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>

<ol>
	<li>Chudakov, D. M., Matz, M. V., Lukyanov, S., Lukyanov, K. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+proteins+and+their+applications+in+imaging+living+cells+and+tissues.">Fluorescent proteins and their applications in imaging living cells and tissues.</a> Physiol Rev. 90 (3), 1103-1163 (2010).</li><li>Costantini, L. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+palette+of+fluorescent+proteins+optimized+for+diverse+cellular+environments.">A palette of fluorescent proteins optimized for diverse cellular environments.</a> Nat Commun. 6, 7670 (2015).</li><li>Gibson, T. J., Seiler, M., Veitia, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+transience+of+transient+overexpression.">The transience of transient overexpression.</a> Nat Methods. 10 (8), 715-721 (2013).</li><li>Huh, W. K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Global+analysis+of+protein+localization+in+budding+yeast.">Global analysis of protein localization in budding yeast.</a> Nature. 425 (6959), 686-691 (2003).</li><li>Venken, K. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=MiMIC:+a+highly+versatile+transposon+insertion+resource+for+engineering+Drosophila+melanogaster+genes.">MiMIC: a highly versatile transposon insertion resource for engineering Drosophila melanogaster genes.</a> Nat Methods. 8 (9), 737-743 (2011).</li><li>Quinones-Coello, A. T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Exploring+strategies+for+protein+trapping+in+Drosophila.">Exploring strategies for protein trapping in Drosophila.</a> Genetics. 175 (3), 1089-1104 (2007).</li><li>Sarov, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+genome-wide+resource+for+the+analysis+of+protein+localisation+in+Drosophila.">A genome-wide resource for the analysis of protein localisation in Drosophila.</a> Elife. 5, e12068 (2016).</li><li>Ran, F. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genome+engineering+using+the+CRISPR-Cas9+system.">Genome engineering using the CRISPR-Cas9 system.</a> Nat Protoc. 8, 2281-2308 (2013).</li><li>Port, F., Chen, H. M., Lee, T., Bullock, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optimized+CRISPR/Cas+tools+for+efficient+germline+and+somatic+genome+engineering+in+Drosophila.">Optimized CRISPR/Cas tools for efficient germline and somatic genome engineering in Drosophila.</a> Proc Natl Acad Sci U S A. 111 (29), E2967-E2976 (2014).</li><li>Gratz, S. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Highly+specific+and+efficient+CRISPR/Cas9-catalyzed+homology-directed+repair+in+Drosophila.">Highly specific and efficient CRISPR/Cas9-catalyzed homology-directed repair in Drosophila.</a> Genetics. 196 (4), 961-971 (2014).</li><li>Kanca, O., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+efficient+CRISPR-based+strategy+to+insert+small+and+large+fragments+of+DNA+using+short+homology+arms.">An efficient CRISPR-based strategy to insert small and large fragments of DNA using short homology arms.</a> Elife. 8, e51539 (2019).</li><li>Lamb, A. M., Walker, E. A., Wittkopp, P. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tools+and+strategies+for+scarless+allele+replacement+in+Drosophila+using+CRISPR/Cas9.">Tools and strategies for scarless allele replacement in Drosophila using CRISPR/Cas9.</a> Fly (Austin). 11 (1), 53-64 (2017).</li><li>Xiao, Y., Yuan, Y., Jimenez, M., Soni, N., Yadlapalli, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Clock+proteins+regulate+spatiotemporal+organization+of+clock+genes+to+control+circadian+rhythms.">Clock proteins regulate spatiotemporal organization of clock genes to control circadian rhythms.</a> Proc Natl Acad Sci U S A. 118 (28), e2019756118 (2021).</li><li>Costantini, L. M., Fossati, M., Francolini, M., Snapp, E. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Assessing+the+tendency+of+fluorescent+proteins+to+oligomerize+under+physiologic+conditions.">Assessing the tendency of fluorescent proteins to oligomerize under physiologic conditions.</a> Traffic. 13 (5), 643-649 (2012).</li><li>Snapp, E. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+proteins:+a+cell+biologist's+user+guide.">Fluorescent proteins: a cell biologist's user guide.</a> Trends Cell Biol. 19 (11), 649-655 (2009).</li><li>Kanca, O., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+expanded+toolkit+for+Drosophila+gene+tagging+using+synthesized+homology+donor+constructs+for+CRISPR-mediated+homologous+recombination.">An expanded toolkit for Drosophila gene tagging using synthesized homology donor constructs for CRISPR-mediated homologous recombination.</a> Elife. 11, e76077 (2022).</li><li>Nitabach, M. N., Taghert, P. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organization+of+the+Drosophila+circadian+control+circuit.">Organization of the Drosophila circadian control circuit.</a> Curr Biol. 18 (2), R84-R93 (2008).</li><li>Carvalho, G. B., Ja, W. W., Benzer, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Non-lethal+PCR+genotyping+of+single+Drosophila.">Non-lethal PCR genotyping of single Drosophila.</a> Biotechniques. 46 (4), 312-314 (2009).</li><li>Gummadova, J. O., Coutts, G. A., Glossop, N. R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+the+Drosophila+Clock+promoter+reveals+heterogeneity+in+expression+between+subgroups+of+central+oscillator+cells+and+identifies+a+novel+enhancer+region.">Analysis of the Drosophila Clock promoter reveals heterogeneity in expression between subgroups of central oscillator cells and identifies a novel enhancer region.</a> J Biol Rhythms. 24 (5), 353-367 (2009).</li></ol>

