Thanks to Jason Wright for suggesting the Golden Gate Assembly cloning strategy and Katherine Helming and members of Orkin lab, particularly Jian Xu, Guoji Guo, Elenoe Smith, and Partha Das for helpful discussions. This work was supported by NIH R01HL032259 and P30DK049216 (Center of Excellence in Molecular Hematology) to S.H.O. and NIDDK K08DK093705 to D.E.B.

1. CRISPR Design
<ol>
	<li>Design sgRNAs manually or using freely available online tools7. Use these tools to help identify guide sequences that minimize identical genomic matches or near-matches to reduce risk of cleavage away from target sites (off-target effects). Ensure that the guide sequences consist of a 20-mer (&#8220;protospacer sequence&#8221;) upstream of an &#8220;NGG&#8221; sequence (&#8220;protospacer adjacent motif&#8221; or PAM) at the genomic recognition site. 
	NOTE: Figure 1 describes possible deletion strategies for genes and non-coding elements. For creating a gene knockout, two sgRNA located within exons will enrich even monoallelic deletion clones for loss of function. This is due to the high frequency of indels formed on non-deleted alleles12, which are likely to cause frameshift mutations leading to nonsense mediated decay of the mRNA transcript (Figure 1B).</li>
	<li>Use the example guides for the intended deletion of Pim1 in mouse (Mus musculus; Table 1, Figure 2A). 
	NOTE: In this example, sgRNA-A&#8217;s protospacer sequence and PAM happen to fall on the bottom (Crick) strand while sgRNA-B&#8217;s protospacer sequence and PAM fall on the top (Watson) strand (Figure 2A). However, DSB will occur independent of orientation of the protospacer sequence/PAM relative to the top or bottom strand.</li>
	<li>Determine the reverse complement (rc) of each guide sequence. Example reverse complement sequences of the Pim1 sgRNA from Table 1 are found in Table 2.</li>
	<li>Obtain 24- or 25-mer oligos for each guide and its associated reverse complement including additional nucleotides for cloning and expression purposes. 
	NOTE: Here we discuss the usage of the pSpCas9(BB) plasmid (pX330) (Addgene plasmid ID 42230). This plasmid allows for the simultaneous expression of sgRNA and SpCas9, but does not contain markers for selection7. Other constructs may be utilized, such as pX458 (Addgene plasmid ID 48138) or pX459 (Addgene plasmid ID 48139), which include GFP and puromycin as selectable markers, respectively, or constructs in which multiple sgRNAs may be expressed from a single plasmid.
	<ol>
		<li>Add &#8220;CACC&#8221; before the 20-mer guide sequence and &#8220;AAAC&#8221; before the guide&#8217;s reverse complement for cloning into the pX330 vector using BbsI restriction enzyme (Table 3).</li>
		<li>Add a G nucleotide after the CACC sequence and before the 20-mer if the first position of the 20-mer is not G. sgRNA expression from the U6 promoter of the pX330 vector is enhanced by the inclusion of a G nucleotide after the CACC sequence. Add a C at the 3&#8217; end of the reverse complement oligo (e.g., sgRNA-A in Table 4). The resultant oligos would be 25-mer oligos.</li>
		<li>However, if the first position of the 20-mer (protospacer sequence) is G, do not add another G (e.g., sgRNA-B in Table 4) and do not add C to the final position of the reverse complement oligo. In this case, the resultant oligos would be 24-mer oligos (Table 4).</li>
	</ol>
	</li>
</ol>
2. Design Deletion Screening Primers
<ol>
	<li>Design one set of primers internal to the sequence to be deleted (&#8220;non-deletion band&#8221;) and another set of primers upstream and downstream of the sgRNA cleavage sites (&#8220;deletion band&#8221;; Figures 1 - 2). In the absence of deletion, the &#8220;deletion band&#8221; is often too large to efficiently amplify. Typically use primers at least 100 bp from the predicted cleavage site to ensure detection would not be impacted by a small indel at the sgRNA target site.
	<ol>
		<li>Design additional primers to analyze for scarring (small indels produced at the sgRNA cleavage site without the intended deletion). Use a pair of forward and reverse primers flanking each sgRNA target site (within 150 - 350 bp) to amplify the sgRNA target site to examine for scarring. This may be useful to characterize the non-deleted allele in monoallelic deletion clones.</li>
	</ol>
	</li>
	<li>For small deletions (as the &#8220;deletion band&#8221; may still amplify), resolve amplicons on an agarose gel to determine if size is consistent with the presence or absence of deletion. For this approach, the internal primers described in step 2.1 may be omitted.</li>
</ol>
3. CRISPR Cloning
<ol>
	<li>Anneal and phosphorylate oligos.
	<ol>
		<li>Resuspend oligos at a concentration of 100 &#956;M in ddH2O.</li>
		<li>Prepare a 10 &#956;l reaction mix for each guide and its reverse complement: 1.0 &#956;l sgRNA 24- or 25-mer oligo (100 &#956;M; see step 1.4), 1.0 &#956;l sgRNA 24- or 25-mer reverse complement oligo (100 &#956;M; see step 1.4), 1.0 &#956;l 10x T4 Ligation Buffer, 6.5 &#956;l ddH2O, and 0.5 &#956;l T4 Polynucleotide Kinase (PNK) (10,000 U/ml). 
		NOTE: Phosphorylated oligos may be ordered instead. For this approach, the use of T4 PNK is omitted.</li>
		<li>Anneal in a thermocycler using the following parameters: 37 &#176;C for 30 min; 95 &#176;C for 5 min and then ramp down to 25 &#176;C at 5 &#176;C/min.</li>
		<li>Dilute oligos 1:10 in ddH2O (e.g., 1.0 &#956;l annealed oligos + 9.0 &#956;l ddH2O to yield a concentration of 1 &#956;M).</li>
	</ol>
	</li>
	<li>Ligate annealed oligos into pX330 using a Golden Gate assembly cloning strategy26.
	<ol>
		<li>Prepare a 50 &#956;l reaction mix: 100 ng circular pX330 vector, 1.0 &#956;l annealed oligos (1 &#956;M; see step 3.1.4), 5.0 &#956;l restriction enzyme buffer (10x), 4.0 &#956;l (20 U) BbsI restriction enzyme (5,000 U/ml), 5.0 &#956;l ATP (10 mM), 0.25 &#956;l (5 &#956;g) BSA (20 mg/ml), 0.375 &#956;l (750 U) T4 DNA ligase (2,000,000 U/ml), and H2O to final volume of 50 &#956;l. This reaction may be scaled down to a smaller final volume if necessary.</li>
		<li>Run samples in a thermocycler using the following parameters: Cycles 1-20 (37 &#176;C for 5 min, 20 &#176;C for 5 min); Cycle 21 (80 &#176;C for 20 min). These cycling conditions allow for digestion and ligation to occur in one reaction (see step 3.2).</li>
		<li>Transform 10 &#956;l of DH5&#945; E. coli cells with 1 &#956;l of reaction (from 3.2.1 - 3.2.2).</li>
		<li>Plate onto a lysogeny broth (LB) agar plate with 100 &#956;g/ml ampicillin and incubate O/N at 37 &#176;C.</li>
	</ol>
	</li>
	<li>Pick 2 - 3 colonies and inoculate into a mini-prep culture.
	<ol>
		<li>Perform mini-prep for each sample and sequence each colony using a U6 promoter forward primer: CGTAACTTGAAAGTATTTCGATTTCTTGGC. This is a representative sequencing primer; other flanking primers may be utilized.</li>
	</ol>
	</li>
	<li>Choose a sequence-verified colony and inoculate into a maxi-prep culture. Prep size may be scaled based on the required DNA yield.</li>
	<li>Perform maxi-prep for each CRISPR/Cas9 construct.</li>
</ol>
4. Transfecting CRISPRs into Cells of Interest
NOTE: This protocol involves the delivery of CRISPR/Cas9 plasmids by electroporation27. This protocol is described in detail for murine erythroleukemia (MEL) cells, a suspension cell line. The culture medium in all steps consists of DMEM supplemented with 2% penicillin/streptomycin and 1% L-glutamine, which is used for MEL cells. However, transient transfection of CRISPR/Cas9 plasmids may be successfully adapted to numerous cell types using preferred culture conditions and transfection strategies for each cell type. While MEL cells are suspension cells, instructions for adherent cells have also been included.
