There are no conflicts of interest related to this report.

All animal care and experimental procedures followed the guideline from the National Institutes of Health and the Institutional Animal Care and Use Committee (IACUC) at the University of Michigan. Use RNase-free solutions and pipette tips throughout the protocol. Clean the working area, pipettes, tubes, and centrifuge with RNase Decontamination solution following the manufacturer guidelines (see Materials Table).
1. sgRNA Target Selection
<ol>
	<li>Using a web browser, open the webpage:&#160;http://crispor.tefor.net8
	<ol>
		<li>Input ~60 base pairs (bp) of sequence flanking the desired edit of interest (i.e. 30 bp 5&#39; and 3&#39; of desired edit).</li>
		<li>Select the Caenorhabditis elegans genome from the pulldown menu.</li>
		<li>Select the Protospacer Adjacent Motif (20 bp-NGG - SpCas9, SpCas9-HF1, eSPCas9 1.1) from the pulldown menu.</li>
		<li>Click submit. In the results page, choose the top ranked target sequence closest to the edit of interest. If there are no suitable target sites within the flanking 60 bp input sequence, expand the query sequence up to 100 bp (i.e. 50 bp either side of the desired edit).</li>
	</ol>
	</li>
	<li>From an available source, e.g. synthego, obtain a 20-mer target sgRNA with the following specifications: 3 nmol; no modifications.</li>
	<li>Upon receipt, centrifuge lyophilized sgRNA containing tube at &#62;12,000 x g for 1 min at room temperature. Add 60 &#181;L of nuclease-free TE to the tube, and gently re-suspend by pipetting up and down 15 - 20 times using a P200 pipette. The final concentration is 50 &#181;M.</li>
</ol>
2. Homology-directed Repair Template Design
<ol>
	<li>Design an ssODN HDR template containing: 1) the mutation of interest, 2) a unique in-frame restriction endonuclease site, 3) a silent mutation of one or both of the Gs within the NGG PAM sequence, and 4) 50 bp 5&#39; and 3&#39; homology arms flanking the first and last mutation. (Figure 1C)9,10.
	<ol>
		<li>If mutation of the NGG PAM sequence is not possible, introduce 5 - 6 silent mutations within the sgRNA target recognition sequence to prevent sgRNA-mediated cleavage of the ssODN HDR template.</li>
	</ol>
	</li>
	<li>From an available source, e.g. idtdna, obtain an &#34;Ultramer DNA oligonucleotide&#34; with the following parameters: Scale: 4 nmol; Formulation: None; Purification: Standard Desalting.</li>
	<li>Upon receipt, centrifuge lyophilized ssODN containing tube at &#62;12,000 x g for 1 min at room temperature. Gently re-suspend ssODN to a final concentration of 100 &#181;M in nuclease-free ddH2O by pipetting up and down 15 - 20 times using a P200 pipette.</li>
</ol>
3. Design Genotyping Primers
<ol>
	<li>Design a primer set flanking the genome modification to generate a ~400 - 700 bp PCR amplicon11.</li>
	<li>Optimize PCR cycling conditions to produce a single and specific amplicon11.</li>
</ol>
4. Prepare Injection Mix
<ol>
	<li>Centrifuge Table 1 reagents at maximum speed for 2 min at 4 &#176;C.</li>
	<li>In the order listed, add Table 1 reagents to a nuclease-free PCR tube.</li>
	<li>Mix thoroughly by gently pipetting up and down 10 times with a P20 pipette.</li>
	<li>Incubate injection mix for 10 min at room temperature.</li>
</ol>
5. Injection Protocol
<ol>
	<li>Load injection micropipette with approximately one-half of the injection mix12. Save remaining injection mix in case the injection needle clogs or breaks.</li>
	<li>Break micropipette tip so that ~20 - 30 p.s.i. produces a gradual flowing solution12. 
	NOTE: Larger diameter tips negatively affect animal health and decrease F1 progeny yield.</li>
	<li>Inject 10 - 15 young adult worms in both gonadal arms if possible12. If not, injection into one gonadal arm is sufficient.</li>
	<li>Allow injected animals (P0) to recover for 1 - 2 h at room temperature on an OP50-seeded 35 mm nematode growth media (NGM) plate. Subsequently, single injected P0 animals to individual OP50-seeded 35 mm NGM plates using a platinum wire worm pick12.</li>
</ol>
6. Screen P0 Plates and Single mCherry(+) F1s
<ol>
	<li>Two days post-injection, identify P0 plates containing F1 progeny expressing mCherry in the pharynx using a fluorescent stereomicroscope (Figure 1D, day 2 -&#160;3). Choose three P0 plates that contain the most mCherry(+) F1 animals.</li>
	<li>From each of the three selected P0 plates, single 8 - 12 mCherry(+) F1 animals for a total of 24-36 mCherry(+) F1s using a worm pick (Figure 1D, day 2 - 3). 
	NOTE: mCherry(-) animals can also be singled from these P0 plates, but the probability of identifying correctly edited animals is much lower (Figure 2A).</li>
	<li>Allow individual F1s to self-fertilize and lay eggs for 1 - 2 days at room temperature.</li>
</ol>
7. Single Worm PCR and Genotyping
<ol>
	<li>After 1 - 2 days of egg-laying, transfer the F1s into individual PCR strip tube caps containing 7 &#181;L of Worm Lysis Buffer using a worm pick (Table 2, Figure 1D, day 4 - 5).</li>
	<li>Centrifuge PCR tubes at maximum speed for 1 min at room temperature to bring animals to the tube bottom, and freeze tubes at -80 &#176;C for 1 h. 
	NOTE: Worms can be stored at -80 &#176;C indefinitely.</li>
	<li>Lyse frozen worms in a thermocycler using the following program: 60 &#176;C for 60 min, 95 &#176;C for 15 min, 4 &#176;C hold.</li>
	<li>Set-up PCR mastermix as shown in Table 3.</li>
	<li>Add 21 &#181;L of PCR mastermix into clean PCR tubes, and then add 4 &#181;L of worm lysis from step 3. Mix well using a pipette and run PCR program following the manufacturers guidelines (see Materials Table).</li>
	<li>Purify PCR reactions using a DNA Clean and Concentrate kit, following the manufacturer instructions (see Materials Table). Elute DNA by adding 10 &#181;L water to the spin column.</li>
	<li>Set-up restriction enzyme mastermix as shown in Table 4.</li>
	<li>Add 10 &#181;L of the restriction enzyme mastermix to each cleaned PCR reaction and incubate for 1 - 2 h at 37 &#176;C13.</li>
	<li>Separate digested PCR products on a 1.5% agarose gel run at 120 V using 1x&#160;Tris-acetate-EDTA (TAE) buffer14.</li>
</ol>
8. Identification and Sequence Verification of Edited Animals
<ol>
	<li>Examine enzyme digested samples for the presence of bands that indicate potential edited animals14 (Figure 1D, day 4 - 5). 
	NOTE: Load an N2 control to identify potential indel mutations which can be observed as shifts in band size and/or unexpected and complex restriction enzyme digestion banding patterns.</li>
	<li>After identifying potential edited animals, use the remaining 3 &#181;L of worm lysis and repeat the PCR reaction. Do not clean or restriction enzyme digest the PCR reaction after the program is complete. Load the PCR reaction on a 1% agarose gel, separate and extract the band using a gel extraction kit15 (see Materials Table).</li>
	<li>Sanger sequence16 gel extracted DNA using the F1 primer to identify correctly edited animals (Figure 1A) 
	NOTE: Heterozygote reads will contain overlaid peaks due to the presence of the wild type and engineered allele.</li>
	<li>To obtain homozygous animals from verified heterozygous edited animals, single 8 - 12 F2 animals and perform steps 7 - 8.</li>
</ol>

