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>

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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><tbody><tr><td>CRISPRevolution sgRNA&amp;nbsp; EZ Kit</td><td>Synthego Inc.</td><td></td><td>chimeric sgRNA</td></tr><tr><td>Nuclease-free TE</td><td>Synthego Inc.</td><td>provided with the sgRNA kit EZ kit</td><td></td></tr><tr><td>Nuclease-free water</td><td>Synthego Inc.</td><td>provided with the sgRNA kit EZ kit</td><td></td></tr><tr><td>4 nmole Ultramer DNA Oligo</td><td>Integrated DNA Technologies</td><td></td><td>ssODN HDR template</td></tr><tr><td>Alt-R S.p. HiFi Cas9 Nuclease 3NLS</td><td>Integrated DNA Technologies</td><td>1078728</td><td>Cas9 protein</td></tr><tr><td>pCFJ90&amp;nbsp;</td><td>Addgene</td><td>19327</td><td>Pmyo-2::mCherry Marker Plasmid</td></tr><tr><td>KCl</td><td>Sigma</td><td>P5405</td><td></td></tr><tr><td>HEPES</td><td>Sigma</td><td>H4034</td><td></td></tr><tr><td>DNA Clean &amp;amp; Concentrator</td><td>Zymo Research</td><td>D4004</td><td></td></tr><tr><td>Zymoclean Gel DNA Recovery Kit</td><td>Zymo Research</td><td>D4002</td><td></td></tr><tr><td>Q5 Hot Start High-Fidelity 2X Master Mix</td><td>New England Biolabs</td><td>M0494L</td><td></td></tr><tr><td>Proteinase K</td><td>Sigma</td><td>P2308</td><td></td></tr><tr><td>Glass Borosilicate Glass Micropipettes</td><td>Sutter Instruments</td><td>BF100-78-10</td><td>OD: 1.0mm. ID: 0.78mm</td></tr><tr><td>Trizma Hydrochloride</td><td>Sigma</td><td>T5941</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma</td><td>M2393</td><td></td></tr><tr><td>NP-40</td><td>Sigma</td><td>74385</td><td></td></tr><tr><td>Tween-20</td><td>Fisher Scientific</td><td>BP337-100</td><td></td></tr><tr><td>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/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/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 ...

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Genetics

7.5K Views.  University of Michigan. Hier presenteren we een methode om het genoom van C. elegans ingenieur met behulp van CRISPR-Cas9 ribonucleoproteins en homologie afhankelijke reparatie sjablonen.

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

Efficient Generation of hiPSC Neural Lineage Specific Knockin Reporters Using the CRISPR/Cas9 and Cas9 Double Nickase System

Gene targeting is a critical approach for characterizing gene functions in modern biomedical research. However, the efficiency of gene targeting in human cells has been low, which prevents the generation of human cell lines at a desired rate. The past two years have witnessed a rapid progression on improving efficiency of genetic manipulation by genome editing tools such as the CRISPR (Clustered Regularly Interspaced Short Palindromic Repeats)/Cas (CRISPR-associated) system. This manuscript describes a protocol for generating lineage specific human induced pluripotent stem cell (hiPSC) reporters using CRISPR/Cas system assisted homologous recombination. Procedures for obtaining necessary components for making neural lineage reporter lines using the CRISPR/Cas system, focusing on construction of targeting vectors and single guide RNAs, are described. This protocol can be extended to platform establishment and mutation correction in hiPSCs.

Genome editing tools such as the CRISPR (Clustered Regularly Interspaced Short Palindromic Repeats)/Cas (CRISPR-associated) system have greatly improved gene targeting efficiency in human induced pluripotent stem cells (hiPSCs). This manuscript describes a protocol for generating lineage specific hiPSC reporter using CRISPR/Cas system assisted homologous recombination.

Gene targeting is a critical approach for characterizing gene functions in modern biomedical research. However, the efficiency of gene targeting in human ...

Developmental Biology

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 ...

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 ...

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

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

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

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

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

Enhanced Genome Editing with Cas9 Ribonucleoprotein in Diverse Cells and Organisms

Site-specific eukaryotic genome editing with CRISPR (clustered regularly interspaced short palindromic repeats)-Cas (CRISPR-associated) systems has quickly become a commonplace amongst researchers pursuing a wide variety of biological questions. Users most often employ the Cas9 protein derived from Streptococcus pyogenes in a complex with an easily reprogrammed guide RNA (gRNA). These components are introduced into cells, and through a base pairing with a complementary region of the double-stranded DNA (dsDNA) genome, the enzyme cleaves both strands to generate a double-strand break (DSB). Subsequent repair leads to either random insertion or deletion events (indels) or the incorporation of experimenter-provided DNA at the site of the break.
The use of a purified single-guide RNA and Cas9 protein, preassembled to form an RNP and delivered directly to cells, is a potent approach for achieving highly efficient gene editing. RNP editing particularly enhances the rate of gene insertion, an outcome that is often challenging to achieve. Compared to the delivery via a plasmid, the shorter persistence of the Cas9 RNP within the cell leads to fewer off-target events. 
Despite its advantages, many casual users of CRISPR gene editing are less familiar with this technique. To lower the barrier to entry, we outline detailed protocols for implementing the RNP strategy in a range of contexts, highlighting its distinct benefits and diverse applications. We cover editing in two types of primary human cells, T cells and hematopoietic stem/progenitor cells (HSPCs). We also show how Cas9 RNP editing enables the facile genetic manipulation of entire organisms, including the classic model roundworm Caenorhabditis elegans and the more recently introduced model crustacean, Parhyale hawaiensis.