<ol>
	<li>Ensure there are 2 x 106 cells per CRISPR pair. Resuspend 2 x 106 cells in 100 &#956;l of electroporation solution and add to electroporation cuvette.
	<ol>
		<li>Add 5 &#956;g of each CRISPR/Cas9 construct (10 &#956;g total). Add 0.5 &#956;g of GFP expression construct.</li>
	</ol>
	</li>
	<li>Electroporate cells with 250 volts for 5 msec in a 2 mm cuvette using an electroporation system. 
	NOTE: Alternatively use another transfection method such as cationic liposome-based transfection. Optimize transfection conditions for each cell line with a reporter construct to ensure robust plasmid delivery before attempting genome editing.</li>
	<li>Immediately transfer solution from cuvette into 1 ml of culture media after electroporation. Minimize the time between electroporation and transferring the solution into media to enhance cell viability.</li>
	<li>Incubate at 30 - 37 &#176;C for 24 - 72 hr. 30 &#176;C may enhance genome editing efficiency, but 37 &#176;C is acceptable.</li>
</ol>
5. Fluorescence Activated Cell Sorting (FACS) of Transfected Cells
<ol>
	<li>Prepare cells for FACS by filtering them through a 50 &#956;m filter into a FACS tube.</li>
	<li>FACS sort the top ~3% of GFP positive cells in order to enrich for cells that received high levels of the CRISPR/Cas9 constructs.</li>
	<li>Plate sorted cells individually into 96-well round-bottom plates using sorter or by using limiting dilution at 30 cells per 96-well round-bottom plate. Optimize plating the cell type used at limiting dilution prior to performing this step to reliably obtain approximately 30 cells per 96-well plate.</li>
	<li>Include 100 &#956;l per well of cell culture media.</li>
	<li>For the remaining sorted cells (&#8220;bulk&#8221;) that were not plated, freeze half of the cells for future plating. Plate the other half for screening and primer validation (see step 6). 
	NOTE: This protocol is for suspension cells. Adherent cells can either grow as individual cells in 96-well flat bottom plate or in a 10 cm dish at low concentration so that individual single-cell derived clones can be picked and moved to a flat bottom 96-well plate.</li>
	<li>Allow the bulk cells to incubate at 37 &#176;C for 3 - 7 days and allow the clones to incubate at 37 &#176;C for 7 - 14 days. Vary these times depending on the doubling time of the cell line used. 
	NOTE: This incubation time allows for sufficient cell proliferation for screening genomic DNA (gDNA) for the intended deletion by PCR (see steps 6.1 and 7.1). The bulk cells have sufficiently proliferated when the concentration exceeds ~100,000 cells/ml or for adherent cells, the cells have reached ~80% confluence. The clones have sufficiently proliferated once macroscopically visible with ~2 mm diameter.</li>
</ol>
6. Primer Validation and Screening for CRISPR/Cas9-Mediated Deletion
<ol>
	<li>Isolate gDNA from parental and bulk sorted cells by resuspending parental and bulk cell pellets in 50 &#956;l of DNA extraction solution. 
	NOTE: Generally ~100,000 cells are used for DNA extraction, although a wide range of cell numbers is acceptable. The bulk sorted cells are composed of a polyclonal population exposed to sgRNA-A and sgRNA-B (see step 5). The purpose of the following PCR is to validate primers and verify the presence of intended genomic deletion.</li>
	<li>Run sample in thermocycler and run the following program: 65 &#176;C for 6 min, 98 &#176;C for 2 min to extract gDNA. Measure the DNA concentration. 
	NOTE: While steps 6.1 and 6.2 recommend an efficient method for DNA extraction, any method for genomic DNA isolation may be utilized to be able to perform PCR in step 6.3.</li>
	<li>Assemble a 20 &#956;l PCR with the following components: 10 &#956;l 2x PCR mix, 0.5 &#956;l forward primer (10 &#956;M), 0.5 &#956;l reverse primer (10 &#956;M), 50-100 ng gDNA, and H2O up to 20 &#956;l. Use the primers designed in step 2 above. Conduct PCR for &#8220;non-deletion band&#8221; and &#8220;deletion band&#8221; in separate reactions. 
	NOTE: Numerous polymerases may be used for step 6.3.
	<ol>
		<li>Run samples in a thermocycler using the following parameters: 95 &#176;C for 15 min, 35 cycles of (95 &#176;C for 30 sec, 60 &#176;C for 1 min, 72 &#176;C for 1 min), and 72 &#176;C for 10 min. Optimize PCR conditions for each primer pair designed based on testing the bulk sorted cells.</li>
	</ol>
	</li>
	<li>Run samples on 2% agarose gel at 10 V/cm using 1x Tris-acetate-EDTA (TAE) buffer.</li>
	<li>Examine samples for the presence/absence of non-deletion and deletion bands (Figure 2). Consider multiplexing the &#8220;deletion&#8221; and &#8220;non-deletion&#8221; PCR primer pairs in a single reaction. Optimize multiplexing in a polyclonal population (i.e., bulk sorted cells) before screening individual clones. It is critical that the deletion and non-deletion amplicons be easily resolved on an agarose gel for multiplexing.</li>
</ol>
7. Screening CRISPR/Cas9 Clones for Deletions and Clone Selection
<ol>
	<li>For suspension cells, transfer all clones to a single 96-well plate that already contains 50 &#956;l cell culture media per well for a final volume of 150 &#956;l. This facilitates screening by allowing a multichannel pipette to be used for the remainder of the steps in step 7.
	<ol>
		<li>Transfer 50 &#956;l from each well (leaving 100 &#956;l in each well) to a 96-well PCR plate using a multichannel pipette.</li>
		<li>Centrifuge PCR plate at 400 x g for 5 min and remove supernatant by flicking the PCR plate over a sink. Add 50 &#956;l of DNA extraction solution per well and resuspend. Continue to step 7.3 for suspension cells.</li>
	</ol>
	</li>
	<li>For adherent cells, aspirate media. Add 20 &#956;l of 0.05% trypsin-EDTA to each well with a clone present.
	<ol>
		<li>Resuspend cells in 200 &#956;l of media. Pipette mix to detach cells.</li>
		<li>Plate 100 &#956;l each into two separate 96-well flat-bottom plates. Keep one plate to allow for clones to grow and use the other plate to screen each clone for deletions.</li>
		<li>Add an additional 100 &#956;l to each well for a total volume of 200 &#956;l. Wait 24 - 72 hr to allow cells to grow.</li>
		<li>Aspirate media. Add 50 &#956;l DNA extraction solution per well, resuspend and transfer to 96-well PCR plate. Continue to step 7.3.</li>
	</ol>
	</li>
	<li>Extract the gDNA from clones. Run sample in thermocycler: 65 &#176;C for 6 min and 98 &#176;C for 2 min to extract gDNA.</li>
	<li>Screen each clone using the same PCR primers and reaction conditions optimized on the bulk cells (see step 6).</li>
	<li>Select the clones identified with the desired deletion and move to larger plate or flask for growth.</li>
</ol>
8. Validation of Biallelic Deletion Clones
<ol>
	<li>In order to characterize obtained clones and validate a successful knockout, evaluate clones at the DNA as well as RNA and/or protein levels.
	<ol>
		<li>To evaluate the DNA, amplify deletion bands from biallelic deletion clones with a proofreading polymerase and clone the amplicons (e.g., with a PCR cloning kit) into a plasmid vector. Transform the plasmid into DH5&#945; E. coli cells and plate onto LB agar plates with the relevant antibiotic. Select multiple colonies, mini-prep each one, and subject each clone to Sanger sequencing to characterize each deletion allele28&#8211;30. Repeating the PCR test for deletion after the initial screen ensures that the correct clone was selected and reproducibility of results.</li>
		<li>To evaluate the RNA, perform RT-qPCR for gene expression of the relevant gene30,31.</li>
		<li>To evaluate the protein, perform an immunoblot using an antibody against the relevant protein33.</li>
	</ol>
	</li>
</ol>

