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

Summary

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.

Abstract

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 φC31 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 φC31 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 φC31 integrase. We report two representative results of φC31-mediated site-directed modification: the single integration of a transgenic cassette in An. stephensi and RMCE in An. gambiae mosquitoes. φC31-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 φC31 system an essential component of the genetic toolkit for the transgenic manipulation of malaria mosquitoes and other insect vectors.

Introduction

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 Anopheles^2,3,4. However, the random nature of TE integration can lead to undesirable modifications such as gene knockouts (insertional mutagenesis) and significant position effects on transgene expression^5,6,7,8. Multiple insertions are also a common occurrence when using piggyBac^5,9, 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 transposase^10,11,12, and their limited size of DNA cargo (10-15 kb in length) with transformation efficiency declining with increasing size of the donor plasmid^13,14.

Site-directed integration approaches were introduced to circumvent these issues. The most common site-directed genome modification in mosquitoes is that mediated by the φC31 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 φC31 (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 factor^16,17. 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 >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 φC31-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 φC31-mediated single integration have been developed and validated in An. gambiae^19,20,21,22 and in An. stephensi^23,24,25 (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.

φC31-mediated integration has been applied extensively to insect genome editing from the model organism D. melanogaster²⁷, to the mosquitoes Aedes aegypti^13,28, Ae. albopictus²⁹, An. gambiae¹⁹, and An. stephensi²⁴, as well as other insects including Ceratitis capitata³⁰ and Bombyx mori³¹.

A limitation of φC31-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 φC31 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. gambiae^9,26,33, Ae. aegypti³⁴, Plutella xylostella³⁴, and B. mori³⁵. Several docking lines for RMCE have been developed and validated in An. gambiae^5,9,26 (Table 1). To our knowledge, RMCE is yet to be explored in other Anopheles vectors species.

To date, the φC31 system has been used widely in Anopheles mosquitoes to introduce and study a variety of molecules including antimalaria effectors^19,24,36, components of the GAL4/UAS system to overexpress and knockdown genes for insecticide resistance studies^9,33, regulatory elements, reporter genes^5,21,37, and gene-drive elements^26,38.

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 φC31 integrase. The major malaria vectors An. gambiae and An. stephensi are used as specific examples, however these protocols are applicable to other Anopheles species.

Figure 1. Site-directed genome modifications, single integration and recombinase-mediated cassette exchange (RMCE) , using the φC31 system. The φC31 integrase (INT, grey double arrow) catalyzes the recombination between the attB site(s) (purple striped) present in a donor plasmid and the attP site(s) (blue striped) present in a receiving docking line, which results in the formation of hybrid sites attL and attR. A) Integration is achieved when single attB and attP sites recombine and results in the presence of two integrated markers (blue and red). B) RMCE occurs when two attB/P sites recombine simultaneously and results in the replacement of the cassette between the att sites of the docking line (blue marker) with that carried by the donor plasmid (red marker). C) Partial nucleotide sequences of attP (blue) and attB (purple) and the hybrid sites attL/R. Recombination occurs between the 'TT' core sequences highlighted in bold black. Please click here to view a larger version of this figure.

Protocol

NOTE: A schematic workflow of the illustrated protocol is shown in Figure 2.

1. Design of φC31 attB -tagged plasmids (Figure 3)

1. Create attB donor plasmids carrying the following essential components
   1. Dominant fluorescent marker
      1. Choose a promoter to drive the expression of the fluorescent marker.
         ​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.
      2. Choose a fluorescent protein (FP) that is compatible with that of the receiving docking line so that they are readily distinguishable.
         ​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.
   2. attB recombination site(s)
      1. Use a single attB site for integration of a transgenic cassette (single-attB design) (Figure 3A).
      2. 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.
   3. Desired transgene cargo
      1. 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.
   4. Plasmid backbone components
      1. 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).

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 φC31 integrase under the regulation of the Drosophila Hsp70 promoter⁴⁰.

1. 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.
2. Combine appropriate amounts of the two plasmids to obtain a mix with a final concentration of 350 ng/µL of the donor plasmid and 150 ng/µL of the helper plasmid when resuspended in injection buffer.
   NOTE: When calculating the necessary volume of mix, consider that 10-15 µL are sufficient for each day of planned injections and DNA can be prepared in advance and stored at -20 °C. Integrase helper plasmid concentrations of 60-500 ng/µL and donor plasmid concentrations of 85-200 ng/µL have also been reported^21,22,26,41.
3. 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 µg/µL) improves precipitation efficiency.
   NOTE: Stopping point - The precipitate can be stored at -20 °C overnight.
4. Centrifuge at 15,000 x g for 20 min at 4 °C, discard the supernatant, and wash the pellet with 1 mL of ice-cold 70% EtOH.
5. Wash the pellet with 1 mL of ice-cold 70% EtOH and centrifuge at 15,000 x g for 5 min at room temperature.
6. Discard the supernatant without disturbing the pellet and air dry.
7. Resuspend the pellet in 1x injection buffer (0.1 mM Na₃PO₄, 5 mM KCl, pH 7.2, 0.22 µm filter sterilized) to reach a total final concentration of 500 ng/µL.
   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/µL.
8. Ensure that the DNA is thoroughly resuspended, prepare aliquots of 10-15 µL each and store them at -20 °C.
9. On the day of injection, thaw one aliquot and centrifuge at 15,000 x g for 5 min to remove any particulate residues.
   ​NOTE: An alternative method for particulate removal is to filter the solution through a 0.22 µm filter. Avoid the presence of particulate residues in the injection mix as they lead to needle blockage during embryo microinjection.

