One-step CRISPR-based Strategy for Endogenous Gene Tagging in Drosophila melanogaster

1. Design and construction of gene editing reagents

NOTE: To carry out endogenous tagging, it is essential to identify the various isoforms of the desired gene. Depending on the specific goals of the experiment, a fluorescent tag can be incorporated either internally or at the N- or C-termini of the selected protein isoform(s). For optimal CRISPR-mediated editing, it's crucial to position the two sgRNAs suitably close to the intended knock-in site. Subsequently, to enable editing through homologous recombination, a donor vector encompassing sequences homologous to the target gene, along with the fluorescent tag, should be prepared. Below is a step-by-step guide to these processes (Figure 1).

1. Design sgRNA as described below.
   1. Design sgRNAs for Drosophila using these websites: http://tools.flycrispr.molbio.wisc.edu/targetFinder/ or http://www.flyrnai.org/crispr2/. 
   2. For the flycrispr website, enter the sequence ±100 bp surrounding the desired knock-in location(N terminal, C terminal, or any genomic locus of interest). If using the nos-Cas9 attP2 as the injection background, choose the genome option Drosophila melanogaster (nos-Cas9 III, BDSC #78782). The Nos-Cas9 system ensures germline-specific Cas9 expression.
   3. To improve sgRNA expression efficiency, select the CRISPR targets with 5' G option, as sgRNAs that commence with a guanine will enhance editing activity. Then, click the Find CRISPR Targets button. A list of possible sgRNA sequences will be displayed.
   4. For each sequence, click the Evaluate button. This action checks for any potential off targets for each sequence based on the previously selected genome. From the provided list, choose 2 sgRNAs that have 0 off targets. Ensure one sgRNA is upstream and the other is downstream relative to the knock-in location. Ideally, their positions should be as near to the knock-in site as possible (<200 bp), but they must not overlap with one another (Figure 2).
      ​NOTE: It might sometimes be difficult to find appropriate sgRNAs for a given target gene. The gap between sgRNAs and the knock-in location can be increased to ± 200 bp. Using sgRNAs not initiating with G might slightly decrease the positive rate.
2. Perform fluorescent tag selection as described below.
   1. Fluorescent tags offer a diverse range of characteristics, each having their unique advantages and drawbacks^1,2. When determining the ideal fluorescent tag for labeling a specific protein, take care of the key considerations described in the following steps that come into play.
   2. Given that the natural protein expression levels are typically much lower than those seen in overexpression experiments^1,2, start with the tags that are bright and photostable and that fit the experimental setup. Some fluorescent tags, specifically certain RFPs like tdTomato and DsRed, tend to form multimers¹⁴ and may lead to protein aggregation.
   3. For enhanced versatility, link epitope tags (e.g., FLAG or V5) to the fluorescent marker using a flexible linker, which allows for biochemical analyses such as Western Blot or Co-Immunoprecipitation.
3. For linker design, follow the steps described below.
   1. For optimal functionality, ensure that both the endogenous protein of interest and the fluorescent tag fold separately into their respective functional forms. To achieve this, use a linker peptide ranging from 2-20 amino acids in length. For example, place two glycine residues between the last amino acid of the Period protein and the first amino acid of the knock-in fluorescent protein mNeonGreen.
   2. Ensure that these linker peptide sequences comprise amino acids like glycine and serine, which impart flexibility to the structure¹⁵. This ensures that the natural behavior and location of the endogenous protein remain unaltered.
4. Design donor vector as described below.
   ​NOTE: The donor vector introduces the fluorescent tag to the desired endogenous location through the process of homology-directed repair, which is initiated by CRISPR cleavage. As such, besides the fluorescent tag, the donor vector must possess sequences that are homologous on both sides of the tag.
   1. Begin the donor vector design by organizing the DNA sequence of the linker peptide and the fluorescent tag and ensure there are homologous sequences flanking both ends (homology arms). For optimal results, ensure the homology arms encompass at least 800 nucleotides on each side of the intended knock-in site (Figure 2).
      ​NOTE: Recent reports suggest that homology arms can be shorter (100-200 bp)¹⁶.
   2. Ensure that any PAM sequences within the donor plasmid are modified in such a way that the resulting protein's amino acid sequence remains unchanged. If the PAM sequence is located in the UTR, remove it entirely.
   3. The positioning of START and STOP codon depends on the location of the tag. For internal tagging, ensure the fluorescent tag is in frame with the coding sequence of the target gene. For N-terminus tagging, position the fluorescent tag before the endogenous start codon. For C-terminus tagging, situate the fluorescent tag before the natural stop codon and make sure to retain only a single start codon to avoid potential translation issues.
5. Synthesize sgRNA and donor plasmids as described below.
   1. For sgRNA plasmid creation, follow the U6-gRNA cloning guidelines, which can be found at flycrispr.org/protocols/grna/. Employ the alternative restriction enzyme-based cloning approach (referred to as strategy C in the provided resource) to insert gRNA sequence into the pU6-BbsI-chiRNA plasmid (Addgene, #45946).