<ol>
	<li>Dickinson, D. J., Goldstein, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CRISPR-Based+Methods+for+Caenorhabditis+elegans+Genome+Engineering.">CRISPR-Based Methods for Caenorhabditis elegans Genome Engineering.</a> Genetics. 202, 885-901 (2016).</li><li>Doudna, J. A., Charpentier, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genome+editing.+The+new+frontier+of+genome+engineering+with+CRISPR-Cas9.">Genome editing. The new frontier of genome engineering with CRISPR-Cas9.</a> Science. 346, 1258096 (2014).</li><li>Ma, D., Liu, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genome+Editing+and+Its+Applications+in+Model+Organisms.">Genome Editing and Its Applications in Model Organisms.</a> Genomics Proteomics Bioinformatics. 13, 336-344 (2015).</li><li>Sander, J. D., Joung, J. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CRISPR-Cas+systems+for+editing,+regulating+and+targeting+genomes.">CRISPR-Cas systems for editing, regulating and targeting genomes.</a> Nat Biotechnol. 32, 347-355 (2014).</li><li>Kim, H. M., Colaiacovo, M. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CRISPR-Cas9-Guided+Genome+Engineering+in+C.+elegans.">CRISPR-Cas9-Guided Genome Engineering in C. elegans.</a> Curr Protoc Mol Biol. 115, 31-31 (2016).</li><li>Yin, H., Kauffman, K. J., Anderson, D. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Delivery+technologies+for+genome+editing.">Delivery technologies for genome editing.</a> Nat Rev Drug Discov. 16, 387-399 (2017).</li><li>Kim, S., Kim, D., Cho, S. W., Kim, J., Kim, J. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Highly+efficient+RNA-guided+genome+editing+in+human+cells+via+delivery+of+purified+Cas9+ribonucleoproteins.">Highly efficient RNA-guided genome editing in human cells via delivery of purified Cas9 ribonucleoproteins.</a> Genome Res. 24, 1012-1019 (2014).</li><li>Haeussler, 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=Evaluation+of+off-target+and+on-target+scoring+algorithms+and+integration+into+the+guide+RNA+selection+tool+CRISPOR.">Evaluation of off-target and on-target scoring algorithms and integration into the guide RNA selection tool CRISPOR.</a> Genome Biol. 17, 148 (2016).</li><li>Paix, A., Folkmann, A., Seydoux, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Precision+genome+editing+using+CRISPR-Cas9+and+linear+repair+templates+in+C.+elegans.">Precision genome editing using CRISPR-Cas9 and linear repair templates in C. elegans.</a> Methods. , (2017).</li><li>Prior, H., Jawad, A. K., MacConnachie, L., Beg, A. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Highly+Efficient,+Rapid+and+Co-CRISPR-Independent+Genome+Editing+in+Caenorhabditis+elegans.">Highly Efficient, Rapid and Co-CRISPR-Independent Genome Editing in Caenorhabditis elegans.</a> G3 (Bethesda). 7, 3693-3698 (2017).</li><li>JoVE Science Education Database. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Basic+Methods+in+Cellular+and+Molecular+Biology.+PCR:+The+Polymerase+Chain+Reaction.">Basic Methods in Cellular and Molecular Biology. PCR: The Polymerase Chain Reaction.</a> J Vis Exp. , (2017).</li><li>Berkowitz, L. A., Knight, A. L., Caldwell, G. A., Caldwell, 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=Generation+of+stable+transgenic+C.+elegans+using+microinjection.">Generation of stable transgenic C. elegans using microinjection.</a> J Vis Exp. , (2008).</li><li>JoVE Science Education Database. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=+Basic+Methods+in+Cellular+and+Molecular+Biology.+Restriction+Enzyme+Digests."> Basic Methods in Cellular and Molecular Biology. Restriction Enzyme Digests.</a> J Vis Exp. , (2017).</li><li>JoVE Science Education Database. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Basic+Methods+in+Cellular+and+Molecular+Biology.+DNA+Gel+Electrophoresis+.">Basic Methods in Cellular and Molecular Biology. DNA Gel Electrophoresis .</a> J Vis Exp. , (2017).</li><li>JoVE Science Education Database. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Basic+Methods+in+Cellular+and+Molecular+Biology.+Gel+Purification.">Basic Methods in Cellular and Molecular Biology. Gel Purification.</a> J Vis Exp. , (2017).</li><li>Estrada-Rivadeneyra, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sanger+sequencing.">Sanger sequencing.</a> FEBS J. , (2017).</li><li>Ludolph, A. C., Brettschneider, J., Weishaupt, J. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Amyotrophic+lateral+sclerosis.">Amyotrophic lateral sclerosis.</a> Curr Opin Neurol. 25, 530-535 (2012).</li><li>Evans, T. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> WormBook. , (2006).</li><li>Mello, C. C., Kramer, J. M., Stinchcomb, D., Ambros, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficient+gene+transfer+in+C.elegans:+extrachromosomal+maintenance+and+integration+of+transforming+sequences.">Efficient gene transfer in C.elegans: extrachromosomal maintenance and integration of transforming sequences.</a> EMBO J. 10, 3959-3970 (1991).</li><li>Jinek, 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+programmable+dual-RNA-guided+DNA+endonuclease+in+adaptive+bacterial+immunity.">A programmable dual-RNA-guided DNA endonuclease in adaptive bacterial immunity.</a> Science. 337, 816-821 (2012).</li><li>Boyd, S. D., 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+Balanced+Look+at+the+Implications+of+Genomic+(and+Other+&amp;#34;Omics&amp;#34;)+Testing+for+Disease+Diagnosis+and+Clinical+Care.">A Balanced Look at the Implications of Genomic (and Other &amp;#34;Omics&amp;#34;) Testing for Disease Diagnosis and Clinical Care.</a> Genes (Basel). 5, 748-766 (2014).</li><li>Saccon, R. A., Bunton-Stasyshyn, R. K., Fisher, E. M., Fratta, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Is+SOD1+loss+of+function+involved+in+amyotrophic+lateral+sclerosis?.">Is SOD1 loss of function involved in amyotrophic lateral sclerosis?.</a> Brain. 136, 2342-2358 (2013).</li><li>Acevedo-Arozena, 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=A+comprehensive+assessment+of+the+SOD1G93A+low-copy+transgenic+mouse,+which+models+human+amyotrophic+lateral+sclerosis.">A comprehensive assessment of the SOD1G93A low-copy transgenic mouse, which models human amyotrophic lateral sclerosis.</a> Dis Model Mech. 4, 686-700 (2011).</li><li>Gurney, M. E., 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=Motor+neuron+degeneration+in+mice+that+express+a+human+Cu,Zn+superoxide+dismutase+mutation.">Motor neuron degeneration in mice that express a human Cu,Zn superoxide dismutase mutation.</a> Science. 264, 1772-1775 (1994).</li></ol>

