We thank George Watase and Josie Clowney for discussions during the initial stages of the protocol development. The work was supported by funds from the NIH (grant no. R35GM133737 to S.Y.), Alfred P. Sloan Fellowship (to S. Y.) and McKnight Scholar Award (to S. Y.).

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&#39;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).
<ol>
	<li>Design sgRNA as described below.
	<ol>
		<li>Design sgRNAs for Drosophila using these websites:&#160;http://tools.flycrispr.molbio.wisc.edu/targetFinder/ or http://www.flyrnai.org/crispr2/.&#160;</li>
		<li>For the flycrispr website, enter the sequence &#177;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.</li>
		<li>To improve sgRNA expression efficiency, select the CRISPR targets with 5&#39; 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.</li>
		<li>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 (&#60;200 bp), but they must not overlap with one another (Figure 2). 
		&#8203;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 &#177; 200 bp. Using sgRNAs not initiating with G might slightly decrease the positive rate.</li>
	</ol>
	</li>
	<li>Perform fluorescent tag selection as described below.
	<ol>
		<li>Fluorescent tags offer a diverse range of characteristics, each having their unique advantages and drawbacks1,2. 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.</li>
		<li>Given that the natural protein expression levels are typically much lower than those seen in overexpression experiments1,2, 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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>For linker design, follow the steps described below.
	<ol>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Design donor vector as described below. 
	&#8203;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.
	<ol>
		<li>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). 
		&#8203;NOTE: Recent reports suggest that homology arms can be shorter (100-200 bp)16.</li>
		<li>Ensure that any PAM sequences within the donor plasmid are modified in such a way that the resulting protein&#39;s amino acid sequence remains unchanged. If the PAM sequence is located in the UTR, remove it entirely.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Synthesize sgRNA and donor plasmids as described below.
	<ol>
		<li>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).</li>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Use the following injection and crossing scheme.
	<ol>
		<li>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 &#181;g/&#181;L, and the donor plasmid concentration should be above 1.0 &#181;g/&#181;L. 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.</li>
		<li>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&#176;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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Use the following single-leg PCR protocol.
	<ol>
		<li>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&#176;C.</li>
		<li>Lysis buffer preparation: Prepare 10 &#181;L 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 &#181;g/mL Proteinase K. Prepare the lysis buffer as a master mix, then distribute 10 &#181;L into each well of a 96-well PCR plate. Ensure the buffer stays cold throughout the leg dissection process (Figure 3B).</li>
		<li>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).</li>
		<li>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.</li>
		<li>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.</li>
		<li>Post-dissection care: House the fly hotel in a vented incubator maintained at 25 &#176;C and ~60% humidity. While flies can survive overnight, it&#39;s recommended to complete subsequent steps and mating within the same day.</li>
		<li>Lysis protocol: Place the severed legs in the lysis buffer solution in a thermocycler set at 37 &#176;C for 90 min, followed by 95 &#176;C for 5 min, finally held at 4 &#176;C. This will extract sufficient DNA from the single leg of each fly (Figure 3B).</li>
		<li>PCR reaction: Organize the PCR reaction as instructed by the manufacturer, utilizing 1.5 &#181;L of the fly leg lysate as the DNA source.</li>
		<li>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). 
		&#8203;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.</li>
	</ol>
	</li>
	<li>Carry out the following post-PCR cross set-up.
	<ol>
		<li>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).</li>
		<li>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.</li>
	</ol>
	</li>
</ol>

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The authors declare no competing interests.

The results demonstrate a streamlined, simple one-step CRISPR-based strategy for tagging endogenous genes in Drosophila. In the protocol, we use two gRNAs to induce DNA double-strand break and homologous recombination more efficiently. The gRNAs and the donor plasmid are injected into fruit fly embryos, which express Cas9 enzyme in their germline. These embryos are subsequently cultivated under standard conditions until adulthood, at which point they are crossed with balancer flies to produce the first generatio...

