have been developed and validated in An. gambiae5,9,26 (Table 1). To our knowledge, RMCE is yet to be explored in other Anopheles vectors species.
To date, the &#966;C31 system has been used widely in Anopheles mosquitoes to introduce and study a variety of molecules including antimalaria effectors19,24,36, components of the GAL4/UAS system to overexpress and knockdown genes for insecticide resistance studies9,33, regulatory elements, reporter genes5,21,37, and gene-drive elements26,38.
This protocol describes how to perform 1) site-directed integration of an attB-flanked cargo and 2) RMCE of a construct flanked by inverted attB sites into the genome of Anopheles docking lines. This is achieved by using two plasmids: a donor attB-tagged plasmid carrying the transgene of interest, and a helper plasmid expressing the &#966;C31 integrase. The major malaria vectors An. gambiae and An. stephensi are used as specific examples, however these protocols are applicable to other Anopheles species.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62146/62146fig01.jpg" /> 
Figure 1. Site-directed genome modifications, single integration and recombinase-mediated cassette exchange (RMCE) , using the&#160;&#966;C31&#160;system.&#160;The&#160;&#966;C31&#160;integrase (INT, grey double arrow) catalyzes the recombination between the&#160;attB&#160;site(s) (purple striped) present in a donor plasmid and the&#160;attP&#160;site(s) (blue striped) present in a receiving docking line, which results in the formation of hybrid sites&#160;attL&#160;and&#160;attR. A) Integration is achieved when single&#160;attB&#160;and&#160;attP&#160;sites recombine and results in the presence of two integrated markers (blue and red). B) RMCE occurs when two&#160;attB/P sites recombine simultaneously and results in the replacement of the cassette between the&#160;att&#160;sites of the docking line (blue marker) with that carried by the donor plasmid (red marker). C) Partial nucleotide sequences of&#160;attP&#160;(blue) and&#160;attB&#160;(purple) and the hybrid sites&#160;attL/R. Recombination occurs between the &#39;TT&#39; core sequences highlighted in bold black.&#160;<a href="https://www.jove.com/files/ftp_upload/62146/62146fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1.5 mL eppendorf tubes</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>8-well microslides</td>
			<td>VWR</td>
			<td>MARI1216690</td>
			<td></td>
		</tr>
		<tr>
			<td>DNeasy Blood &#38; Tissue Kit</td>
			<td>Qiagen</td>
			<td>69504</td>
			<td></td>
		</tr>
		<tr>
			<td>EndoFree&#160;Plasmid&#160;Maxi&#160;Kit (10)</td>
			<td>Qiagen</td>
			<td>12362</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol, Absolute, Molecular Biology Grade</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Filter set CFP for Leica MZ FLIII Excitation 436/20 nm, extinction 480/40 nm</td>
			<td>Leica</td>
			<td>10446363</td>
			<td></td>
		</tr>
		<tr>
			<td>Filter set dsRED for Leica MZ FLIII Excitation 545/30 nm, extinction 620/60 nm</td>
			<td>Leica</td>
			<td>10447079</td>
			<td></td>
		</tr>
		<tr>
			<td>Filter set YFP customised for Leica MZ FLIII</td>
			<td>Omega Optical</td>
			<td>500QM25, 500QM35</td>
			<td></td>
		</tr>
		<tr>
			<td>Halocarbon oil 27</td>
			<td>Sigma</td>
			<td>H8773</td>
			<td></td>
		</tr>
		<tr>
			<td>Halocarbon oil 700</td>
			<td>Sigma</td>
			<td>H8898</td>
			<td></td>
		</tr>
		<tr>
			<td>Petri dishes</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium&#160;chloride</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Chloride</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium phosphate dibasic</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium&#160;Acetate&#160;Solution (3 M),&#160;pH&#160;5.2</td>
			<td>Thermo Fisher Scientific (Life Technologies)</td>
			<td>R1181</td>
			<td></td>
		</tr>
		<tr>
			<td>Stable brush Size 0</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

site-directed &#966;c31-mediated integration and cassette exchange in anopheles vectors of malaria