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D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Purifying+Plasmid+DNA+from+Bacterial+Colonies+Using+the+Qiagen+Miniprep+Kit.">Purifying Plasmid DNA from Bacterial Colonies Using the Qiagen Miniprep Kit.</a> J. Vis. Exp. (6), e247 (2007).</li><li>Froger, A., Hall, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transformation+of+Plasmid+DNA+into+E.+coli+Using+the+Heat+Shock+Method.">Transformation of Plasmid DNA into E. coli Using the Heat Shock Method.</a> J. Vis. Exp. (6), e253 (2007).</li><li>Finney, M., Nisson, P. E., Rashtchian, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Molecular+cloning+of+PCR+products.">Molecular cloning of PCR products.</a> Current Protocols in Molecular Biology. Chapter 15. (Unit 15.4), (2001).</li><li>Gordanpour, A., Nam, R. K., Sugar, L., Bacopulos, S., Seth, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=MicroRNA+Detection+in+Prostate+Tumors+by+Quantitative+Real-time+PCR+(qPCR).">MicroRNA Detection in Prostate Tumors by Quantitative Real-time PCR (qPCR).</a> J. Vis. Exp. (63), e3874 (2012).</li><li>Schularick, N. M., Clark, J. J., Hansen, M. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Primary+Culture+of+Human+Vestibular+Schwannomas.">Primary Culture of Human Vestibular Schwannomas.</a> J. Vis. Exp. (89), e51093 (2014).</li><li>Eslami, A., Lujan, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Western+Blotting:+Sample+Preparation+to+Detection.">Western Blotting: Sample Preparation to Detection.</a> J. Vis. Exp. (44), e2359 (2010).</li><li>McVey, M., Lee, S. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=MMEJ+repair+of+double-strand+breaks+(director&#8217;s+cut):+deleted+sequences+and+alternative+endings.">MMEJ repair of double-strand breaks (director&#8217;s cut): deleted sequences and alternative endings.</a> Trends in Genetics. 24 (11), 529-538 (2008).</li><li>Symington, L. S., Gautier, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Double-strand+break+end+resection+and+repair+pathway+choice.">Double-strand break end resection and repair pathway choice.</a> Annual Review of Genetics. 45, 247-271 (2011).</li><li>Fu, Y., 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=High-frequency+off-target+mutagenesis+induced+by+CRISPR-Cas+nucleases+in+human+cells.">High-frequency off-target mutagenesis induced by CRISPR-Cas nucleases in human cells.</a> Nature Biotechnology. 31 (9), 822-826 (2013).</li><li>Wu, X., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genome-wide+binding+of+the+CRISPR+endonuclease+Cas9+in+mammalian+cells.">Genome-wide binding of the CRISPR endonuclease Cas9 in mammalian cells.</a> Nature Biotechnology. , (2014).</li><li>Hsu, P. 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=DNA+targeting+specificity+of+RNA-guided+Cas9+nucleases.">DNA targeting specificity of RNA-guided Cas9 nucleases.</a> Nature Biotechnology. 31 (9), 827-832 (2013).</li><li>Fu, Y., 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=Improving+CRISPR-Cas+nuclease+specificity+using+truncated+guide+RNAs.">Improving CRISPR-Cas nuclease specificity using truncated guide RNAs.</a> Nature Biotechnology. 32, 279-284 (2014).</li><li>Alem&#225;n, L. M., Doench, J., Sharp, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=a+Comparison+of+siRNA-induced+off-target+RNA+and+protein+effects.">a Comparison of siRNA-induced off-target RNA and protein effects.</a> RNA. 13 (3), 385-395 (2007).</li><li>Anderson, E. 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=Experimental+validation+of+the+importance+of+seed+complement+frequency+to+siRNA+specificity+Experimental+validation+of+the+importance+of+seed+complement+frequency+to+siRNA+specificity.">Experimental validation of the importance of seed complement frequency to siRNA specificity Experimental validation of the importance of seed complement frequency to siRNA specificity.</a> RNA. 14, 853-861 (2008).</li><li>Marine, S., 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=Common+seed+analysis+to+identify+off-target+effects+in+siRNA+screens.">Common seed analysis to identify off-target effects in siRNA screens.</a> Journal of Biomolecular Screening. 17 (3), 370-378 (2012).</li></ol>