We thank members of the Beg laboratory for critical reading of this manuscript. Strains were provided by the Caenorhabditis Genetics Center, which is funded by NIH Office of Research Infrastructure Programs (P40 OD010440). Research in the Beg laboratory is supported by grants from the NIH (R01 NS094678) and Muscular Dystrophy Association (MDA382300) to A.A.B.

The CRISPR-Cas9 system is a powerful and effective tool for precisely modifying the genome of model organisms. Here, we demonstrate that chimeric sgRNAs20 coupled with ssODN HDR templates enable the highly efficient generation of genomic point mutations in C. elegans. Importantly, we demonstrate that RNP delivery produces high editing efficiency when fluorescence is used as a co-selection marker, highlighting the ease and reliability of the technique.
The majority of CRISPR-Cas9 methods for C. elegans genome engineering rely on specific genetic backgrounds or co-CRISPR strategies1. The co-CRISPR method has proven to be an incredibly useful tool that increases the likelihood of obtaining a successful genome edit. However, co-CRISPR methods require introducing a phenotype-selectable edit at a marker locus that enriches for the genome edit of interest. While powerful, introducing selectable phenotype-bearing mutations may be undesirable if the strain to edit or the desired point mutation of interest itself exhibits phenotypes that might be exacerbated or suppressed by the marker phenotype. Moreover, co-CRISPR strategies necessitate creating an additional double strand break at a marker locus that may contribute to off-target effects. Here, we present a method that uses a transient fluorescent marker that not only serves to identify successfully injected P0 animals, but also enriches for proper genome edits. Our data demonstrate this method significantly enriches for successfully edited animals in comparison to non-fluorescent animals, and provides several distinct advantages: 1) only one targeting site-specific sgRNA is required, 2) a 5-fold reduction in Cas9 and sgRNA concentration, 3) strain- and phenotype-selection independence, and 4) a nominal screening workload (~24 F1s). Robust CRISPR-Cas9 genome editing in the F1 generation was observed using this method. Notably, the PCR-restriction enzyme-based detection strategy described enables the unequivocal detection of mono- and bi-allelic edited animals in the F1 generation. Finally, the genotyping strategy facilitates identification of potential indel mutations, which can be observed as PCR band shifts when compared to N2 controls, that are either not susceptible to restriction digest or yield complex bands when digested.
The advent of next generation sequencing coupled with large scale genomic efforts has accelerated the identification of genomic variants that segregate with disease21. However, functional tests of pathogenicity have not been demonstrated for the majority of variants in cellular and organismal model systems. In this study, we targeted the sod-1 gene in C. elegans to introduce the widely studied ALS-associated SOD-1G93A mutation. To date, this mutation has been modeled and studied in C. elegans and mice primarily using transgenic overexpression. Although overexpression of variants in animal models may recapitulate key cellular, molecular and behavioral aspects of disease, it is increasingly clear that &#39;dose&#39; is an extenuating factor in determining phenotypes22. For example, high copy number SOD-1G93A mice carrying ~24 copies of the mutant human SOD-1 gene exhibit greatly accelerated disease course compared to SOD-1G93Adl mice, which carry only ~8 - 10 copies23,24. Here, we demonstrate that our CRISPR-Cas9 RNP-based method can be utilized to rapidly generate human variant-associated mutations in the orthologous worm gene without requiring transgenic overexpression, in an effort to more precisely replicate the genetic context of disease. Our method provides a feasible platform to produce and test genomic variants of unknown significance in the context of endogenous regulatory control and expression, which precludes the confines of overexpression. Finally, we have also used this method to tag endogenous genes with fluorescent proteins (i.e. GFP), which will provide an additional tool for visualizing how variants affect protein localization, aggregation, and turnover.