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.5M EDTA pH8.0</td>
			<td>Invitrogen</td>
			<td>AM9260G</td>
			<td></td>
		</tr>
		<tr>
			<td>5-alpha Competent&#160;E. coli&#160;(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&#39;s Drosophila 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

Research

JoVE Journal

Neuroscience

576 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

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed in sensory hairs called sensilla, which cover the surface of olfactory organs. The surface of each sensillum is covered with tiny pores, through which odorants pass and dissolve in a fluid called sensillum lymph, which bathes the sensory dendrites of the OSNs housed in a given sensillum. The OSN dendrites express odorant receptor (OR) proteins, which in insects function as odor-gated ion channels4, 5. The interaction of odorants with ORs either increases or decreases the basal firing rate of the OSN. This neuronal activity in the form of action potentials embodies the first representation of the quality, intensity, and temporal characteristics of the odorant6, 7.
Given the easy access to these sensory hairs, it is possible to perform extracellular recordings from single OSNs by introducing a recording electrode into the sensillum lymph, while the reference electrode is placed in the lymph of the eye or body of the insect. In Drosophila, sensilla house between one and four OSNs, but each OSN typically displays a characteristic spike amplitude. Spike sorting techniques make it possible to assign spiking responses to individual OSNs. This single sensillum recording (SSR) technique monitors the difference in potential between the sensillum lymph and the reference electrode as electrical spikes that are generated by the receptor activity on OSNs1, 2, 8. Changes in the number of spikes in response to the odorant represent the cellular basis of odor coding in insects. Here, we describe the preparation method currently used in our lab to perform SSR on Drosophila melanogaster and Anopheles gambiae, and show representative traces induced by the odorants in a sensillum-specific manner.

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

Electrophysiological responses of olfactory sensory neurons to odorants can be measured in insects using single sensillum recordings. In this video article we will demonstrate how to perform single sensillum recordings in the antennae of the vinegar fly (Drosophila melanogaster) and the maxillary palps of the malaria mosquito (Anopheles gambiae).

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed ...