Fluorescent proteins have emerged as powerful tools for visualizing protein localization, dynamics, and protein-protein interactions in cells within living organisms1,2. Tagging endogenous genes with fluorescent proteins enables long-term live imaging of cellular processes with subcellular resolution. Furthermore, these endogenous gene-labeling techniques have several advantages compared to overexpression and immunostaining approaches. For example, overexpression of proteins might lead to protein misfolding, nonspecific protein-protein interactions, and mislocalization3. To date, researchers have been able to create vast libraries of endogenous genes fused to fluorescent proteins for a few model organisms, such as budding yeast, which can undergo efficient homologous recombination4. In the case of Drosophila melanogaster, various tools such as MiMICs5, transposon-based enhancer traps6, and fosmids7 have been devised to fluorescently tag genes.
The development of CRISPR-Cas9-based tools has made it possible to efficiently perform genome editing, including tagging endogenous proteins, in a wide range of model organism8,9. Cas9 is an RNA-guided enzyme that cleaves double-stranded DNA in a highly efficient and specific manner8, which can then be repaired by a nonhomologous end-joining (NHEJ) pathway leading to point mutations or deletions. Alternatively, the homology-directed repair (HDR) pathway allows the integration of heterologous DNA into the chromosome.
Past methods for CRISPR-based gene tagging in the model organism, Drosophila melanogaster, have relied on a two-step process10,11,12. This involves using a discernible eye-color marker, Dsred, to detect successful HDR events. Afterward, the Dsred marker would be excised using the Cre/loxP recombination system. To streamline and expedite this process, here, we present a simpler and more efficient method. This approach circumvents the need for lethal genotyping and allows for the direct screening of individual flies. Specifically, we employ a PCR-based approach to extract DNA from a small segment of the fly&#39;s middle leg, allowing us to genotype individual flies. Notably, this procedure does not compromise the vitality, movement, or reproductive capabilities of the sampled fly. Only the appropriate individuals, identified through this method, are further developed into stable stocks.
Here, we present a detailed protocol for generating fluorescent protein tagged flies in which endogenous genes are labeled with fluorescent reporters. This technique uses CRISPR-Cas9 genome editing to fuse any desired fluorophore tag to the N- or C- terminal of an endogenous gene. Below, we describe the design and construction of gRNA plasmids and a donor plasmid, one-step screening strategy to select CRISPR-positive flies, and steps to establish stable lines. Specifically, we describe strategies to tag an endogenous core clock Drosophila gene, period, with a fluorophore at the C-terminal. We also briefly describe the control experiments that need to be performed to confirm that the tagged protein is functional and live-imaging techniques to visualize the Period protein in live clock neurons within intact Drosophila brains13. The protocol described here is also broadly applicable to screen candidates for deletions and insertions of specific bits of DNA by CRISPR-Cas9 genome editing.

<table><tbody><tr><td>0.5M EDTA pH8.0</td><td>Invitrogen</td><td>AM9260G</td><td></td></tr><tr><td>5-alpha Competent&amp;nbsp;E. coli&amp;nbsp;(High Efficiency)</td><td>NEB</td><td>C2987H</td><td></td></tr><tr><td>96-well deep well plate</td><td>BRAND</td><td>701354</td><td></td></tr><tr><td>Agar</td><td>Fisher Scientific</td><td>BP1423-500</td><td></td></tr><tr><td>BbsI-HF</td><td>NEB</td><td>R3539S</td><td></td></tr><tr><td>DreamTaq Green PCR Master Mix (2X)</td><td>Thermo Scientific</td><td>K1081</td><td></td></tr><tr><td>D-Sucrose</td><td>Fisher Scientific</td><td>BP220-1</td><td></td></tr><tr><td>EcoRI-HF</td><td>NEB</td><td>R3101S</td><td></td></tr><tr><td>Hydrochloric Acid</td><td>Fisher Scientific</td><td>A142-212</td><td></td></tr><tr><td>Prolong Glass Antifade Mounting Medium</td><td>Invitrogen</td><td>P36982</td><td></td></tr><tr><td>Proteinase K</td><td>QIAGEN</td><td>19131</td><td></td></tr><tr><td>Schneider&#039;s &lt;em&gt;Drosophila&lt;/em&gt; Medium</td><td>Gibco</td><td>21720001</td><td></td></tr><tr><td>Sodium Chloride</td><td>Fisher Scientific</td><td>S271-500</td><td></td></tr><tr><td>T4 DNA ligase</td><td>NEB</td><td>M0202S</td><td></td></tr><tr><td>Tris Base</td><td>Fisher Scientific</td><td>BP152-500</td><td></td></tr><tr><td>XbaI</td><td>NEB</td><td>R0145S</td><td></td></tr></tbody></table>

one-step crispr-based strategy for endogenous gene tagging in drosophila melanogaster