Recent technological advances have radically transformed and accelerated the ability to precisely engineer genomes. In particular, the CRISPR-Cas9 system, which relies on the RNA-guided endonuclease Cas9 to induce a double strand break (DSB) near the target sequence of interest, has been extensively used to accurately engineer the genome of the majority of model organisms used in biomedical research1,2,3,4. Significantly, the use of CRISPR-Cas9 has unlocked genome editing even in difficult species like C. elegans5. Regardless of species, generating point mutations with the CRISPR-Cas9 based genome editing system relies on three core components: 1) Cas9 endonuclease, 2) a single guide RNA (sgRNA) that directs the Cas9 endonuclease to a target sequence, and 3) a user designed homology-directed repair (HDR) template containing the desired edit(s) of interest2.
There are several methods that can be used to introduce the targeting sgRNA and Cas9 nuclease into cells including plasmid, RNA, and viral-based delivery methods6. Recently, direct delivery of pre-complexed sgRNA-Cas9 Ribonucleoproteins (RNPs) has emerged as a powerful and efficient tool in CRISPR-Cas9-based genome editing7. The direct delivery of pre-complexed CRISPR-Cas9 RNPs has several distinct advantages, namely: 1) RNPs bypass the need for cellular transcription and translation, 2) RNPs are rapidly cleared, which may increase specificity by reducing available time for off-target cleavage, and 3) RNPs contain no foreign DNA/RNA elements which circumvents the introduction of non-native sequences into the host genome through random integration. Together, these attributes likely provide a short-lived burst of on-target CRISPR editing while minimizing off-target effects.
We describe a simple and efficient protocol for introducing site-specific genomic changes in C. elegans. This protocol includes targeting sgRNA and single stranded oligonucleotide (ssODN) HDR template design, sgRNA-Cas9 RNP complexing and delivery, and a genotyping strategy for the unequivocal identification of properly edited animals. Using this strategy, not only can the desired site-specific changes be recovered, but other non-specific indel mutations may also be recovered. Thus, our strategy permits the generation of an allelic series using a single strategy, where both mono-allelic, bi-allelic, and indel mutants can be generated in the F1 generation.