Functional genomic analysis and related strategies for genetic control of malaria rely on validated and reproducible methods to accurately modify the genome of Anopheles mosquitoes. Amongst these methods, the &#966;C31 system allows precise and stable site-directed integration of transgenes, or the substitution of integrated transgenic cassettes via recombinase-mediated cassette exchange (RMCE). This method relies on the action of the Streptomyces &#966;C31 bacteriophage integrase to catalyze recombination between two specific attachment sites designated attP (derived from the phage) and attB (derived from the host bacterium). The system uses one or two attP sites that have been integrated previously into the mosquito genome and attB site(s) in the donor template DNA. Here we illustrate how to stably modify the genome of attP-bearing Anopheles docking lines using two plasmids: an attB-tagged donor carrying the integration or exchange template and a helper plasmid encoding the &#966;C31 integrase. We report two representative results of &#966;C31-mediated site-directed modification: the single integration of a transgenic cassette in An. stephensi and RMCE in An. gambiae mosquitoes. &#966;C31-mediated genome manipulation offers the advantage of reproducible transgene expression from validated, fitness neutral genomic sites, allowing comparative qualitative and quantitative analyses of phenotypes. The site-directed nature of the integration also substantially simplifies the validation of the single insertion site and the mating scheme to obtain a stable transgenic line. These and other characteristics make the &#966;C31 system an essential component of the genetic toolkit for the transgenic manipulation of malaria mosquitoes and other insect vectors.