We have no conflicts of interest related to this report.

The CRISPR/Cas9 system may be used to generate genomic deletions of a range of sizes. Although we have observed that the frequency of deletion varies inversely with respect to intended deletion size, we have been able to recover deletions of up to 1 Mb, and deletions up to 100 kb routinely yield multiple biallelic deleted clones. We have observed no loss in efficiency of sequentially introducing deletions into a cell line. This strategy can be used for creation of combinatorial deletion of numerous genes and elements. The process of obtaining biallelic deletion clones can be expedited by estimating the minimum number of clones needed to be screened based on deletion size to obtain the desired number of clones with biallelic deletion12.
The ability to obtain monoallelic deletion with the absence of biallelic deletion at probabilistic distribution could indicate cell lethality associated with complete loss of function. Low frequency or absent deletions could reflect a number of scenarios including poor transfection, inefficient sgRNAs or inefficient PCR screening primers (due to lack of a positive control to validate PCR primers to screen for deletion). GFP+ cells can be used as a surrogate for transfection efficiency (see step 5.2), so a reduction in GFP+ cells likely reflects poor transfection and a resulting decreased deletion efficiency. Using two different sgRNA pairs with independent screening primers can help control for inefficient sgRNA and screening PCR primers and maximize chances of obtaining biallelic deletion clones. Cell sorting for GFP+ cells enriches for deletion alleles. While this step may be omitted, omission will likely necessitate screening more clones to identify those with monoallelic or biallelic deletions. To the degree that transfection efficiency may be optimized, we would expect genome editing efficiency to be enhanced.
The NHEJ events that underlie deletions and local repair result in a series of alleles with a variety of indels at the target sites. The predominant outcome is small ~1 - 10 bp insertions or more commonly deletions at the site of sgRNA-directed cleavage (Figure 2B). Often these alleles appear to be the result of microhomology-based repair34,35. It should be noted that the PCR-based detection strategy we describe will not identify larger or more complicated insertions, deletions, inversions, or rearrangements. Although these events are less common, we have observed clones in which neither deletion nor non-deletion amplicons could be detected, and upon further investigation reflect these more complex outcomes.
We have observed extensive CRISPR/Cas9-mediated &#8220;scarring&#8221; of non-deletion alleles from monoallelic and non-deletion clones (see Figure 2B). These &#8220;scars&#8221; consist of small indels produced at the sgRNA cleavage site without the intended deletion (i.e., deletion of the intervening segment between sgRNAs A and B). These scars often interrupt target recognition by the sgRNA. Therefore we would urge caution in retargeting alleles in cells previously exposed to sgRNAs using the same sgRNAs. A more successful retargeting strategy would utilize unique sgRNA sequences distinct from previously &#8220;scarred&#8221; recognition sites. In cases when a pair of sgRNAs recognizes exonic sequences (Figure 1B, bottom), frameshift alleles may be produced even in the absence of deletion. Therefore, monoallelic deletion clones can be enriched for loss-of-function due to the high frequency of frameshift mutations on the non-deleted allele12.
One concern with the CRISPR/Cas9 system is off-target effects, i.e., genomic modification at unintended sites36&#8211;38. Recent reports have suggested that shorter guide RNAs with 17 - 19 nucleotides can reduce the frequency of CRISPR/Cas9-based off-target effects39. Additionally, a double-nicking strategy using two guides per target site with a nickase can be used to create DSBs while minimizing off-target effects7. Alternatively, analogous to strategies used for RNAi, we suggest that different pairs of sgRNAs with non-overlapping protospacer sequences be used to demonstrate that the observed phenotype is the result of the on-target CRISPR/Cas9 modification as opposed to a potential off-target effect. A convenient approach would be to design at least two adjacent but non-overlapping sgRNA pairs so that a single set of screening primers (see step 2) may be used for multiple sgRNA pairs (Figure 1A). Furthermore, complementing a deletion cell line by reintroducing the missing sequence and/or disrupted gene can substantiate a causal relationship between a given genomic deletion and phenotype.
For biologists working with cellular model systems, RNAi has represented a powerful tool for functional genomics. However, limitations of this approach have included incomplete reduction in target mRNA transcript levels, heterogeneity of effect of independent reagents targeting the same gene, and known off-target effects including seed-based and non-seed effects40&#8211;42. Genome editing strategies promise to address many of these concerns and represent an exciting, complementary approach for prospective genetic perturbation8,36,37. Furthermore, genome editing allows for the study of non-coding genetic elements in a way not possible by RNAi and challenging by conventional targeting approaches25. We encourage generation of genomic deletions by CRISPR/Cas9 as a robust and specific method to produce and characterize loss-of-function alleles.