<table border="0" cellpadding="0" cellspacing="0" width="1144">
	<tbody>
		<tr height="40">
			<td height="40" style="height:40px;width:359px;">CRISPRevolution sgRNA&#160; EZ Kit</td>
			<td style="width:216px;">Synthego Inc.</td>
			<td style="width:344px;"></td>
			<td style="width:227px;">chimeric sgRNA</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Nuclease-free TE</td>
			<td>Synthego Inc.</td>
			<td>provided with the sgRNA kit EZ kit</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Nuclease-free water</td>
			<td>Synthego Inc.</td>
			<td>provided with the sgRNA kit EZ kit</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">4 nmole Ultramer DNA Oligo</td>
			<td>Integrated DNA Technologies</td>
			<td></td>
			<td>ssODN HDR template</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Alt-R S.p. HiFi Cas9 Nuclease 3NLS</td>
			<td>Integrated DNA Technologies</td>
			<td>1078728</td>
			<td>Cas9 protein</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">pCFJ90&#160;</td>
			<td>Addgene</td>
			<td>19327</td>
			<td>Pmyo-2::mCherry Marker Plasmid</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">KCl</td>
			<td>Sigma</td>
			<td>P5405</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">HEPES</td>
			<td>Sigma</td>
			<td>H4034</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">DNA Clean &#38; Concentrator</td>
			<td>Zymo Research</td>
			<td>D4004</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Zymoclean Gel DNA Recovery Kit</td>
			<td>Zymo Research</td>
			<td>D4002</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Q5 Hot Start High-Fidelity 2X Master Mix</td>
			<td>New England Biolabs</td>
			<td>M0494L</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Proteinase K</td>
			<td>Sigma</td>
			<td>P2308</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Glass Borosilicate Glass Micropipettes</td>
			<td>Sutter Instruments</td>
			<td>BF100-78-10</td>
			<td>OD: 1.0mm. ID: 0.78mm</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Trizma Hydrochloride</td>
			<td>Sigma</td>
			<td>T5941</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">MgCl2</td>
			<td>Sigma</td>
			<td>M2393</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">NP-40</td>
			<td>Sigma</td>
			<td>74385</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Tween-20</td>
			<td>Fisher Scientific</td>
			<td>BP337-100</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">RNaseZap Decontamination Solution</td>
			<td>Fisher Scientific</td>
			<td>AM9780</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Mutations in human superoxide dismutase 1 (SOD1) account for ~10 - 20% of familial amyotrophic lateral sclerosis, a devastating neurodegenerative disease that invariably leads to paralysis and death17. Human SOD-1 is an evolutionarily conserved protein sharing 55% identity and 70% similarity with the C. elegans SOD-1 protein (Figure 1B). To demonstrate the simplicity, feasibility, and efficiency of the CRISPR-Cas9 RNP-based approach, we targeted the C. elegans sod-1 gene to introduce the disease-associated human G93A variant in the worm genome (Figure 1A).
To engineer the G93A mutation in the C. elegans genome, an sgRNA was chosen whose PAM recognition site sits 3 bp upstream of the G93 codon (Figure 1C). We designed an ssODN HDR repair template containing: 1) nucleotide changes to convert the G93 codon to A93 (GGA GCA), 2) a silent change to the NGG PAM sequence (CGG CAG) to prevent sgRNA-Cas9-mediated cleavage of the HDR template, 3) a silent mutation (CT) that introduces a unique HindIII restriction enzyme site for genotyping, and 4) 50 bp 5&#39; and 3&#39; homology arms (Figure 1C). A PCR primer set (F1-R1) was designed to produce a single 592 bp PCR product in N2 controls (Figure 1A). An injection mix containing the sgRNA, Cas9, ssODN and a fluorescent marker plasmid (pCFJ90) were mixed together, incubated for 10 min at room temperature, and loaded into a microinjection pipette.
Figure 1D depicts the workflow after the injection mix is loaded into an injection micropipette. On Day 0, 10 - 15 P0 animals were injected in one or both gonad arms using the standard microinjection technique18,19. Injected animals recovered for 1 - 2 h and were then individually plated on OP50-seeded NGM plates. After 2 - 3 days, successful P0 injections were identified by those having mCherry(+) F1 progeny. The top three P0 plates (i.e. those having the greatest number of mCherry(+) F1s) were chosen for subsequent analysis. From each of these three plates (P0-8, P0-10, P0-12), 8 mCherry(+) F1s were singled to individual OP50-seeded 35 mm NGM plates for a total of 24 mCherry(+) F1s. After 2 -&#160;3 days of egg-laying (Day 4 - 5), the individual F1s were transferred into PCR tubes containing worm lysis buffer and frozen for 1 h at -80 &#176;C. Subsequently, the F1s were lysed to liberate genomic DNA, and subjected to PCR. The PCR products were then purified and digested with HindIII for 1 h, and run on a 1.5% agarose gel to identify potential edited animals. In correctly edited animals, HindIII digestion cuts the full-length PCR product (592 bp) into two bands of 370 bp and 222 bp. This genotyping strategy unambiguously differentiates between wild-type (592 bp), heterozygote (592, 370, 222 bp) and homozygote (370, 222 bp) animals. Figure 2A illustrates the genotyping results for the 24 mCherry(+) animals.
To demonstrate that the fluorescent mCherry marker enriches for genome modification, we also picked 8 mCherry(-) animals from each of the three P0 plates. Of the 24 mCherry(+) F1s screened (red bar), 10 contained mono-allelic or bi-allelic modification (42%), 10 were wild type (42%), 3 were potential indels (13%), and there was 1 PCR failure (Figure 2A). In contrast, of the 24 mCherry(-) F1s screened, only 1 animal was correctly modified (4%); whereas, 23 were wild type (96%) (Figure 2A). Figure 2B shows the chromatogram from the full-length gel extracted PCR product (F18-6), demonstrating that the precise nucleotide changes designed in the ssODN HDR template were faithfully incorporated into the genome of this homozygous F1 animal. Notably, F1 animals 10-7, 10-8, and 12-3 exhibited unexpected PCR product sizes and restriction digest banding patterns, suggesting these animals may carry complex indel mutations (Figure 2A). Thus, our method is not only capable of generating desired genome modification(s) with ease, efficiency, and fidelity, but also permits the identification and recovery of unique indel alleles that may shed important and unexpected insight into gene function.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57518/57518fig1.jpg" /> 