The protocol illustrated here enables to generate a stable Anopheles transgenic line in ~10 weeks (assuming a 21-day mosquito life cycle).
Post-injection larval hatching rates in An. gambiae are expected to be generally lower than An. stephensi, however hatching rates between 10-50% have been reported9,20,24,26,33,43,47. Given appropriate injection technique, hatching rates of &#8805;20% are generally sufficient to yield transformants. DNA uptake by the embryos can be assessed by screening young larvae for transient expression of the fluorescent marker. In successful RMCE experiments in An. gambiae using the 3xP3 promoter up to 50% of the surviving G0 larvae showed episomal expression of the marker in the anal papillae48.
Generalized estimates of transformation efficiency are difficult to evaluate among laboratories and even among experiments as transformation depends on a complex interplay of variables including purity, concentration, size, and potential toxicity of the injected DNA, quality of eggs, pre- and post-injection handling of eggs, mosquito rearing, and most importantly the experience of the operator. Transformation rates up to 7% have been obtained for RMCE in An. gambiae (calculated as the number of independent transformation events in the total G0 individuals)9,26,33, and up to 2.2% transformation rate for integration in An. stephensi. We suggest injecting at least 500 embryos, which should lead to the hatching of at least 100 G0 larvae and to 2-7 G0 adult founders from which stably transformed progeny can be obtained. If screening for transient expression in G0 larvae, up to 40 positive larvae can be expected.
Examples of phenotypic validation of transformation via the screening of fluorescent markers regulated by the 3xP3 promoter are reported in Figure 4&#160;and&#160;Figure 5. Figure 4 shows a new An. stephensi line obtained by insertion of a DsRed-marked cassette into a docking line marked with CFP (80.9, Table 1), resulting in G1 progeny expressing both markers as indicated by the red and blue fluorescence detected in the eyes.
RMCE designs are instead expected to result in the replacement of the marker originally inserted into the docking line with that of the donor plasmid. Figure 5A and&#160;Figure B illustrate this marker exchange in an An. gambiae docking line marked with CFP (A11, Table 1) where after successful RMCE the CFP marker is lost and the YFP marker is acquired resulting in yellow (but not blue) eye and nerve cord fluorescence33. Occasionally, RMCE can result in a single integration event instead of the exchange of the desired transgenic cassette as illustrated in Figure 5C, where a larva marked with both the original CFP and the new YFP markers is shown. It is reported that up to 50% of the total number of transformation events are single integrations9, 33.
When screening for the presence of a fluorescent marker it is crucial to distinguish its signal from possible background autofluorescence. This is particularly important when using CFP as Anopheles larvae display natural blue autofluorescence (Figure 6A). Increasing the magnification and focusing on the tissues and organs where fluorescence is expected to be driven by the promoter is necessary to identify true CFP-positive individuals as illustrated in Figure 6B using the 3xP3-CFP marker.
Individual transformants are finally assessed molecularly via PCR to confirm the expected insertion site. Figure 7 reports the PCR validation in individuals from an exchange An. gambiae line showing the two potential orientations of insertion in the mosquito genome33.
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/62146/62146fig04.jpg" /> 
Figure 4. Validation of &#966;C31 single integration in An. stephensi larvae (dorsal view).&#160;A) The docking line (80.9, Table 1) expresses CFP in the eyes under the regulation of the 3xP3 promoter. B) Successful integration results in the expression of the newly acquired DsRed as well as the original CFP marker in the eyes. <a href="https://www.jove.com/files/ftp_upload/62146/62146fig04large.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/62146/62146fig05.jpg" /> 
Figure 5. Validation of &#966;C31 RMCE in An. gambiae larvae (ventral view).&#160;A) The docking line (A10, Table 1) expresses CFP under the regulation of the 3xP3 promoter in the eyes (e) and the nerve cord (nc)5. B) Successful RMCE results in the swap of fluorescent marker from CFP to YFP33. C) Single integration event occurred during RMCE experiment where the transformant larva expresses both the CFP and YFP markers. This larva carries GAL4/UAS components that cause a broad expression pattern of YFP, particularly strong in the abdominal muscles (am). <a href="https://www.jove.com/files/ftp_upload/62146/62146fig05large.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/62146/62146fig06.jpg" /> 
Figure 6. CFP autofluorescence in An. gambiae larvae (dorsal view). A) Side-by-side image of a positive (CFP+) and a negative (CFP-) L4 larva using the CFP filter. B) Close-up image of the larval eyes that reveals a CFP+ vs CFP- individual. <a href="https://www.jove.com/files/ftp_upload/62146/62146fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/62146/62146fig07.jpg" /> 
Figure 7. Molecular validation of the orientation of cassette insertion in representative transgenic An. gambiae created by &#966;C31 RMCE. The transgenic cassette can be inserted in one of two alternative orientations (A or B) in respect to the insertion site. Each PCR reaction (I - IV) uses a combination of primers (5-8)33 designed to give a diagnostic amplification fragment for each orientation as indicated in the schematic plasmid maps. T1: representative transgenic individual carrying orientation of insertion A; T2: representative transgenic individual carrying orientation of insertion B; WT: wild type; DL: docking line; -: reaction negative control where water was used as template.This figure has been modified from Adolfi et al. (2019)33.&#160;<a href="https://www.jove.com/files/ftp_upload/62146/62146fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Species</td>
			<td>Strain</td>
			<td>Name</td>
			<td>attP(s)</td>
			<td>Chromo-some</td>
			<td>Promoter-marker</td>
			<td>Institution of origin</td>
			<td>Reference</td>
		</tr>
		<tr>
			<td>An. stephensi</td>
			<td>Indiana</td>
			<td>26.10b</td>
			<td>Single</td>
			<td>2R</td>
			<td>3xP3-eCFP</td>
			<td>Univ. of California Irvine</td>
			<td>25</td>
		</tr>
		<tr>
			<td>An. stephensi</td>
			<td>Indiana</td>
			<td>44Cb</td>
			<td>Single</td>
			<td>X</td>
			<td>3xP3-eCFP</td>
			<td>Univ. of California Irvine</td>
			<td>23, 24</td>
		</tr>
		<tr>
			<td>An. stephensi</td>
			<td>Indiana</td>
			<td>80.9b</td>
			<td>Single</td>
			<td>2L</td>
			<td>3xP3-eCFP</td>
			<td>Univ. of California Irvine</td>
			<td>This study</td>
		</tr>
		<tr>
			<td>An. gambiae</td>
			<td>G3</td>
			<td>113</td>
			<td>Single</td>
			<td>2R</td>
			<td>3xP3-eCFP</td>
			<td>Univ. of California Irvine</td>
			<td>This study</td>
		</tr>
		<tr>
			<td>An. gambiae</td>
			<td>KIL</td>
			<td>Ec</td>
			<td>Single</td>
			<td>3R</td>
			<td>3xP3-eCFP</td>
			<td>Keele Univ.</td>
			<td>19, 43</td>
		</tr>
		<tr>
			<td>An. gambiae</td>
			<td>G3</td>
			<td>X1</td>
			<td>Single</td>
			<td>2L</td>
			<td>No marker</td>
			<td>Univ. of Strasbourg</td>
			<td>22</td>
		</tr>
		<tr>
			<td>An. gambiae</td>
			<td>G3</td>
			<td>YAttP</td>
			<td>Single</td>
			<td>Y</td>
			<td>3xP3-RFP</td>
			<td>Imperial College London</td>
			<td>21</td>
		</tr>
		<tr>
			<td>An. gambiae</td>
			<td>G3</td>
			<td>A10b</td>
			<td>Double</td>
			<td>2R</td>
			<td>3xP3-eCFP</td>
			<td>Liverpool School Trop. Med.</td>
			<td>5</td>
		</tr>
		<tr>
			<td>An. gambiae</td>
			<td>G3</td>
			<td>A11b</td>
			<td>Double</td>
			<td>2R</td>
			<td>3xP3-eCFP</td>
			<td>Liverpool School Trop. Med.</td>
			<td>9</td>
		</tr>
		<tr>
			<td colspan="8">a. Strain from Johns Hopkins University (gift of M. Jacobs-Lorena) and in culture at the Univ. of California Irvine for &#62;20 years.</td>
		</tr>
		<tr>
			<td colspan="8">b. These lines are available from the authors upon reasonable request.</td>
		</tr>
		<tr>
			<td colspan="8">c. This line is available at the BEI repository www.beiresources.org as MRA-1163.</td>
		</tr>
	</tbody>
</table>
Table 1. Anopheles attP docking lines.