Recent advances in genome engineering technology have allowed for unprecedented opportunities for site-specific modification of the genome. This technology may be utilized to investigate the function of genes and regulatory elements via prospective genetic perturbation. Zinc finger nucleases (ZFNs), transcription-activator like (TAL) effector nucleases (TALENs), and CRISPR/Cas9 RNA-guided nucleases each leverage customizable DNA specificity to localize a nuclease for the introduction of DSBs1&#8211;3. The resulting DSBs can be repaired by indel-forming NHEJ or by homology-directed repair (HDR) using a donor template4.
The CRISPR/Cas9 nuclease pathway, an adaptive immune system in prokaryotic cells5, has been recently adapted for mammalian genome engineering2,3. This tool has been demonstrated to be an inexpensive, efficient, and reliable genome engineering technique6. Briefly, a complex of Streptococcus pyogenes-derived Cas9 nuclease and a sgRNA achieve target recognition via Watson-Crick base-pairing with cognate genomic DNA sequences. sgRNAs include 20-mer sequences complementary to genomic sequences adjacent to an obligate protospacer adjacent motif (PAM) NGG. Cas9 induces a DSB at a predictable position within the target site. Additionally, variants of Cas9 with single-strand cleavage capacity or catalytic inactivity may be used to facilitate &#8220;nicking&#8221; or transcriptional regulation respectively7&#8211;9. CRISPR/Cas9 has been used for a wide range of applications including both knock-in and knockout10,11, large-scale genomic deletions12&#8211;14, pooled library screening for gene discovery15,16, genetic engineering of numerous model organisms10,11,17&#8211;21, as well as gene therapy22,23.
Here we describe a protocol for efficient deletion of desired genomic regions. The protocol includes CRISPR design, cloning, and delivery, as well as deletion, identification, and characterization. Genomic deletions can be generated by the introduction of two CRISPR sgRNAs with Cas9 to induce repair of the resultant two DSBs by NHEJ with deletion of the intervening segment. This strategy has been used to create deletions ranging from one kilobase to over one megabase12. Deletions can be informative for the study of genes and other genetic elements, either in isolation or in combination. There are several potential advantages of genomic deletions as compared to HDR or single-site small indel production. First, this method capitalizes on the high efficiency of NHEJ in many cellular contexts7. The high frequency of deletion limits the number of clones needed to be screened to identify informative clones. Deletion frequency is inversely related to deletion size. Biallelic deletion clones may be retrieved at frequencies at least as great as of probabilistic expectation12. Second, both monoallelic and biallelic deletions may be easily identified and distinguished by conventional PCR, simplifying the screening process. Strategies relying on small indels or point mutations may require RFLP, allele-specific PCR, T7EN1 cleavage assay, Sanger sequencing, RT-qPCR, or immunoblotting, which may be more laborious. Third, by removing a substantial portion of a gene or element of interest, a reliable loss-of-function allele may be obtained. In contrast, frameshift mutations in protein-coding sequences may not always induce nonsense-mediated decay, may produce a hypomorphic or neomorphic allele, or target an exon excluded from an alternate isoform24. Finally, deletions may be particularly revealing for the study of non-coding DNA such as regulatory elements since frameshift mutations as produced by single-site indels would not be relevant25.