Utilizing a preassembled Cas9 ribonucleoprotein complex (RNP) is a powerful method for precise, efficient genome editing. Here, we highlight its utility across a broad range of cells and organisms, including primary human cells and both classic and emerging model organisms.

Site-specific eukaryotic genome editing with CRISPR (clustered regularly interspaced short palindromic repeats)-Cas (CRISPR-associated) systems has ...

Generation of Defined Genomic Modifications Using CRISPR-CAS9 in Human Pluripotent Stem Cells

Human pluripotent stem cells offer a powerful system to study gene function and model specific mutations relevant to disease. The generation of precise heterozygous genetic modifications is challenging due to CRISPR-CAS9 mediated indel formation in the second allele. Here, we demonstrate a protocol to help overcome this difficulty by using two repair templates in which only one expresses the desired sequence change, while both templates contain silent mutations to prevent re-cutting and indel formation. This methodology is most advantageous for gene editing coding regions of DNA to generate isogenic control and mutant human stem cell lines for studying human disease and biology. In addition, optimization of transfection and screening methodologies have been performed to reduce labor and cost of a gene editing experiment. Overall, this protocol is widely applicable to many genome editing projects utilizing the human pluripotent stem cell model.

This protocol provides a method to facilitate the generation of defined heterozygous or homozygous nucleotide changes using CRISPR-CAS9 in human pluripotent stem cells.

Human pluripotent stem cells offer a powerful system to study gene function and model specific mutations relevant to disease. The generation of precise ...

CRISPR/Cas9 Editing of the C. elegans rbm-3.2 Gene using the dpy-10 Co-CRISPR Screening Marker and Assembled Ribonucleoprotein Complexes.

The bacterial Clustered Regularly Interspaced Short Palindromic Repeat (CRISPR)/Streptococcus pyogenes CRISPR-associated protein (Cas) system has been harnessed by researchers to study important biologically relevant problems. The unparalleled power of the CRISPR/Cas genome editing method allows researchers to precisely edit any locus of their choosing, thereby facilitating an increased understanding of gene function. Several methods for editing the C. elegans genome by CRISPR/Cas9 have been described previously. Here, we discuss and demonstrate a method which utilizes in vitro assembled ribonucleoprotein complexes and the dpy-10 co-CRISPR marker for screening. Specifically, in this article, we go through the step-by-step process of introducing premature stop codons into the C. elegans rbm-3.2 gene by homology-directed repair using this method of CRISPR/Cas9 editing. This relatively simple editing method can be used to study the function of any gene of interest and allows for the generation of homozygous-edited C. elegans by CRISPR/Cas9 editing&#160;in less than two weeks.

CRISPR/Cas9 Editing of the C. elegans rbm-3.2 Gene using the dpy-10 Co-CRISPR Screening Marker and Assembled Ribonucleoprotein Complexes.

The goal of this protocol is to enable seamless CRISPR/Cas9 editing of the C. elegans genome using assembled ribonucleoprotein complexes and the dpy-10 co-CRISPR marker for screening. This protocol can be used to make a variety of genetic modifications in C. elegans including insertions, deletions, gene replacements and codon substitutions.

The bacterial Clustered Regularly Interspaced Short Palindromic Repeat (CRISPR)/Streptococcus pyogenes CRISPR-associated protein (Cas) system has been ...

Construction of Homozygous Mutants of Migratory Locust Using CRISPR/Cas9 Technology

The migratory locust, Locusta migratoria, is not only one of the worldwide plague locusts that caused huge economic losses to human beings but also an important research model for insect metamorphosis. The CRISPR/Cas9 system can accurately locate at a specific DNA locus and cleave within the target site, efficiently introducing double-strand breaks to induce target gene knockout or integrate new gene fragments into the specific locus. CRISPR/Cas9-mediated genome editing is a powerful tool for addressing questions encountered in locust research as well as a promising technology for locust control. This study provides a systematic protocol for CRISPR/Cas9-mediated gene knockout with the complex of Cas9 protein and single guide RNAs (sgRNAs) in migratory locusts. The selection of target sites and design of sgRNA are described in detail, followed by in vitro synthesis and verification of the sgRNAs. Subsequent procedures include egg raft collection and tanned-egg separation to achieve successful microinjection with low mortality rate, egg culture,&#160;preliminary estimation of the mutation rate, locust breeding as well as detection, preservation, and passage of the mutants to ensure population stability of the edited locusts. This method can be used as a reference for CRISPR/Cas9 based gene editing applications in migratory locusts as well as in other insects.

This study provides a systematically optimized procedure of CRISPR/Cas9 ribonuclease-based construction of homozygous locust mutants as well as a detailed method for cryopreservation and resuscitation of the locust eggs.

The migratory locust, Locusta migratoria, is not only one of the worldwide plague locusts that caused huge economic losses to human beings but also an ...

Biology

Single Worm PCR: A Method to Extract and Amplify Genomic DNA

Source: Prior, H., et al. <a href="https://www.jove.com/video/57518/a-rapid-facile-pipeline-for-generating-genomic-point-mutants-c">A Rapid and Facile Pipeline for Generating Genomic Point Mutants in&#160;C. elegans&#160;Using CRISPR/Cas9 Ribonucleoproteins.</a>&#160;J. Vis. Exp.&#160;(2018).
This video describes a technique to rapidly screen a single nematode for a genetic marker by PCR screening. In the example protocol, we screen for CRISPR-based genome editing.

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

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