Watch this Scientific Journal Video about Site-Directed Ãâ€ C31-Mediated Integration and Cassette Exchange in Anopheles Vectors of Malaria at JoVE.com

Site-Directed &#966;C31-Mediated Integration and Cassette Exchange in Anopheles Vectors of Malaria

The protocol describes how to achieve site-directed modifications in the genome of Anopheles malaria mosquitoes using the &#966;C31 system. Modifications described include both the integration and the exchange of transgenic cassettes in the genome of attP-bearing docking lines.

Functional genomic analysis and related strategies for genetic control of malaria rely on validated and reproducible methods to accurately modify the ...

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3.7K Views.  Liverpool School of Tropical Medicine. The protocol describes how to achieve site-directed modifications in the genome of Anopheles malaria mosquitoes using the φC31 system. Modifications described include both the integration and the exchange of transgenic cassettes in the genome of attP-bearing docking lines.

Video: Site-Directed φC31-Mediated Integration and Cassette Exchange in Anopheles Vectors of Malaria

Optogenetic Random Mutagenesis Using Histone-miniSOG in C. elegans

Forward genetic screening in model organisms is the workhorse to discover functionally important genes and pathways in many biological processes. In most mutagenesis-based screens, researchers have relied on the use of toxic chemicals, carcinogens, or irradiation, which requires designated equipment, safety setup, and/or disposal of hazardous materials. We have developed a simple approach to induce heritable mutations in C. elegans using germline-expressed histone-miniSOG, a light-inducible potent generator of reactive oxygen species. This mutagenesis method is free of toxic chemicals and requires minimal laboratory safety and waste management. The induced DNA modifications include single-nucleotide changes and small deletions, and complement those caused by classical chemical mutagenesis. This methodology can also be used to induce integration of extrachromosomal transgenes. Here, we provide the details of the LED setup and protocols for standard mutagenesis and transgene integration.

Optogenetic Random Mutagenesis Using Histone-miniSOG in C. elegans

Genetically-encoded histone-miniSOG induces genome-wide heritable mutations in a blue&#160;light-dependent manner. This mutagenesis method is simple, fast, free of toxic chemicals, and well-suited for forward genetic screening and transgene integration.

Forward genetic screening in model organisms is the workhorse to discover functionally important genes and pathways in many biological processes. In most ...