<table border="0" cellpadding="0" cellspacing="0" width="842">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:388px;">Name of Material/ Equipment</td>
			<td style="width:168px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td style="width:167px;">Comments/Description</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">T4 Polynucleotide Kinase</td>
			<td>New England Biolabs</td>
			<td>M0201S</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">T4 DNA Ligase (with associated ligation buffer)</td>
			<td>New England Biolabs</td>
			<td>M0202T</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Adenosine 5&#39;-Triphosphate (ATP)</td>
			<td>New England Biolabs</td>
			<td>P0756S</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">BSA, Molecular Biology Grade</td>
			<td>New England Biolabs</td>
			<td>B9000S</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">BbsI Restriction Enzyme (with associated NEB Buffer 2.1)</td>
			<td>New England Biolabs</td>
			<td>R0539S</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">pSpCas9(BB) (pX330)</td>
			<td>Addgene</td>
			<td>42230</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">S.O.C. Medium</td>
			<td>Life Technologies</td>
			<td>15544-034</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">BTX ECM 830</td>
			<td>Harvard Apparatus</td>
			<td>45-0052</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">BTX Solution and 2mm Cuvettes</td>
			<td>Harvard Apparatus</td>
			<td>45-0803</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">pmaxGFP Plasmid</td>
			<td>Lonza</td>
			<td>VPA-1003</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">QuickExtract DNA Extraction Solution</td>
			<td>Epicentre</td>
			<td>QE09050</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">HotStarTaq PCR Master Mix Kit</td>
			<td>Qiagen</td>
			<td>203443</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Zero Blunt PCR Cloning Kit</td>
			<td>Life Technologies</td>
			<td>K2700-20</td>
			<td></td>
		</tr>
	</tbody>
</table>

generation of genomic deletions in mammalian cell lines via crispr/cas9

The prokaryotic clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated (Cas) 9 system may be re-purposed for site-specific eukaryotic genome engineering. CRISPR/Cas9 is an inexpensive, facile, and efficient genome editing tool that allows genetic perturbation of genes and genetic elements. Here we present a simple methodology for CRISPR design, cloning, and delivery for the production of genomic deletions. In addition, we describe techniques for deletion, identification, and characterization. This strategy relies on cellular delivery of a pair of chimeric single guide RNAs (sgRNAs) to create two double strand breaks (DSBs) at a locus in order to delete the intervening DNA segment by non-homologous end joining (NHEJ) repair. Deletions have potential advantages as compared to single-site small indels given the efficiency of biallelic modification, ease of rapid identification by PCR, predictability of loss-of-function, and utility for the study of non-coding elements. This approach can be used for efficient loss-of-function studies of genes and genetic elements in mammalian cell lines.