3. Microinjection of embryos from an Anopheles docking line

1. 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).
2. 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.
3. Perform embryo microinjections
   1. 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.
   2. 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.
   3. Transfer eggs immediately after injection in a Petri dish filled with sterile distilled water (pH 7.2) and return them to insectary conditions.
4. Upon hatching, transfer G₀ larvae into a tray with salted distilled water (0.1% tonic salt) daily and rear to pupae.
5. Record hatching rate (i.e., number of larvae hatched/number of embryos injected).
   ​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.

4. Crossing and screening of transformed individuals

1. [OPTIONAL STEP] Screen G₀ (injected) 1^st or 2^nd instar larvae (L₁-L₂) for transient expression of the fluorescent marker.
   1. Use a fine-tip glass pipette to transfer G₀ L₁-L₂ larvae to a microscope slides with wells. Place one larva in each well.
   2. 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)
   3. Rear G₀ positive individuals separately.
2. Sort G₀ pupae by sex under a stereoscope⁵².
3. 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.
4. 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.
5. Allow adults to mate for 4-5 days and provide females with a blood meal.
   ​NOTE: Blood feed and collect eggs from G₀ females multiple times to maximize the chances of getting transformants from multiple gonotrophic cycles.
6. Blood feed WT individuals at the same time for outcrossing.
7. Collect eggs and rear emerging next generation G₁s.
8. Screen G₁ L₃-L₄ larvae for appropriate fluorescence to identify transformants.
   1. 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.
      ​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.
      1. 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, Figure 4).
         ​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.
      2. 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).
         ​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.
9. Transfer transformed G₁ individuals into a larval tray and rear to pupae. Discard non-fluorescent individuals and individuals with an unexpected marker expression pattern.
10. Sort transformed G₁ pupae by sex and cross them en masse with opposite-sex age-matched WT individuals.
11. Allow adults to mate for 4-5 days, provide a blood meal, collect the eggs, and rear the next generation G₂ progeny.
    1. For single integration experiments, collect eggs directly from the en masse cross as the integration site is identical in all individuals.
    2. 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).
12. 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.
13. 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'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.
14. Allow adult males and females to intercross in the same cage to establish the new transgenic line.
    ​NOTE: For RMCE experiments, adult intercross must occur between siblings deriving from a single female until orientation of insertion is determined via molecular analysis.

5. Molecular validation of the insertion site by DNA amplification (PCR)

1. Prepare a map of the predicted insertion site in the genome of the docking line after transformation.
   1. 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).
   2. 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).
2. Design oligonucleotide primers to amplify the insertional junction at either side of the integration locus.
   1. 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).
   2. 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).
3. 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's legs⁴⁶ or pupal cases (L. Grigoraki personal communication).
4. Sequence PCR products to confirm expected sequences.

Figure 2. Workflow diagram for site-directed φC31 genome modification in Anopheles mosquitoes. Please click here to view a larger version of this figure.

Figure 3. Molecular basis of φC31-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. Please click here to view a larger version of this figure.

Discussion

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 φC31 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 promoter^22,26. Other protocols describe the use of in vitro transcribed messenger RNA (mRNA) as source of φC31 integrase^19,24,43. 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. gambiae^{9,21,22,26,33,37} 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 φC31 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 ones^43,48. 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 site^9,33. 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'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 editing^26,41,50,51. 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 φC31 system remains a very valuable tool to perform direct phenotypic comparisons between transgenes at the same genomic locations.

Materials

Name	Company	Catalog Number	Comments
1.5 mL eppendorf tubes
8-well microslides	VWR	MARI1216690
DNeasy Blood & Tissue Kit	Qiagen	69504
EndoFree Plasmid Maxi Kit (10)	Qiagen	12362
Ethanol, Absolute, Molecular Biology Grade
Filter set CFP for Leica MZ FLIII Excitation 436/20 nm, extinction 480/40 nm	Leica	10446363
Filter set dsRED for Leica MZ FLIII Excitation 545/30 nm, extinction 620/60 nm	Leica	10447079
Filter set YFP customised for Leica MZ FLIII	Omega Optical	500QM25, 500QM35
Halocarbon oil 27	Sigma	H8773
Halocarbon oil 700	Sigma	H8898
Petri dishes
Potassium chloride
Sodium Chloride
Sodium phosphate dibasic
Sodium Acetate Solution (3 M), pH 5.2	Thermo Fisher Scientific (Life Technologies)	R1181
Stable brush Size 0