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

A growing number of genetically encoded tools are becoming available that allow non-invasive manipulation of the neural activity of specific neurons in Drosophila melanogaster1. Chief among these are optogenetic tools, which enable the activation or silencing of specific neurons in the intact and freely moving animal using bright light. Channelrhodopsin (ChR2) is a light-activated cation channel that, when activated by blue light, causes depolarization of neurons that express it. ChR2 has been effective for identifying neurons critical for specific behaviors, such as CO2 avoidance, proboscis extension and giant-fiber mediated startle response2-4. However, as the intense light sources used to stimulate ChR2 also stimulate photoreceptors, these optogenetic techniques have not previously been used in the visual system. Here, we combine an optogenetic approach with a mutation that impairs phototransduction to demonstrate that activation of a cluster of loom-sensitive neurons in the fly's optic lobe, Foma-1 neurons, can drive an escape behavior used to avoid collision. We used a null allele of a critical component of the phototransduction cascade, phospholipase C-&beta;, encoded by the norpA gene, to render the flies blind and also use the Gal4-UAS transcriptional activator system to drive expression of ChR2 in the Foma-1 neurons. Individual flies are placed on a small platform surrounded by blue LEDs. When the LEDs are illuminated, the flies quickly take-off into flight, in a manner similar to visually driven loom-escape behavior. We believe that this technique can be easily adapted to examine other behaviors in freely moving flies.

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

Genetically encoded optogenetic tools enable noninvasive manipulation of specific neurons in the Drosophila brain. Such tools can identify neurons whose activation is sufficient to elicit or suppress particular behaviors. Here we present a method for activating Channelrhodopsin2 that is expressed in targeted neurons in freely walking flies.

A  growing number of genetically encoded tools are becoming available that allow non-invasive  manipulation of the neural activity of specific neurons in ...

Determination of the Spontaneous Locomotor Activity in Drosophila melanogaster

Drosophila melanogaster has been used as an excellent model organism to study environmental and genetic manipulations that affect behavior. One such behavior is spontaneous locomotor activity. Here we describe our protocol that utilizes Drosophila population monitors and a tracking system that allows continuous monitoring of the spontaneous locomotor activity of flies for several days at a time. This method is simple, reliable, and objective and can be used to examine the effects of aging, sex, changes in caloric content of food, addition of drugs, or genetic manipulations that mimic human diseases.

Determination of the Spontaneous Locomotor Activity in Drosophila melanogaster

Drosophila melanogaster are useful in studying genetic or environmental manipulations that affect behaviors such as spontaneous locomotor activity.&#160; Here we describe a protocol that utilizes monitors with infrared beams and data analysis software to quantify spontaneous locomotor activity.&#160;

Drosophila melanogaster has been used as an excellent model organism to study environmental and genetic manipulations that affect behavior. One such ...

Straightforward Assay for Quantification of Social Avoidance in Drosophila melanogaster

Drosophila melanogaster is an emerging model to study different aspects of social interactions. For example, flies avoid areas previously occupied by stressed conspecifics due to an odorant released during stress known as the Drosophila stress odorant (dSO). Through the use of the T-maze apparatus, one can quantify the avoidance of the dSO by responder flies in a very affordable and robust assay. Conditions necessary to obtain a strong performance are presented here. A stressful experience is necessary for the flies to emit dSO, as well as enough emitter flies to cause a robust avoidance response to the presence of dSO. Genetic background, but not their group size, strongly altered the avoidance of the dSO by the responder flies. Canton-S and Elwood display a higher performance in avoiding the dSO than Oregon and Samarkand strains. This behavioral assay will allow identification of mechanisms underlying this social behavior, and the assessment of the influence of genes and environmental conditions on both emission and avoidance of the dSO. Such an assay can be included in batteries of simple diagnostic tests used to identify social deficiencies of mutants or environmental conditions of interest.

Straightforward Assay for Quantification of Social Avoidance in Drosophila melanogaster

Here, we present a protocol to quantify the avoidance of stressed individuals. This paradigm is powerful yet user-friendly and can be used to assess the influence of genes and environment on one kind of social interaction in Drosophila melanogaster.

Drosophila melanogaster is an emerging model to study different aspects of social interactions. For example, flies avoid areas previously occupied by ...