The goal of this experiment was the deletion of Pim1 in MEL cells. Use of multiple non-overlapping sgRNA pairs (i.e., independent protospacer sequences) may help to control for off-target effects (Figure 1A). A consistent phenotype would be more likely to result from an on-target effect as opposed to a common off-target effect shared by multiple independent protospacer sequences. Each pair would lead to production of a unique deletion breakpoint. If close together (i.e., n less than ~150 bp), the same screening primers could be used to detect deletions produced by each set of sgRNAs. Genomic deletions may disrupt genes by using sgRNA pairs in various locations with respect to the gene (Figure 1B). For example, the sgRNA pair may flank a gene for deletion of the entire gene body; the pair could be located within two exons, with the potential to create frameshift indels even if one or both alleles were not deleted; or the pair may flank a specific exon to allow disruption of a particular isoform.
The deletion strategy used for Pim1 was to design flanking sgRNAs to delete the entire gene body, an 8 kb deletion (Figure 2). This strategy was chosen in part due to the relatively small size of the Pim1 gene. This example shows one PAM (green) on the top (Watson) strand and one PAM on the bottom (Crick) strand; however, DSB is independent of PAM sequence localization to the top or bottom strand. sgRNA pairs can have both PAM sequences on the top strand, both on the bottom strand, or one of each. Using the protocol described above, two sgRNA were designed, cloned into the pX330 expression vector, and delivered to MEL cells by electroporation along with a GFP reporter (Figure 2A). The top 3% of GFP+ cells were sorted two days post-electroporation and plated clonally at limiting dilution. Screening primers were designed as described in step 2 and as shown in Figure 2. PCR conditions were optimized using gDNA isolated from parental MEL cells and from &#8220;bulk&#8221; sorted cells.
10 days after plating, gDNA was isolated from all clones and screened for deletion via PCR, which identified non-deletion, monoallelic and biallelic deletion clones according to the patterns of non-deletion (ND) and deletion (D) amplicons (Figure 3). Non-deletion clones were identified as having the presence of the non-deletion amplicon and the absence of the deletion amplicon. Monoallelic clones were identified as having the presence of both the non-deletion and deletion amplicons. Biallelic clones were identified as having the absence of the non-deletion amplicon and the presence of the deletion amplicon. For this deletion, 400 clones were screened which identified 126 monoallelic deletion clones and 32 biallelic deletion clones (it is important to note that deletion frequency varies with deletion size12). Biallelic deletion clones were selected and moved to flasks with 8 ml of media. After allowing 5 days for expansion, each clone was retested by PCR of gDNA to confirm biallelic deletion and deletion amplicons were subjected to Sanger sequencing to identify the precise deletion (Figure 2B). Heterogeneity within the deletion amplicons reflects imperfect indel-forming NHEJ repair. Sequencing of the non-deletion allele in monoallelelic deletion clones uncovered indels in the majority of cases, demonstrating that even the non-deleted allele is frequently edited by CRISPR/Cas9, which may be important for applications where both monoallelic and biallelic deletion clones are required. RNA was isolated from biallelic deletion clones and analyzed by RT-qPCR to confirm loss of Pim1 expression (Figure 4).
<img alt="Figure 1" src="/files/ftp_upload/52118/52118fig1highres.jpg" /> 
Figure 1. Schematic of possible deletion strategies. (A) Two example sgRNA pairs for genomic deletions (shown in black and orange, respectively). The blue arrows indicate primers to detect the non-deletion amplicon and red arrows indicate primers to detect the deletion amplicon. sgRNA positions 17 and 18 are highlighted in red and blue at sgRNA-Sites-A1/A2 and are highlighted in purple and orange at sgRNA-Sites-B1/B2 with a red line indicating the predicted Cas9 cleavage between positions 17 and 18. (B) CRISPR-directed cleavages are shown as vertical black lines. The blue arrows indicate primers to detect the non-deletion amplicon and red arrows indicate primers to detect the deletion amplicon. <a href="https://www.jove.com/files/ftp_upload/52118/52118fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/52118/52118fig2highres.jpg" /> 
Figure 2. Strategy to produce and detect deletion of Pim1, including sequencingdeletion and non-deletion alleles. (A) Schematic of the PCR-based screening strategy to identify Pim1 deletion clones. One primer pair is located internal to the deletion (blue arrows) and one primer pair is localized external to the deletion (red arrows). sgRNA sequences (protospacer sequences) are shown in purple. The vertical red lines indicate the predicted Cas9 cleavage between positions 17 and 18 of the sgRNA sequence. (B) Sanger sequencing reveals indel formation at the sgRNA recognition site. sgRNA sequences are shown in purple and PAM sequences in green. Deletion events are shown by an equivalent number of dash marks and insertions are highlighted in blue. Vertical red lines indicate predicted cleavage site, between positions 17 and 18 of the sgRNA. <a href="https://www.jove.com/files/ftp_upload/52118/52118fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/52118/52118fig3highres.jpg" /> 
Figure 3. Representative gel identifying non-deletion, monoallelic, and biallelic clones. For each clone, the left lane represents the non-deletion amplicon (&#8220;ND&#8221; in blue) and the right lane represents the deletion amplicon (&#8220;D&#8221; in red). Individual clones are separated by dotted lines. The three monoallelic deletion clones are clones 5, 6, and 2. The three biallelic deletion clones are clones 15, 17, and 13. As seen in the sequencing data, the deletion band for clone 13 has a smaller size due to larger deletions at the deletion junction.
<img alt="Figure 4" src="/files/ftp_upload/52118/52118fig4highres.jpg" /> 
Figure 4. Loss of Pim1 expression in biallelic deletion clones. Pim1 expression was calculated for two biallelic deletion clones by RT-qPCR. Data was normalized to Gapdh using the 2-&#916;Ct method.
<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td></td>
			<td>Protospacer Sequence</td>
		</tr>
		<tr>
			<td>sgRNA-A</td>
			<td>5&#8217;-TAGGAAAATGACTCGTCACT-3&#8217;</td>
		</tr>
		<tr>
			<td>sgRNA-B</td>
			<td>5&#8217;-GTCTATCCTATCTCGATAAA-3&#8217;</td>
		</tr>
	</tbody>
</table>
Table 1. 20-mer protospacer sequences for two sgRNA for the deletion of Pim1.
<table border="0" cellpadding="0" cellspacing="0" width="543">
	<tbody>
		<tr height="23">
			<td height="23" style="height:23px;width:101px;"></td>
			<td style="width:443px;">Reverse Complement of Protospacer Sequence</td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;">sgRNA-A-rc</td>
			<td>5&#8217;-AGTGACGAGTCATTTTCCTA-3&#8217;</td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;">sgRNA-B-rc</td>
			<td>5&#8217;-TTTATCGAGATAGGATAGAC-3&#8217;</td>
		</tr>
	</tbody>
</table>
Table 2. Reverse complement of the protospacer sequences for the two sgRNA from Table 1.
<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td></td>
			<td>Sequences</td>
		</tr>
		<tr>
			<td>sgRNA-A</td>
			<td>5&#8217;-CACCTAGGAAAATGACTCGTCACT-3&#8217;</td>
		</tr>
		<tr>
			<td>sgRNA-A-rc</td>
			<td>5&#8217;-AAACAGTGACGAGTCATTTTCCTA-3&#8217;</td>
		</tr>
		<tr>
			<td>sgRNA-B</td>
			<td>5&#8217;-CACCGTCTATCCTATCTCGATAAA-3&#8217;</td>
		</tr>
		<tr>
			<td>sgRNA-B-rc</td>
			<td>5&#8217;-AAACTTTATCGAGATAGGATAGAC-3&#8217;</td>
		</tr>
	</tbody>
</table>
Table 3. Protospacer sequences and their reverse complements with &#8220;CACC&#8221; and &#8220;AAAC&#8221; added for cloning into the pX330 vector using BbsI restriction enzyme.
<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td></td>
			<td>Sequences</td>
		</tr>
		<tr>
			<td>sgRNA-A</td>
			<td>5&#8217;- CACCGTAGGAAAATGACTCGTCACT-3&#8217;</td>
		</tr>
		<tr>
			<td>sgRNA-A-rc</td>
			<td>5&#8217;- AAACAGTGACGAGTCATTTTCCTAC-3&#8217;</td>
		</tr>
		<tr>
			<td>sgRNA-B</td>
			<td>5&#8217;- CACCGTCTATCCTATCTCGATAAA-3&#8217;</td>
		</tr>
		<tr>
			<td>sgRNA-B-rc</td>
			<td>5&#8217;- AAACTTTATCGAGATAGGATAGAC-3&#8217;</td>
		</tr>
	</tbody>
</table>
Table 4. sgRNA expression from the U6 promoter of the pX330 vector is enhanced by the addition of a G nucleotide after the CACC sequence and before the 20-mer. The addition of an extra G nucleotide requires the addition of a C nucleotide at the 3&#8217; end of the reverse complement oligo (e.g., sgRNA-A). However, if the first position of the 20-mer (protospacer sequence) is already a G nucleotide, there is no need to add another G (e.g., sgRNA-B) and no need to add C to the final position of the reverse complement oligo.

Watch this Scientific Journal Video about Generation of Genomic Deletions in Mammalian Cell Lines via CRISPR/Cas9 at JoVE.com

Generation of Genomic Deletions in Mammalian Cell Lines via CRISPR/Cas9

CRISPR/Cas9 is a robust system to produce disruption of genes and genetic elements. Here we describe a protocol for the efficient creation of genomic deletions in mammalian cell lines using CRISPR/Cas9.

The prokaryotic clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated (Cas) 9 system may be re-purposed for site-specific ...

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94.8K Views.  Harvard Medical School. CRISPR/Cas9 is a robust system to produce disruption of genes and genetic elements. Here we describe a protocol for the efficient creation of genomic deletions in mammalian cell lines using CRISPR/Cas9.