Figure 1. CRISPR-Cas9 engineering of the C. eleganssod-1 locus. (A) Schematic illustration depicting the exon-intron structure of the sod-1 locus in C. elegans. The red bar denotes the location of G93 in exon 3. The primer pair set is noted by the red arrows (F1-R1). (B) Sequence alignment of human and C. elegans (worm) SOD-1 proteins. The targeted G93 residue is highlighted in red. (C) Top, a section of genomic DNA surrounding the G93 codon is shown as a reference, and the PAM (green) and sgRNA target (blue) sites are highlighted. Bottom, the single stranded oligonucleotide (ssODN) homology directed repair (HDR) template is shown containing: the codon edit changing G93 to A93, a silent change to the PAM motif to prevent cleavage by the sgRNA-Cas9 complex, a silent change to introduce a unique HindIII restriction enzyme site (yellow box), and flanking 5&#39; and 3&#39; 50 bp homology arms (black bars). All nucleotide changes designed in the ssODN are highlighted red. (D) Schematic illustration of the pipeline for generating and identifying CRISPR-Cas9 RNP-based point mutants. On day 0, inject 10 - 15 P0 animals with injection mix. On day 2 - 3, identify successfully injected P0 animals by those containing fluorescent F1 progeny, and select three P0 plates with the most fluorescent progeny. From each of the three P0 plates, single 8 fluorescent F1 progeny. On day 4 - 5, (a) step 1, individual F1s are picked and placed into PCR tubes containing Lysis Buffer; (b) step 2, after freezing at -80 &#176;C, PCR tubes containing F1 worms are lysed to release genomic DNA, which is used as a PCR template. The PCR products are then cleaned and digested with the unique restriction enzyme; (c) step 3, enzyme digested products are resolved on an agarose gel where wild type (+/+), heterozygous (m/+), and homozygous (m/m) animals can be identified by restriction digest banding patterns. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57518/57518fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57518/57518fig2.jpg" /> 
Figure 2. Genotyping data for sod-1 (G93A) mutants.(A)Gel image of F1 animals that were individually lysed, PCRed and digested with HindIII. For each P0 plate (P0-8, P0-10, P0-12), 8 mCherry(+) and 8 mCherry(-) F1 animals were analyzed. Both mono-allelic (blue numbers) and bi-allelic (red numbers) edited animals were recovered. Size shifted PCR products or unpredicted HindIII digested bands were also recovered that are indicative of indel mutants (yellow numbers). N2 controls are shown as a reference for PCR product sizing and enzyme specificity. (B) Top, codon spacing and amino acid translations are shown above for the wild type (WT) and below for G93A modified strands. The desired edit is highlighted by a red box, and the silent changes that create an in-frame HindIII restriction site are highlighted by a yellow box. All designed nucleotide changes are shown in bold. Bottom, representative chromatogram from homozygous F1 animal 8-6. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57518/57518fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Reagent</td>
			<td>Volume</td>
			<td>Final Concentration</td>
		</tr>
		<tr>
			<td>ddH2O</td>
			<td>6.3 &#956;L</td>
			<td>----</td>
		</tr>
		<tr>
			<td>KCl (4M)</td>
			<td>0.94 &#956;L</td>
			<td>300 mM</td>
		</tr>
		<tr>
			<td>HEPES (0.5M) pH 7.4</td>
			<td>0.5 &#956;L</td>
			<td>20 mM</td>
		</tr>
		<tr>
			<td>pCFJ90 (25 ng/&#956;L)</td>
			<td>1.25 &#956;L</td>
			<td>2.5 ng/&#956;L</td>
		</tr>
		<tr>
			<td>ssODN (500 ng/&#956;L)</td>
			<td>1.25 &#956;L</td>
			<td>50 ng/&#956;L</td>
		</tr>
		<tr>
			<td>sgRNA (50 &#956;M)</td>
			<td>1.25 &#956;L</td>
			<td>5 &#956;M</td>
		</tr>
		<tr>
			<td>Cas9 (61 &#956;M)</td>
			<td>1.03 &#956;L</td>
			<td>5 &#956;M</td>
		</tr>
		<tr>
			<td>Final Volume</td>
			<td>12.5 &#956;L</td>
			<td>----</td>
		</tr>
	</tbody>
</table>
Table 1. CRISPR-Cas9 RNP Injection Mix. All reagents should be re-suspended in nuclease-free ddH2O. RNase free techniques should be used when handling reagents and when making the injection mix.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Reagent</td>
			<td>[Final]</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>50 mM</td>
		</tr>
		<tr>
			<td>Tris-HCl pH 8.3</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>2.5 mM</td>
		</tr>
		<tr>
			<td>NP-40</td>
			<td>0.45%</td>
		</tr>
		<tr>
			<td>Tween-20</td>
			<td>0.45%</td>
		</tr>
		<tr>
			<td>Proteinase K</td>
			<td>1 mg/mL</td>
		</tr>
	</tbody>
</table>
Table 2. Worm Lysis Buffer Recipe. Proteinase K should be added fresh before each use.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Reagent</td>
			<td>1x</td>
			<td>24x</td>
		</tr>
		<tr>
			<td>2X Q5 Mastermix</td>
			<td>12.5 &#956;L</td>
			<td>300 &#956;L</td>
		</tr>
		<tr>
			<td>Primer F1 (10 &#956;M)</td>
			<td>1.25 &#956;L</td>
			<td>30 &#956;L</td>
		</tr>
		<tr>
			<td>Primer R1 (10 &#956;M)</td>
			<td>1.25 &#956;L</td>
			<td>30 &#956;L</td>
		</tr>
		<tr>
			<td>Worm Lysis</td>
			<td>4 &#956;L</td>
			<td>----</td>
		</tr>
		<tr>
			<td>ddH2O</td>
			<td>6 &#956;L</td>
			<td>144 &#956;L</td>
		</tr>
		<tr>
			<td>Final Volume</td>
			<td>25 &#956;L</td>
			<td>504 &#956;L</td>
		</tr>
	</tbody>
</table>
Table 3. PCR Mastermix. The PCR mastermix should be mixed well by pipetting until the solution is homogeneous. 21 &#181;L of the PCR mastermix should be added to clean PCR tubes, and then 4 &#181;L of individual worm lysate should be added to properly labeled tubes.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Reagent</td>
			<td>1x</td>
			<td>24x</td>
		</tr>
		<tr>
			<td>10X Enzyme Buffer</td>
			<td>2 &#956;L</td>
			<td>48 &#956;L</td>
		</tr>
		<tr>
			<td>Cleaned PCR Reaction</td>
			<td>10 &#956;L</td>
			<td>----</td>
		</tr>
		<tr>
			<td>ddH2O</td>
			<td>7 &#956;L</td>
			<td>168 &#956;L</td>
		</tr>
		<tr>
			<td>Restriction Enzyme (10 U/&#956;L)</td>
			<td>1 &#956;L</td>
			<td>24 &#956;L</td>
		</tr>
		<tr>
			<td>Final Volume</td>
			<td>20 &#956;L</td>
			<td>240 &#956;L</td>
		</tr>
	</tbody>
</table>
Table 4. Restriction Enzyme Mastermix. The restriction enzyme mastermix should be mixed well by pipetting until the solution is homogeneous. 10 &#181;L of the restriction enzyme mastermix should be added to 10 &#181;L of cleaned PCR product