Conditions Affecting Social Space in Drosophila melanogaster

The social space assay described here can be used to quantify social interactions of Drosophila melanogaster &#8212; or other small insects &#8212; in a straightforward manner. As we previously demonstrated 1, in a two-dimensional chamber, we first force the flies to form a tight group, subsequently allowing them to take their preferred distance from each other. After the flies have settled, we measure the distance to the closest neighbor (or social space), processing a static picture with free online software (ImageJ). The analysis of the distance to the closest neighbor allows researchers to determine the effects of genetic and environmental factors on social interaction, while controlling for potential confounding factors. Diverse factors such as climbing ability, time of day, sex, and number of flies, can modify social spacing of flies. We thus propose a series of experimental controls to mitigate these confounding effects. This assay can be used for at least two purposes. First, researchers can determine how their favorite environmental shift (such as isolation, temperature, stress or toxins) will impact social spacing 1,2. Second, researchers can dissect the genetic and neural underpinnings of this basic form of social behavior 1,3. Specifically, we used it as a diagnostic tool to study the role of orthologous genes thought to be involved in social behavior in other organisms, such as candidate genes for autism in humans 4.

Conditions Affecting Social Space in Drosophila melanogaster

The effect of genes and environment on social space of Drosophila melanogaster can be quantified through a powerful but straightforward analytical paradigm. We show here different factors that can affect this social space, and thus need to be taken into consideration when designing experiments in this paradigm.

The social space assay described here can be used to quantify social interactions of Drosophila melanogaster &#8212; or other small insects &#8212; in a ...

High-resolution Quantification of Odor-guided Behavior in Drosophila melanogaster Using the Flywalk Paradigm

In their natural environment, insects such as the vinegar fly Drosophila melanogaster are bombarded with a huge amount of chemically distinct odorants. To complicate matters even further, the odors detected by the insect nervous system usually are not single compounds but mixtures whose composition and concentration ratios vary. This leads to an almost infinite amount of different olfactory stimuli which have to be evaluated by the nervous system.
To understand which aspects of an odor stimulus determine its evaluation by the fly, it is therefore desirable to efficiently examine odor-guided behavior towards many odorants and odor mixtures. To directly correlate behavior to neuronal activity, behavior should be quantified in a comparable time frame and under identical stimulus conditions as in neurophysiological experiments. However, many currently used olfactory bioassays in Drosophila neuroethology are rather specialized either towards efficiency or towards resolution.
Flywalk, an automated odor delivery and tracking system, bridges the gap between efficiency and resolution. It allows the determination of exactly when an odor packet stimulated a freely walking fly, and to determine the animal&#180;s dynamic behavioral reaction.

High-resolution Quantification of Odor-guided Behavior in Drosophila melanogaster Using the Flywalk Paradigm

The automated tracking system Flywalk is used for high-resolution quantification of odor-guided behavior in Drosophila melanogaster.

In their natural environment, insects such as the vinegar fly Drosophila melanogaster are bombarded with a huge amount of chemically distinct odorants. To ...

A Behavioral Assay for Mechanosensation of MARCM-based Clones in Drosophila melanogaster

Because of the structural and functional homology to the hair cells of the mammalian inner ear, the neurons that innervate the Drosophila external sense organs provide an excellent model system for the study of mechanosensation. This protocol describes a simple touch behavior in fruit flies which can be used to identify mutations that interfere with mechanosensation. The tactile stimulation of a macrochaete bristle on the thorax of flies elicits a grooming reflex from either the first or third leg. Mutations that interfere with mechanotransduction (such as NOMPC), or with other aspects of the reflex arc, can inhibit the grooming response. A traditional screen of adult behaviors would have missed mutants that have essential roles during development. Instead, this protocol combines the touch screen with mosaic analysis with a repressible cell marker (MARCM) to allow for only limited regions of homozygous mutant cells to be generated and marked by the expression of green fluorescent protein (GFP). By testing MARCM clones for abnormal behavioral responses, it is possible to screen a collection of lethal p-element mutations to search for new genes involved in mechanosensation that would have been missed by more traditional methods.

A Behavioral Assay for Mechanosensation of MARCM-based Clones in Drosophila melanogaster

In order to identify novel mutations affecting mechanosensation, we designed an assay that measures the behavioral response to tactile stimulation of fly bristles in mutant clones generated by the MARCM method. The combination of techniques allows for the identification of mechanosensitive mutations that would otherwise be lethal.

Because of the structural and functional homology to the hair cells of the mammalian inner ear, the neurons that innervate the Drosophila external sense ...