Rearing and Double-stranded RNA-mediated Gene Knockdown in the Hide Beetle, Dermestes maculatus

Advances in genomics have raised the possibility of probing biodiversity at an unprecedented scale. However, sequence alone will not be informative without tools to study gene function. The development and sharing of detailed protocols for the establishment of new model systems in laboratories, and for tools to carry out functional studies, is thus crucial for leveraging the power of genomics. Coleoptera (beetles) are the largest clade of insects and occupy virtually all types of habitats on the planet. In addition to providing ideal models for fundamental research, studies of beetles can have impacts on pest control as they are often pests of households, agriculture, and food industries. Detailed protocols for rearing and maintenance of D. maculatus laboratory colonies and for carrying out dsRNA-mediated interference in D. maculatus are presented. Both embryonic and parental RNAi procedures-including apparatus set up, preparation, injection, and post-injection recovery-are described. Methods are also presented for analyzing embryonic phenotypes, including viability, patterning defects in hatched larvae, and cuticle preparations for unhatched larvae. These assays, together with in situ hybridization and immunostaining for molecular markers, make D. maculatus an accessible model system for basic and applied research. They further provide useful information for establishing procedures in other emerging insect model systems.

Rearing and Double-stranded RNA-mediated Gene Knockdown in the Hide Beetle, Dermestes maculatus

Here, we present protocols for rearing an intermediate-germ beetle, Dermestes maculatus (D. maculatus) in the lab. We also share protocols for embryonic and parental RNAi and methods for analyzing embryonic phenotypes to study gene function in this species.

Advances in genomics have raised the possibility of probing biodiversity at an unprecedented scale. However, sequence alone will not be informative ...

Bidirectional Retroviral Integration Site PCR Methodology and Quantitative Data Analysis Workflow

Integration Site (IS) assays are a critical component of the study of retroviral integration sites and their biological significance. In recent retroviral gene therapy studies, IS assays, in combination with next-generation sequencing, have been used as a cell-tracking tool to characterize clonal stem cell populations sharing the same IS. For the accurate comparison of repopulating stem cell clones within and across different samples, the detection sensitivity, data reproducibility, and high-throughput capacity of the assay are among the most important assay qualities. This work provides a detailed protocol and data analysis workflow for bidirectional IS analysis. The bidirectional assay can simultaneously sequence both upstream and downstream vector-host junctions. Compared to conventional unidirectional IS sequencing approaches, the bidirectional approach significantly improves IS detection rates and the characterization of integration events at both ends of the target DNA. The data analysis pipeline described here accurately identifies and enumerates identical IS sequences through multiple steps of comparison that map IS sequences onto the reference genome and determine sequencing errors. Using an optimized assay procedure, we have recently published the detailed repopulation patterns of thousands of Hematopoietic Stem Cell (HSC) clones following transplant in rhesus macaques, demonstrating for the first time the precise time point of HSC repopulation and the functional heterogeneity of HSCs in the primate system. The following protocol describes the step-by-step experimental procedure and data analysis workflow that accurately identifies and quantifies identical IS sequences.

This manuscript describes the experimental procedure and software analysis for a bidirectional integration site assay that can simultaneously analyze upstream and downstream vector-host junction DNA. Bidirectional PCR products can be used for any downstream sequencing platform. The resulting data are useful for a high-throughput, quantitative comparison of integrated DNA targets.

Integration Site (IS) assays are a critical component of the study of retroviral integration sites and their biological significance. In recent retroviral ...

Retroviral Scanning: Mapping MLV Integration Sites to Define Cell-specific Regulatory Regions

Moloney murine leukemia (MLV) virus-based retroviral vectors integrate predominantly in acetylated enhancers and promoters. For this reason, mLV integration sites can be used as functional markers of active regulatory elements. Here, we present a retroviral scanning tool, which allows the genome-wide identification of cell-specific enhancers and promoters. Briefly, the target cell population is transduced with an mLV-derived vector and genomic DNA is digested with a frequently cutting restriction enzyme. After ligation of genomic fragments with a compatible DNA linker, linker-mediated polymerase chain reaction (LM-PCR) allows the amplification of the virus-host genome junctions. Massive sequencing of the amplicons is used to define the mLV integration profile genome-wide. Finally, clusters of recurrent integrations are defined to identify cell-specific regulatory regions, responsible for the activation of cell-type specific transcriptional programs.
The retroviral scanning tool allows the genome-wide identification of cell-specific promoters and enhancers in prospectively isolated target cell populations. Notably, retroviral scanning represents an instrumental technique for the retrospective identification of rare populations (e.g. somatic stem cells) that lack robust markers for prospective isolation.