The study of protein subcellular localization, dynamics, and regulation in live cells has been profoundly transformed by the advent of techniques that allow the tagging of endogenous genes to produce fluorescent fusion proteins. These methods enable researchers to visualize protein behavior in real time, providing valuable insights into their functions and interactions within the cellular environment. Many current gene tagging studies employ a two-step process where visible markers, such as eye color changes, are used to identify genetically modified organisms in the first step, and the visible marker is excised in the second step. Here, we present a one-step protocol to perform precise and rapid endogenous gene tagging in Drosophila melanogaster, which enables screening for engineered lines without the visible eye marker, offering a significant advantage over past methods. To screen for successful gene-tagging events, we employ a PCR-based technique to genotype individual flies by analyzing a small segment from their middle leg. Flies that pass the screening criteria are then used to produce stable stocks. Here, we detail the design and construction of CRISPR editing plasmids and methods for screening and confirmation of engineered lines. Together, this protocol improves the efficiency of endogenous gene tagging in Drosophila significantly and enables studies of cellular processes in vivo.

In our experiments, we typically found a success rate of ~20%-30% in screening for various endogenous tagged genes on all chromosomes (X, II, III). The efficiency of this process can vary based on several factors, such as gRNA selection, the nature of the gene, and injection quality. Here, we illustrate the results from our strategy to tag the endogenous period gene with the mNeonGreen fluorophore13. The period gene, positioned on the X chromosome...

Watch this Scientific Journal Video about One-step CRISPR-based Strategy for Endogenous Gene Tagging in Drosophila melanogaster at JoVE.com

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

Here, we present a simplified endogenous gene tagging protocol for Drosophila, which utilizes a PCR-based technique for marker-free identification of successful genetic modifications, facilitating the development of stable knock-in lines.

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

The study of protein subcellular localization, dynamics, and regulation in live cells has been profoundly transformed by the advent of techniques that ...

one-step-crispr-based-strategy-for-endogenous-gene-tagging-drosophila

Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

615 Views.  University of Michigan. Here, we present a simplified endogenous gene tagging protocol for Drosophila, which utilizes a PCR-based technique for marker-free identification of successful genetic modifications, facilitating the development of stable knock-in lines.

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

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

Characterization of the clustered, regularly interspaced, short, palindromic repeat (CRISPR) system of Streptococcus pyogenes has enabled the development of a customizable platform to rapidly generate gene modifications in a wide variety of organisms, including zebrafish. CRISPR-based genome editing uses a single guide RNA (sgRNA) to target a CRISPR-associated (Cas) endonuclease to a genomic DNA (gDNA) target of interest, where the Cas endonuclease generates a double-strand break (DSB). Repair of DSBs by error-prone mechanisms lead to insertions and/or deletions (indels). This can cause frameshift mutations that often introduce a premature stop codon within the coding sequence, thus creating a protein-null allele. CRISPR-based genome engineering requires only a few molecular components and is easily introduced into zebrafish embryos by microinjection. This protocol describes the methods used to generate CRISPR reagents for zebrafish microinjection and to identify fish exhibiting germline transmission of CRISPR-modified genes. These methods include in vitro transcription of sgRNAs, microinjection of CRISPR reagents, identification of indels induced at the target site using a PCR-based method called a heteroduplex mobility assay (HMA), and characterization of the indels using both a low throughput and a powerful next-generation sequencing (NGS)-based approach that can analyze multiple PCR products collected from heterozygous fish. This protocol is streamlined to minimize both the number of fish required and the types of equipment needed to perform the analyses. Furthermore, this protocol is designed to be amenable for use by laboratory personal of all levels of experience including undergraduates, enabling this powerful tool to be economically employed by any research group interested in performing CRISPR-based genomic modification in zebrafish.

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

Targeted genome editing in the model system Danio rerio (zebrafish) has been greatly facilitated by the emergence of CRISPR-based approaches. Herein, we describe a streamlined, robust protocol for generation and identification of CRISPR-derived nonsense alleles that incorporates the heteroduplex mobility assay and identification of mutations using next-generation sequencing.