Foraging Path-length Protocol for Drosophila melanogaster Larvae

The Drosophila melanogaster larval path-length phenotype is an established measure used to study the genetic and environmental contributions to behavioral variation. The larval path-length assay was developed to measure individual differences in foraging behavior that were later linked to the foraging gene. Larval path-length is an easily scored trait that facilitates the collection of large sample sizes, at minimal cost, for genetic screens. Here we provide a detailed description of the current protocol for the larval path-length assay first used by Sokolowski. The protocol details how to reproducibly handle test animals, perform the behavioral assay and analyze the data. An example of how the assay can be used to measure behavioral plasticity in response to environmental change, by manipulating feeding environment prior to performing the assay, is also provided. Finally, appropriate test design as well as environmental factors that can modify larval path-length such as food quality, developmental age and day effects are discussed.

Foraging Path-length Protocol for Drosophila melanogaster Larvae

We provide a detailed protocol for a Drosophila melanogaster foraging path-length assay. We discuss the preparation and handling of test animals, how to perform the assay and analyze the data.

The Drosophila melanogaster larval path-length phenotype is an established measure used to study the genetic and environmental contributions to behavioral ...

The CApillary FEeder Assay Measures Food Intake in Drosophila melanogaster

For most animals, feeding is an essential behavior for securing survival, and it influences development, locomotion, health and reproduction. Ingestion of the right type and quantity of food therefore has a major influence on quality of life. Research on feeding behavior focuses on the underlying processes that ensure actual feeding and unravels the role of factors regulating internal energy homeostasis and the neuronal bases of decision-making. The model organism Drosophila melanogaster, with its great variety of genetically traceable tools for labeling and manipulating single neurons, allows mapping of neuronal networks and identification of molecular signaling cascades involved in the regulation of food intake. This report demonstrates the CApillary FEeder assay (CAFE) and shows how to measure food intake in a group of flies for time spans ranging from hours to days. This easy-to-use assay consists of glass capillaries filled with liquid food that flies can freely access and feed on. Food consumption in the assay is accurately determined using simple measurement tools. Herein we describe step-by-step the method from setup to successful execution of the CAFE assay, and provide practical examples to analyze the food intake of a group of flies under controlled conditions. The reader is guided through possible limitations of the assay, and advantages and disadvantages of the method compared to other feeding assays in D. melanogaster are evaluated.

The CApillary FEeder Assay Measures Food Intake in Drosophila melanogaster

The CApillary FEeder (CAFE) assay is a simple, budget-friendly, highly reliable method for investigating mechanisms underlying food intake. Used with the highly versatile genetic model organism Drosophila melanogaster, it provides a powerful means of gaining new insights into regulatory mechanisms of food intake.

For most animals, feeding is an essential behavior for securing survival, and it influences development, locomotion, health and reproduction. Ingestion of ...

A Simple One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the molecular and cellular basis of higher brain functions. A whole-mount preparation of adult brains with well-preserved morphology is critical for such whole brain-based studies, but can be technically challenging and time-consuming. This protocol describes an easy-to-learn, one-step dissection approach of an adult fly head in less than 10 s, while keeping the intact brain attached to the rest of the body to facilitate subsequent processing steps. The procedure helps remove most of the eye and tracheal tissues normally associated with the brain that can interfere with the later imaging step, and also places less demand on the quality of the dissecting forceps. Additionally, we describe a simple method that allows convenient flipping of the mounted brain samples on a coverslip, which is important for imaging both sides of the brains with similar signal intensity and quality. As an example of the protocol, we present an analysis of dopaminergic (DA) neurons in adult brains of WT (w1118) flies. The high efficacy of the dissection method makes it particularly useful for large-scale adult brain-based studies in Drosophila.

A Simple One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

The adult Drosophila brain is a valuable system for studying neuronal circuitry, higher brain functions, and complex disorders. An efficient method to dissect whole brain tissue from the small fly head will facilitate brain-based studies. Here we describe a simple, one-step dissection protocol of adult brains with well-preserved morphology.

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the ...