Here, we describe a protocol for genome-wide mapping of the integration sites of Moloney murine leukemia virus-based retroviral vectors in human cells.

Moloney murine leukemia (MLV) virus-based retroviral vectors integrate predominantly in acetylated enhancers and promoters. For this reason, mLV ...

CRISPR/Cas9-mediated Targeted Integration In Vivo Using a Homology-mediated End Joining-based Strategy

As a promising genome editing platform, the CRISPR/Cas9 system has great potential for efficient genetic manipulation, especially for targeted integration of transgenes. However, due to the low efficiency of homologous recombination (HR) and various indel mutations of non-homologous end joining (NHEJ)-based strategies in non-dividing cells, in vivo genome editing remains a great challenge. Here, we describe a homology-mediated end joining (HMEJ)-based CRISPR/Cas9 system for efficient in vivo precise targeted integration. In this system, the targeted genome and the donor vector containing homology arms (~800 bp) flanked by single guide RNA (sgRNA) target sequences are cleaved by CRISPR/Cas9. This HMEJ-based strategy achieves efficient transgene integration in mouse zygotes, as well as in hepatocytes in vivo. Moreover, a HMEJ-based strategy offers an efficient approach for correction of fumarylacetoacetate hydrolase (Fah) mutation in the hepatocytes and rescues Fah-deficiency induced liver failure mice. Taken together, focusing on targeted integration, this HMEJ-based strategy provides a promising tool for a variety of applications, including generation of genetically modified animal models and targeted gene therapies.

CRISPR/Cas9-mediated Targeted Integration In Vivo Using a Homology-mediated End Joining-based Strategy

The clustered regularly interspaced short palindromic repeats/CRISPR associated protein 9 (CRISPR/Cas9) system provides a promising tool for genetic engineering, and opens up the possibility of targeted integration of transgenes. We describe a homology-mediated end joining (HMEJ)-based strategy for efficient DNA targeted integration in vivo and targeted gene therapies using CRISPR/Cas9.

As a promising genome editing platform, the CRISPR/Cas9 system has great potential for efficient genetic manipulation, especially for targeted integration ...

A Protocol for the Production of Integrase-deficient Lentiviral Vectors for CRISPR/Cas9-mediated Gene Knockout in Dividing Cells

Lentiviral vectors are an ideal choice for delivering gene-editing components to cells due to their capacity for stably transducing a broad range of cells and mediating high levels of gene expression. However, their ability to integrate into the host cell genome enhances the risk of insertional mutagenicity and thus raises safety concerns and limits their usage in clinical settings. Further, the persistent expression of gene-editing components delivered by these integration-competent lentiviral vectors (ICLVs) increases the probability of promiscuous gene targeting. As an alternative, a new generation of integrase-deficient lentiviral vectors (IDLVs) has been developed that addresses many of these concerns. Here the production protocol of a new and improved IDLV platform for CRISPR-mediated gene editing and list the steps involved in the purification and concentration of such vectors is described and their transduction and gene-editing efficiency using HEK-293T cells was demonstrated. This protocol is easily scalable and can be used to generate high titer IDLVs that are capable of transducing cells in vitro and in vivo. Moreover, this protocol can be easily adapted for the production of ICLVs.

We describe the production strategy of integrase-deficient lentiviral vectors (IDLVs) as vehicles for delivering CRISPR/Cas9 to cells. With an ability to mediate quick and robust gene editing in cells, IDLVs present a safer and equally effective vector platform for gene delivery compared to integrase-competent vectors.

Lentiviral vectors are an ideal choice for delivering gene-editing components to cells due to their capacity for stably transducing a broad range of cells ...