Characterization of the clustered, regularly interspaced, short, palindromic repeat (CRISPR) system of Streptococcus pyogenes has enabled the development ...

Genetics

A Rapid and Facile Pipeline for Generating Genomic Point Mutants in C. elegans Using CRISPR/Cas9 Ribonucleoproteins

The clustered regularly interspersed palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9) prokaryotic adaptive immune defense system has been co-opted as a powerful tool for precise eukaryotic genome engineering. Here, we present a rapid and simple method using chimeric single guide RNAs (sgRNA) and CRISPR-Cas9 Ribonucleoproteins (RNPs) for the efficient and precise generation of genomic point mutations in C. elegans. We describe a pipeline for sgRNA target selection, homology-directed repair (HDR) template design, CRISPR-Cas9-RNP complexing and delivery, and a genotyping strategy that enables the robust and rapid identification of correctly edited animals. Our approach not only permits the facile generation and identification of desired genomic point mutant animals, but also facilitates the detection of other complex indel alleles in approximately 4 - 5 days with high efficiency and a reduced screening workload.

A Rapid and Facile Pipeline for Generating Genomic Point Mutants in C. elegans Using CRISPR/Cas9 Ribonucleoproteins

Here, we present a method to engineer the genome of C. elegans using CRISPR-Cas9 ribonucleoproteins and homology dependent repair templates.

The clustered regularly interspersed palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9) prokaryotic adaptive immune defense system has been ...

Endogenous Protein Tagging in Human Induced Pluripotent Stem Cells Using CRISPR/Cas9

A protocol is presented for generating human induced pluripotent stem cells (hiPSCs) that express endogenous proteins fused to in-frame&#160;N- or C-terminal fluorescent tags. The prokaryotic CRISPR/Cas9 system (clustered regularly interspaced short palindromic repeats/CRISPR-associated 9) may be used to introduce large exogenous sequences into genomic loci via homology directed repair (HDR). To achieve the desired knock-in, this protocol employs the ribonucleoprotein (RNP)-based approach where wild type Streptococcus pyogenes Cas9 protein, synthetic 2-part guide RNA (gRNA), and a donor template plasmid are delivered to the cells via electroporation. Putatively edited cells expressing the fluorescently tagged proteins are enriched by fluorescence activated cell sorting (FACS). Clonal lines are then generated and can be analyzed for precise editing outcomes. By introducing the fluorescent tag at the genomic locus of the gene of interest, the resulting subcellular localization and dynamics of the fusion protein can be studied under endogenous regulatory control, a key improvement over conventional overexpression systems. The use of hiPSCs as a model system for gene tagging provides the opportunity to study the tagged proteins in diploid, nontransformed cells. Since hiPSCs can be differentiated into multiple cell types, this approach provides the opportunity to create and study tagged proteins in a variety of isogenic cellular contexts.

Described here is a protocol for tagging endogenously expressed proteins with fluorescent tags in human induced pluripotent stem cells using CRISPR/Cas9. Putatively edited cells are enriched by fluorescence activated cell sorting and clonal cell lines are generated.

A protocol is presented for generating human induced pluripotent stem cells (hiPSCs) that express endogenous proteins fused to in-frame&#160;N- or ...

In Vivo Functional Study of Disease-associated Rare Human Variants Using Drosophila