   2. Transform in DH5a E. coli (NEB, C2987H) following standard transformation protocols provided by the manufacturer. For verification, conduct Sanger sequencing on the resulting clones using T3 or T7 standard sequencing primers.
   3. For the creation of donor plasmid, acquire double-stranded DNA with the desired donor sequence from commercial suppliers (e.g., IDT). Integrate it into the pBlueScriptII SK(-) multiple cloning site. Sequence the donor plasmid to ensure sgRNA or PAM sequences are correctly modified and there are no SNPs present in the plasmid.
      NOTE: An alternative to the integration using pBlueScriptII is to PCR the homology arms and the fluorescent tag segments separately and consolidate them into the backbone using techniques like Golden Gate Assembly or Gibson Assembly.
6. Use the following injection and crossing scheme.
   1. To acquire germline-edited flies, co-inject gRNA plasmids and the donor plasmid into the appropriate Cas9 background (e.g., nos-Cas9 III, BDSC 78782). If utilizing commercial injection services, follow instructions from the vendor to perform plasmid preparation.
      NOTE: Typically, the gRNA plasmid concentration should exceed 0.5 µg/µL, and the donor plasmid concentration should be above 1.0 µg/µL. A common timeline from sending out plasmid samples to receiving injected embryos is approximately 2 weeks. A strategy to screen for gene editing events on the X chromosome employing the FM7/L balancer strain is presented here. For genes on the second or third chromosome, employ the appropriate balancer strains.
   2. For the collection of injected flies (G0), anesthetize with carbon dioxide and separate male flies and virgin females. Establish fly crosses by pairing each G0 male with at least two FM7/L virgin females in individual narrow CT food vials. Place crossed vials in a vented incubator maintained at 25°C and ~60% humidity until F1 flies eclose (FM7/L is the most commonly used X chromosome balancer). Similarly, pair each G0 female with FM7/Y male flies.
   3. Anesthetize and collect at least 10 virgin female offspring (F1) from both G0 male and female crosses. From G0 female crosses, gather at least 10 male F1 flies (Figure 3A). Ensure each F1 fly is tagged with the details of its parentage and stored separately. Proceed to the screening step.
7. Use the following single-leg PCR protocol.
   1. Primer creation: Design a primer pair encompassing the editing site of the target gene. Ideally, the melting temperatures of the primers should be similar, around 55°C.
   2. Lysis buffer preparation: Prepare 10 µL of lysis buffer for every fly sample. The buffer consists of 10 mM Tris (pH 8.2), 1 mM EDTA (pH 8.0), 25 mM NaCl, and 400 µg/mL Proteinase K. Prepare the lysis buffer as a master mix, then distribute 10 µL into each well of a 96-well PCR plate. Ensure the buffer stays cold throughout the leg dissection process (Figure 3B).
   3. Tube setup: Prepare glass capillary tubes, filling one end with sucrose-agar food (2% agar and 4% sucrose), and place these in a 96-well deep-well plate, which will be referred to as the fly hotel (Figure 3C).
   4. Leg dissection: Place the carbon dioxide anesthetization pad under a stereo microscope, anesthetize one candidate fly and sever one of the middle legs. Place the leg into the lysis buffer, ensuring complete immersion. It is not necessary to macerate the leg, directly proceed to the lysis protocol step (Figure 3B). Hold the fly wing with a pair of fine metal forceps, move the fly to the fly hotel, and seal the tube with yarn immediately.
   5. Control samples: Two wild-type flies may be used as negative controls following the same leg dissection procedure. A diluted donor plasmid serves as the positive control.
   6. Post-dissection care: House the fly hotel in a vented incubator maintained at 25 °C and ~60% humidity. While flies can survive overnight, it's recommended to complete subsequent steps and mating within the same day.
   7. Lysis protocol: Place the severed legs in the lysis buffer solution in a thermocycler set at 37 °C for 90 min, followed by 95 °C for 5 min, finally held at 4 °C. This will extract sufficient DNA from the single leg of each fly (Figure 3B).
   8. PCR reaction: Organize the PCR reaction as instructed by the manufacturer, utilizing 1.5 µL of the fly leg lysate as the DNA source.
   9. Gel electrophoresis: Subject the PCR products to agarose gel electrophoresis. Interpret the results from the gel based on band size. Typically, negative control flies will exhibit a lower band, and positive plasmid controls display a higher band. Ensure specific band sizes are consistent with the original donor plasmid design. Female positive flies will display two bands as they are heterozygous, and male positives will present a single band (Figure 3B).
      ​NOTE: Usually, screening of 70 flies is achievable in a single day. Ensure to identify a minimum of 10 positive F1 flies across 4 to 5 distinct G0 crosses, as certain F1s might be sterile due to off-target mutations.
8. Carry out the following post-PCR cross set-up.
   1. Based on the gel results, select the positive flies from the fly hotel and individually cross them with balancer flies (FM7/L for tagging X chromosome genes) to produce the F2 generation. Once the F2 generation emerges, establish sibling crosses to create homozygous stable stocks (Figure 3D).
   2. Use homozygous stocks for Sanger sequencing of the editing site to ensure accuracy. Backcross validated fly lines with wild-type flies to remove background mutations. Screen each generation using the single-leg PCR protocol.