Watch this Scientific Journal Video about A Rapid and Facile Pipeline for Generating Genomic Point Mutants in C. elegans Using CRISPR/Cas9 Ribonucleoproteins at JoVE.com

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.

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 ...

a-rapid-facile-pipeline-for-generating-genomic-point-mutants-c

Nanyang Technological University

Research

JoVE Journal

Genetics

7.5K Views.  University of Michigan. Here, we present a method to engineer the genome of C. elegans using CRISPR-Cas9 ribonucleoproteins and homology dependent repair templates.

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

Optogenetic Random Mutagenesis Using Histone-miniSOG in C. elegans

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

Optogenetic Random Mutagenesis Using Histone-miniSOG in C. elegans

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

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

Using a Fluorescent PCR-capillary Gel Electrophoresis Technique to Genotype CRISPR/Cas9-mediated Knockout Mutants in a High-throughput Format

The development of programmable genome-editing tools has facilitated the use of reverse genetics to understand the roles specific genomic sequences play in the functioning of cells and whole organisms. This cause has been tremendously aided by the recent introduction of the CRISPR/Cas9 system-a versatile tool that allows researchers to manipulate the genome and transcriptome in order to, among other things, knock out, knock down, or knock in genes in a targeted manner. For the purpose of knocking out a gene, CRISPR/Cas9-mediated double-strand breaks recruit the non-homologous end-joining DNA repair pathway to introduce the frameshift-causing insertion or deletion of nucleotides at the break site. However, an individual guide RNA may cause undesirable off-target effects, and to rule these out, the use of multiple guide RNAs is necessary. This multiplicity of targets also means that a high-volume screening of clones is required, which in turn begs the use of an efficient high-throughput technique to genotype the knockout clones. Current genotyping techniques either suffer from inherent limitations or incur high cost, hence rendering them unsuitable for high-throughput purposes. Here, we detail the protocol for using fluorescent PCR, which uses genomic DNA from crude cell lysate as a template, and then resolving the PCR fragments via capillary gel electrophoresis. This technique is accurate enough to differentiate one base-pair difference between fragments and hence is adequate in indicating the presence or absence of a frameshift in the coding sequence of the targeted gene. This precise knowledge effectively precludes the need for a confirmatory sequencing step and allows users to save time and cost in the process. Moreover, this technique has proven to be versatile in genotyping various mammalian cells of various tissue origins targeted by guide RNAs against numerous genes, as shown here and elsewhere.

The genotyping technique described here, which couples fluorescent polymerase chain reaction (PCR) to capillary gel electrophoresis, allows for high-throughput genotyping of nuclease-mediated knockout clones. It circumvents limitations faced by other genotyping techniques and is more cost effective than sequencing methods.

The development of programmable genome-editing tools has facilitated the use of reverse genetics to understand the roles specific genomic sequences play ...

Selection-dependent and Independent Generation of CRISPR/Cas9-mediated Gene Knockouts in Mammalian Cells

The CRISPR/Cas9 genome engineering system has revolutionized biology by allowing for precise genome editing with little effort. Guided by a single guide RNA (sgRNA) that confers specificity, the Cas9 protein cleaves both DNA strands at the targeted locus. The DNA break can trigger either non-homologous end joining (NHEJ) or homology directed repair (HDR). NHEJ can introduce small deletions or insertions which lead to frame-shift mutations, while HDR allows for larger and more precise perturbations. Here, we present protocols for generating knockout cell lines by coupling established CRISPR/Cas9 methods with two options for downstream selection/screening. The NHEJ approach uses a single sgRNA cut site and selection-independent screening, where protein production is assessed by dot immunoblot in a high-throughput manner. The HDR approach uses two sgRNA cut sites that span the gene of interest. Together with a provided HDR template, this method can achieve deletion of tens of kb, aided by the inserted selectable resistance marker. The appropriate applications and advantages of each method are discussed.

Recent advances in the ability to genetically manipulate somatic cell lines hold great potential for basic and applied research. Here, we present two approaches for CRISPR/Cas9 generated knockout production and screening in mammalian cell lines, with and without the use of selectable markers.

The CRISPR/Cas9 genome engineering system has revolutionized biology by allowing for precise genome editing with little effort. Guided by a single guide ...

Primordial Germ Cell Transplantation for CRISPR/Cas9-based Leapfrogging in Xenopus

The creation of mutant lines by genome editing is accelerating genetic analysis in many organisms. CRISPR/Cas9 methods have been adapted for use in the African clawed frog, Xenopus, a longstanding model organism for biomedical research. Traditional breeding schemes for creating homozygous mutant lines with CRISPR/Cas9-targeted mutagenesis have several time-consuming and laborious steps. To facilitate the creation of mutant embryos, particularly to overcome the obstacles associated with knocking out genes that are essential for embryogenesis, a new method called leapfrogging was developed. This technique leverages the robustness of Xenopus embryos to &#34;cut and paste&#34; embryological methods. Leapfrogging utilizes the transfer of primordial germ cells (PGCs) from efficiently-mutagenized donor embryos into PGC-ablated wildtype siblings. This method allows for the efficient mutation of essential genes by creating chimeric animals with wildtype somatic cells that carry a mutant germline. When two F0 animals carrying &#34;leapfrog transplants&#34; (i.e., mutant germ cells) are intercrossed, they produce homozygous, or compound heterozygous, null F1 embryos, thus saving a full generation time to obtain phenotypic data. Leapfrogging also provides a new approach for analyzing maternal effect genes, which are refractory to F0 phenotypic analysis following CRISPR/Cas9 mutagenesis. This manuscript details the method of leapfrogging, with special emphasis on how to successfully perform PGC transplantation.

Primordial Germ Cell Transplantation for CRISPR/Cas9-based Leapfrogging in Xenopus

Genes essential for survival pose technical hurdles for creating mutant lines. Leapfrogging circumvents lethality by combining genome editing with primordial germ cell transplantation to create wild-type animals carrying germline mutations. Leapfrogging also permits the efficient generation of homozygous null mutants in the F1 generation. Here, the transplantation step is demonstrated.