Advances in sequencing technology have made whole-genome and whole-exome datasets more accessible for both clinical diagnosis and cutting-edge human genetics research. Although a number of in silico algorithms have been developed to predict the pathogenicity of variants identified in these datasets, functional studies are critical to determining how specific genomic variants affect protein function, especially for missense variants. In the Undiagnosed Diseases Network (UDN) and other rare disease research consortia, model organisms (MO) including Drosophila, C. elegans, zebrafish, and mice are actively used to assess the function of putative human disease-causing variants. This protocol describes a method for the functional assessment of rare human variants used in the Model Organisms Screening Center Drosophila Core of the UDN. The workflow begins with gathering human and MO information from multiple public databases, using the MARRVEL web resource to assess whether the variant is likely to contribute to a patient&#39;s condition as well as design effective experiments based on available knowledge and resources. Next, genetic tools (e.g., T2A-GAL4 and UAS-human cDNA lines) are generated to assess the functions of variants of interest in Drosophila. Upon development of these reagents, two-pronged functional assays based on rescue and overexpression experiments can be performed to assess variant function. In the rescue branch, the endogenous fly genes are &#34;humanized&#34;&#160;by replacing the orthologous Drosophila gene with reference or variant human transgenes. In the overexpression branch, the reference and variant human proteins are exogenously driven in a variety of tissues. In both cases, any scorable phenotype (e.g., lethality, eye morphology, electrophysiology) can be used as a read-out, irrespective of the disease of interest. Differences observed between reference and variant alleles suggest a variant-specific effect, and thus likely pathogenicity. This protocol allows rapid, in vivo assessments of putative human disease-causing variants of genes with known and unknown functions.

In Vivo Functional Study of Disease-associated Rare Human Variants Using Drosophila

The goal of this protocol is to outline the design and performance of in vivo experiments in Drosophila melanogaster to assess the functional consequences of rare gene variants associated with human diseases.

Advances in sequencing technology have made whole-genome and whole-exome datasets more accessible for both clinical diagnosis and cutting-edge human ...

An Efficient Strategy for Generating Tissue-specific Binary Transcription Systems in Drosophila by Genome Editing

Binary transcription systems are powerful genetic tools widely used for visualizing and manipulating cell fate and gene expression in specific groups of cells or tissues in model organisms. These systems contain two components as separate transgenic lines. A driver line expresses a transcriptional activator under the control of tissue-specific promoters/enhancers, and a reporter/effector line harbors a target gene placed downstream to the binding site of the transcription activator. Animals harboring both components induce tissue-specific transactivation of a target gene expression. Precise spatiotemporal expression of the gene in targeted tissues is critical for unbiased interpretation of cell/gene activity. Therefore, developing a method for generating exclusive cell/tissue-specific driver lines is essential. Here we present a method to generate highly tissue-specific targeted expression system by employing a &#34;Clustered Regularly Interspaced Short Palindromic Repeat/CRISPR-associated&#34; (CRISPR/Cas)-based genome editing technique. In this method, the endonuclease Cas9 is targeted by two chimeric guide RNAs (gRNA) to specific sites in the first coding exon of a gene in the Drosophila genome to create double-strand breaks (DSB). Subsequently, using an exogenous donor plasmid containing the transactivator sequence, the cell-autonomous repair machinery enables homology-directed repair (HDR) of the DSB, resulting in precise deletion and replacement of the exon with the transactivator sequence. The knocked-in transactivator is expressed exclusively in cells where the cis-regulatory elements of the replaced gene are functional. The detailed step-by-step protocol presented here for generating a binary transcriptional driver expressed in Drosophila fgf/branchless-producing epithelial/neuronal cells can be adopted for any gene- or tissue-specific expression.

An Efficient Strategy for Generating Tissue-specific Binary Transcription Systems in Drosophila by Genome Editing

Here, we present a method for generating tissue-specific binary transcription systems in Drosophila by replacing the first coding exon of genes with transcription drivers. The CRISPR/Cas9-based method places a transactivator sequence under the endogenous regulation of a replaced gene, and consequently facilitates transctivator expression exclusively in gene-specific spatiotemporal patterns.

Binary transcription systems are powerful genetic tools widely used for visualizing and manipulating cell fate and gene expression in specific groups of ...

Developmental Biology

phiC31 Integrase-Mediated Embryo Transgenesis in Drosophila

Embryo Microinjection for Transgenesis in Drosophila

Transgenesis in Drosophila is an essential approach to studying gene function at the organism level. Embryo microinjection is a crucial step for the construction of transgenic flies. Microinjection requires some types of equipment, including a microinjector, a micromanipulator, an inverted microscope, and a stereo microscope.&#160;Plasmids isolated with a plasmid miniprep kit are qualified for microinjection. Embryos at the pre-blastoderm or syncytial blastoderm stage, where nuclei share a common cytoplasm, are subjected to microinjection. A cell strainer eases the process of dechorionating embryos. The optimal time for dechorionation and desiccation of embryos needs to be determined experimentally. To increase the efficiency of embryo microinjection, needles prepared by a puller need to be beveled by a needle grinder. In the process of grinding needles, we utilize a foot air pump with a pressure gauge to avoid the capillary effect of the needle tip. We routinely inject 120-140 embryos for each plasmid and obtain at least one transgenic line for around 85% of plasmids. This article takes the phiC31 integrase-mediated transgenesis in Drosophila as an example and&#160;presents a detailed protocol for embryo microinjection for transgenesis in Drosophila.