CRISPR/Cas9 Gene Editing to Make Conditional Mutants of Human Malaria Parasite P. falciparum

Malaria is a significant cause of morbidity and mortality worldwide. This disease, which primarily affects those living in tropical and subtropical regions, is caused by infection with Plasmodium parasites. The development of more effective drugs to combat malaria can be accelerated by improving our understanding of the biology of this complex parasite. Genetic manipulation of these parasites is key to understanding their biology; however, historically the genome of P. falciparum has been difficult to manipulate. Recently, CRISPR/Cas9 genome editing has been utilized in malaria parasites, allowing for easier protein tagging, generation of conditional protein knockdowns, and deletion of genes. CRISPR/Cas9 genome editing has proven to be a powerful tool for advancing the field of malaria research. Here, we describe a CRISPR/Cas9 method for generating glmS-based conditional knockdown mutants in P. falciparum. This method is highly adaptable to other types of genetic manipulations, including protein tagging and gene knockouts.

CRISPR/Cas9 Gene Editing to Make Conditional Mutants of Human Malaria Parasite P. falciparum

We describe a method for generating glmS-based conditional knockdown mutants in Plasmodium falciparum using CRISPR/Cas9 genome editing.

Malaria is a significant cause of morbidity and mortality worldwide. This disease, which primarily affects those living in tropical and subtropical ...

Characterizing Histone Post-translational Modification Alterations in Yeast Neurodegenerative Proteinopathy Models

Neurodegenerative diseases, such as amyotrophic lateral sclerosis (ALS) and Parkinson&#39;s disease (PD), cause the loss of hundreds of thousands of lives each year. Effective treatment options able to halt disease progression are lacking. Despite the extensive sequencing efforts in large patient populations, the majority of ALS and PD cases remain unexplained by genetic mutations alone. Epigenetics mechanisms, such as the post-translational modification of histone proteins, may be involved in neurodegenerative disease etiology and progression and lead to new targets for pharmaceutical intervention. Mammalian in vivo and in vitro models of ALS and PD are costly and often require prolonged and laborious experimental protocols. Here, we outline a practical, fast, and cost-effective approach to determining genome-wide alterations in histone modification levels using Saccharomyces cerevisiae as a model system. This protocol allows for comprehensive investigations into epigenetic changes connected to neurodegenerative proteinopathies that corroborate previous findings in different model systems while significantly expanding our knowledge of the neurodegenerative disease epigenome.

This protocol outlines experimental procedures to characterize genome-wide changes in the levels of histone post-translational modifications (PTM) occurring in connection with the overexpression of proteins associated with ALS and Parkinson&#39;s disease in Saccharomyces cerevisiae models. After SDS-PAGE separation, individual histone PTM levels are detected with modification-specific antibodies via Western blotting.

Neurodegenerative diseases, such as amyotrophic lateral sclerosis (ALS) and Parkinson&#39;s disease (PD), cause the loss of hundreds of thousands of lives ...

Determining the Egg Fertilization Rate of Bemisia tabaci Using a Cytogenetic Technique

A few species of sap-sucking whiteflies are some of the most damaging terrestrial pests worldwide because of the crop damage they inflict and plant viruses they vector. Despite numerous studies of the biology of these species in different environments, a key life history parameter, offspring sex ratios, has received little attention, yet is important for predicting population dynamics. The primary sex ratio (sex ratio at oviposition) of Bemisia tabaci has never been reported but can be found by determining the egg fertilization rate of this haplodiploid insect. The technique involves the dechorionation of eggs with bleach, a series of fixation steps, and the application of the general DNA fluorescent stain, DAPI (4&#8242;,6-diamidino-2-phenylindole, a DNA-binding fluorescent dye), to bind to female and male pronuclei. Here, we present the technique, and an example of its application, to test whether an endosymbiotic bacterium, Rickettsia sp. nr. bellii, influenced the primary sex ratio of B. tabaci. This method may assist in population studies of whiteflies, or in determining if sex allocation exists with certain environmental stimuli.

Determining the Egg Fertilization Rate of Bemisia tabaci Using a Cytogenetic Technique

We present a simple cytogenetic technique using 4&#8242;,6-diamidino-2-phenylindole (DAPI) to determine the fertilization rate and primary sex ratio of the haplodiploid invasive pest Bemisia tabaci.

A few species of sap-sucking whiteflies are some of the most damaging terrestrial pests worldwide because of the crop damage they inflict and plant ...

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 ...