The creation of mutant lines by genome editing is accelerating genetic analysis in many organisms. CRISPR/Cas9 methods have been adapted for use in the ...

A Standard Methodology to Examine On-site Mutagenicity As a Function of Point Mutation Repair Catalyzed by CRISPR/Cas9 and SsODN in Human Cells

Combinatorial gene editing using CRISPR/Cas9 and single-stranded oligonucleotides is an effective strategy for the correction of single-base point mutations, which often are responsible for a variety of human inherited disorders. Using a well-established cell-based model system, the point mutation of a single-copy mutant eGFP gene integrated into HCT116 cells has been repaired using this combinatorial approach. The analysis of corrected and uncorrected cells reveals both the precision of gene editing and the development of genetic lesions, when indels are created in uncorrected cells in the DNA sequence surrounding the target site. Here, the specific methodology used to analyze this combinatorial approach to the gene editing of a point mutation, coupled with a detailed experimental strategy to measuring indel formation at the target site, is outlined. This protocol outlines a foundational approach and workflow for investigations aimed at developing CRISPR/Cas9-based gene editing for human therapy. The conclusion of this work is that on-site mutagenesis takes place as a result of CRISPR/Cas9 activity during the process of point mutation repair. This work puts in place a standardized methodology to identify the degree of mutagenesis, which should be an important and critical aspect of any approach destined for clinical implementation.

This protocol outlines the workflow of a CRISPR/Cas9-based gene editing system for the repair of point mutations in mammalian cells. Here, we use a combinatorial approach to gene editing with a detailed follow-on experimental strategy for measuring indel formation at the target site&#8212;in essence, analyzing onsite mutagenesis.

Combinatorial gene editing using CRISPR/Cas9 and single-stranded oligonucleotides is an effective strategy for the correction of single-base point ...

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

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

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

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

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

Highly Efficient Gene Disruption of Murine and Human Hematopoietic Progenitor Cells by CRISPR/Cas9

Advances in the hematopoietic stem cell (HSCs) field have been aided by methods to genetically engineer primary progenitor cells as well as animal models. Complete gene ablation in HSCs required the generation of knockout mice from which HSCs could be isolated, and gene ablation in primary human HSCs was not possible. Viral transduction could be used for knock-down approaches, but these suffered from variable efficacy. In general, genetic manipulation of human and mouse hematopoietic cells was hampered by low efficiencies and extensive time and cost commitments. Recently, CRISPR/Cas9 has dramatically expanded the ability to engineer the DNA of mammalian cells. However, the application of CRISPR/Cas9 to hematopoietic cells has been challenging, mainly due to their low transfection efficiencies, the toxicity of plasmid-based approaches and the slow turnaround time of virus-based protocols.
A rapid method to perform CRISPR/Cas9-mediated gene editing in murine and human hematopoietic stem and progenitor cells with knockout efficiencies of up to 90% is provided in this article. This approach utilizes a ribonucleoprotein (RNP) delivery strategy with a streamlined three-day workflow. The use of Cas9-sgRNA RNP allows for a hit-and-run approach, introducing no exogenous DNA sequences in the genome of edited cells and reducing off-target effects. The RNP-based method is fast and straightforward: it does not require cloning of sgRNAs, virus preparation or specific sgRNA chemical modification. With this protocol, scientists should be able to successfully generate knockouts of a gene of interest in primary hematopoietic cells within a week, including downtimes for oligonucleotide synthesis. This approach will allow a much broader group of users to adapt this protocol for their needs.

A protocol for fast CRISPR/Cas9-mediated gene disruption in mouse and human primary hematopoietic cells is described in this article. Cas9-sgRNA ribonucleoproteins are introduced via electroporation with sgRNAs generated through in vitro transcription and commercial Cas9. High editing efficiencies are achieved with limited time and financial cost.

Advances in the hematopoietic stem cell (HSCs) field have been aided by methods to genetically engineer primary progenitor cells as well as animal models. ...

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

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

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

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

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

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 ...

CRISPR/Cas9 Ribonucleoprotein-mediated Precise Gene Editing by Tube Electroporation

Gene editing nucleases, represented by CRISPR-associated protein 9 (Cas9), are becoming mainstream tools in biomedical research. Successful delivery of CRISPR/Cas9 elements into the target cells by transfection is a prerequisite for efficient gene editing. This protocol demonstrates that tube electroporation (TE) machine-mediated delivery of CRISPR/Cas9 ribonucleoprotein (RNP), along with single-stranded oligodeoxynucleotide (ssODN) donor templates to different types of mammalian cells, leads to robust precise gene editing events. First, TE was applied to deliver CRISPR/Cas9 RNP and ssODNs to induce disease-causing mutations in the interleukin 2 receptor subunit gamma (IL2RG) gene and sepiapterin reductase (SPR) gene in rabbit fibroblast cells. Precise mutation rates of 3.57%-20% were achieved as determined by bacterial TA cloning sequencing. The same strategy was then used in human iPSCs on several clinically relevant genes including epidermal growth factor receptor (EGFR), myosin binding protein C, cardiac (Mybpc3), and hemoglobin subunit beta (HBB). Consistently, highly precise mutation rates were achieved (11.65%-37.92%) as determined by deep sequencing (DeepSeq). The present work demonstrates that tube electroporation of CRISPR/Cas9 RNP represents an efficient transfection protocol for gene editing in mammalian cells.

Presented here is a protocol for efficient CRISPR/Cas9 ribonucleoprotein-mediated gene editing in mammalian cells using tube electroporation.

Gene editing nucleases, represented by CRISPR-associated protein 9 (Cas9), are becoming mainstream tools in biomedical research. Successful delivery of ...