Author Spotlight: Optimizing Embryo Microinjection for Transgenesis in Drosophila

This article takes the phiC31 integrase-mediated transgenesis in Drosophila as an example and presents an optimized protocol for embryo microinjection,&#160;a crucial step for creating transgenic flies.

Transgenesis in Drosophila is an essential approach to studying gene function at the organism level. Embryo microinjection is a crucial step for the ...

A Single-fly Assay for Foraging Behavior in Drosophila

For many animals, hunger promotes changes in the olfactory system in a manner that facilitates the search for appropriate food sources. In this video article, we describe an automated assay to measure the effect of hunger or satiety on olfactory dependent food search behavior in the adult fruit fly Drosophila melanogaster. In a light-tight box illuminated by red light that is invisible to fruit flies, a camera linked to custom data acquisition software monitors the position of six flies simultaneously. Each fly is confined to walk in individual arenas containing a food odor at the center. The testing arenas rest on a porous floor that functions to prevent odor accumulation. Latency to locate the odor source, a metric that reflects olfactory sensitivity under different physiological states, is determined by software analysis. Here, we discuss the critical mechanics of running this behavioral paradigm and cover specific issues regarding fly loading, odor contamination, assay temperature, data quality, and statistical analysis.

A Single-fly Assay for Foraging Behavior in Drosophila

In this video article, we describe an automated assay to measure the effect of hunger or satiety on olfactory dependent food search behavior in the adult fruit fly Drosophila melanogaster.

For many animals, hunger promotes changes in the olfactory system in a manner that facilitates the search for appropriate food sources. In this video ...

Recombineering Homologous Recombination Constructs in Drosophila

The continued development of techniques for fast, large-scale manipulation of endogenous gene loci will broaden the use of Drosophila melanogaster as a genetic model organism for human-disease related research. Recent years have seen technical advancements like homologous recombination and recombineering. However, generating unequivocal null mutations or tagging endogenous proteins remains a substantial effort for most genes. Here, we describe and demonstrate techniques for using recombineering-based cloning methods to generate vectors that can be used to target and manipulate endogenous loci in vivo. Specifically, we have established a combination of three technologies: (1) BAC transgenesis/recombineering, (2) ends-out homologous recombination and (3) Gateway technology to provide a robust, efficient and flexible method for manipulating endogenous genomic loci. In this protocol, we provide step-by-step details about how to (1) design individual vectors, (2) how to clone large fragments of genomic DNA into the homologous recombination vector using gap repair, and (3) how to replace or tag genes of interest within these vectors using a second round of recombineering. Finally, we will also provide a protocol for how to mobilize these cassettes in vivo to generate a knockout, or a tagged gene via knock-in. These methods can easily be adopted for multiple targets in parallel and provide a means for manipulating the Drosophila genome in a timely and efficient manner.

Recombineering Homologous Recombination Constructs in Drosophila

Homologous recombination techniques greatly advance Drosophila genetics by enabling the creation of molecularly precise mutations. The recent adoption of recombineering allows one to manipulate large pieces of DNA and transform them into Drosophila6. The methods presented here combine these techniques to rapidly generate large homologous recombination vectors.

The continued development of techniques for fast, large-scale manipulation of endogenous gene loci will broaden the use of Drosophila melanogaster as a ...

Biology

Ubiquitous and Tissue-specific RNA Targeting in Drosophila Melanogaster using CRISPR/CasRx

CasRx, a member of the RNA-targeting Cas13 family, is a promising new addition of the CRISPR/Cas technologies in efficient gene transcript reduction with an attractive off-target profile at both cellular and organismal levels. It is recently reported that the CRISPR/CasRx system can be used to achieve ubiquitous and tissue-specific gene transcript reduction in Drosophila melanogaster. This paper details the methods from the recent work, consisting of three parts: 1) ubiquitous&#160;in vivo endogenous RNA targeting using a two-component CasRx system; 2) ubiquitous in vivo exogenous RNA targeting using a three-component CasRx system; and 3) tissue-specific in vivo RNA targeting using a three-component CasRx system. The effects of RNA targeting observed include targeted gene specific phenotypic changes, targeted RNA transcript reduction, and occasional lethality phenotypes associated with high expression of CasRx protein and collateral activity. Overall, these results showed that the CasRx system is capable of target RNA transcript reduction at the organismal level in a programmable and efficient manner, demonstrating that in vivo transcriptome targeting, and engineering is feasible and lays the foundation for future in vivo CRISPR-based RNA targeting technologies.

Ubiquitous and Tissue-specific RNA Targeting in Drosophila Melanogaster using CRISPR/CasRx

This article outlines a detailed protocol for using the RNA-targeting Cas13D enzyme (RfxCas13D) in flies.

CasRx, a member of the RNA-targeting Cas13 family, is a promising new addition of the CRISPR/Cas technologies in efficient gene transcript reduction with ...

Immunology and Infection

Visualizing and Tracking Endogenous mRNAs in Live Drosophila melanogaster Egg Chambers

Fluorescence-based imaging techniques, in combination with developments in light microscopy, have revolutionized how cell biologists conduct live cell imaging studies. Methods for detecting RNAs have expanded greatly since seminal studies linked site-specific mRNA localization to gene expression regulation. Dynamic mRNA processes can now be visualized via approaches that detect mRNAs, coupled with microscopy set-ups that are fast enough to capture the dynamic range of molecular behavior. The molecular beacon technology is a hybridization-based approach capable of direct detection of endogenous transcripts in living cells. Molecular beacons are hairpin-shaped, internally quenched, single-nucleotide discriminating nucleic acid probes, which fluoresce only upon hybridization to a unique target sequence. When coupled with advanced fluorescence microscopy and high-resolution imaging, they enable one to perform spatial and temporal tracking of intracellular movement of mRNAs. Although this technology is the only method capable of detecting endogenous transcripts, cell biologists have not yet fully embraced this technology due to difficulties in designing such probes for live cell imaging. A new software application, PinMol, allows for enhanced and rapid design of probes best suited to efficiently hybridize to mRNA target regions within a living cell. In addition, high-resolution, real-time image acquisition and current, open source image analysis software allow for a refined data output, leading to a finer evaluation of the complexity underlying the dynamic processes involved in the mRNA&#39;s life cycle.
Here we present a comprehensive protocol for designing and delivering molecular beacons into Drosophila melanogaster egg chambers. Direct and highly specific detection and visualization of endogenous maternal mRNAs is performed via spinning disc confocal microscopy. Imaging data is processed and analyzed using object detection and tracking in Icy software to obtain details about the dynamic movement of mRNAs, which are transported and localized to specialized regions within the oocyte.

Visualizing and Tracking Endogenous mRNAs in Live Drosophila melanogaster Egg Chambers

Here, we present a protocol for the visualization, detection, analysis and tracking of endogenous mRNA trafficking in live Drosophila melanogaster egg chamber using molecular beacons, spinning disc confocal microscopy, and open-source analysis software.

Fluorescence-based imaging techniques, in combination with developments in light microscopy, have revolutionized how cell biologists conduct live cell ...

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

Summary

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Chapters in this video

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

A Rapid and Facile Pipeline for Generating Genomic Point Mutants in C. elegans Using CRISPR/Cas9 Ribonucleoproteins

Endogenous Protein Tagging in Human Induced Pluripotent Stem Cells Using CRISPR/Cas9

In Vivo Functional Study of Disease-associated Rare Human Variants Using Drosophila

An Efficient Strategy for Generating Tissue-specific Binary Transcription Systems in Drosophila by Genome Editing

Embryo Microinjection for Transgenesis in Drosophila

A Single-fly Assay for Foraging Behavior in Drosophila

Recombineering Homologous Recombination Constructs in Drosophila

Ubiquitous and Tissue-specific RNA Targeting in Drosophila Melanogaster using CRISPR/CasRx

Visualizing and Tracking Endogenous mRNAs in Live Drosophila melanogaster Egg Chambers