Video: Generation of Genomic Deletions in Mammalian Cell Lines via CRISPR/Cas9

MISSION esiRNA for RNAi Screening in Mammalian Cells

RNA interference (RNAi) is a basic cellular mechanism for the control of gene expression. RNAi is induced by short double-stranded RNAs also known as small interfering RNAs (siRNAs). The short double-stranded RNAs originate from longer double stranded precursors by the activity of Dicer, a protein of the RNase III family of endonucleases. The resulting fragments are components of the RNA-induced silencing complex (RISC), directing it to the cognate target mRNA. RISC cleaves the target mRNA thereby reducing the expression of the encoded protein1,2,3. RNAi has become a powerful and widely used experimental method for loss of gene function studies in mammalian cells utilizing small interfering RNAs.
Currently two main methods are available for the production of small interfering RNAs. One method involves chemical synthesis, whereas an alternative method employs endonucleolytic cleavage of target specific long double-stranded RNAs by RNase III in vitro. Thereby, a diverse pool of siRNA-like oligonucleotides is produced which is also known as endoribonuclease-prepared siRNA or esiRNA. A comparison of efficacy of chemically derived siRNAs and esiRNAs shows that both triggers are potent in target-gene silencing. Differences can, however, be seen when comparing specificity. Many single chemically synthesized siRNAs produce prominent off-target effects, whereas the complex mixture inherent in esiRNAs leads to a more specific knockdown10.
In this study, we present the design of genome-scale MISSION esiRNA libraries and its utilization for RNAi screening exemplified by a DNA-content screen for the identification of genes involved in cell cycle progression. We show how to optimize the transfection protocol and the assay for screening in high throughput. We also demonstrate how large data-sets can be evaluated statistically and present methods to validate primary hits. Finally, we give potential starting points for further functional characterizations of validated hits.

Here we use a human esiRNA library in a high-throughput screen for genes involved in cell division. We demonstrate how to set up and conduct an esiRNA screens, as well as how to analyze and validate the results.

RNA interference (RNAi) is a basic cellular mechanism for the control of gene expression. RNAi is induced by short double-stranded RNAs also known as ...

The Green Monster Process for the Generation of Yeast Strains Carrying Multiple Gene Deletions

Phenotypes for a gene deletion are often revealed only when the mutation is tested in a particular genetic background or environmental condition1,2. There are examples where many genes need to be deleted to unmask hidden gene functions3,4. Despite the potential for important discoveries, genetic interactions involving three or more genes are largely unexplored. Exhaustive searches of multi-mutant interactions would be impractical due to the sheer number of possible combinations of deletions. However, studies of selected sets of genes, such as sets of paralogs with a greater a priori chance of sharing a common function, would be informative.
In the yeast Saccharomyces cerevisiae, gene knockout is accomplished by replacing a gene with a selectable marker via homologous recombination. Because the number of markers is limited, methods have been developed for removing and reusing the same marker5,6,7,8,9,10. However, sequentially engineering multiple mutations using these methods is time-consuming because the time required scales linearly with the number of deletions to be generated.
Here we describe the Green Monster method for routinely engineering multiple deletions in yeast11. In this method, a green fluorescent protein (GFP) reporter integrated into deletions is used to quantitatively label strains according to the number of deletions contained in each strain (Figure 1). Repeated rounds of assortment of GFP-marked deletions via yeast mating and meiosis coupled with flow-cytometric enrichment of strains carrying more of these deletions lead to the accumulation of deletions in strains (Figure 2). Performing multiple processes in parallel, with each process incorporating one or more deletions per round, reduces the time required for strain construction.
The first step is to prepare haploid single-mutants termed 'ProMonsters,' each of which carries a GFP reporter in a deleted locus and one of the 'toolkit' loci&#x2014;either Green Monster GMToolkit-a or GMToolkit-&alpha; at the can1&Delta; locus (Figure 3). Using strains from the yeast deletion collection12, GFP-marked deletions can be conveniently generated by replacing the common KanMX4 cassette existing in these strains with a universal GFP-URA3 fragment. Each GMToolkit contains: either the a- or &alpha;-mating-type-specific haploid selection marker1 and exactly one of the two markers that, when both GMToolkits are present, collectively allow for selection of diploids.
The second step is to carry out the sexual cycling through which deletion loci can be combined within a single cell by the random assortment and/or meiotic recombination that accompanies each cycle of mating and sporulation.

The Green Monster method enables the rapid assembly of multiple deletions marked with a reporter gene encoding green fluorescent protein. This method is based on driving yeast strains through repeated cycles of sexual assortment of deletions and fluorescence-based enrichment of cells carrying more deletions.

Phenotypes  for a gene deletion are often revealed only when the mutation is tested in a  particular genetic background or environmental condition1,2. ...

Generation of Stable Human Cell Lines with Tetracycline-inducible (Tet-on) shRNA or cDNA Expression

A major approach in the field of mammalian cell biology is the manipulation of the expression of genes of interest in selected cell lines, with the aim to reveal one or several of the gene's function(s) using transient/stable overexpression or knockdown of the gene of interest. Unfortunately, for various cell biological investigations this approach is unsuitable when manipulations of gene expression result in cell growth/proliferation defects or unwanted cell differentiation. Therefore, researchers have adapted the Tetracycline repressor protein (TetR), taken from the E. coli tetracycline resistance operon1, to generate very efficient and tight regulatory systems to express cDNAs in mammalian cells2,3. In short, TetR has been modified to either (1) block initiation of transcription by binding to the Tet-operator (TO) in the promoter region upon addition of tetracycline (termed Tet-off system) or (2) bind to the TO in the absence of tetracycline (termed Tet-on system) (Figure 1). Given the inconvenience that the Tet-off system requires the continuous presence of tetracycline (which has a half-life of about 24 hr in tissue cell culture medium) the Tet-on system has been more extensively optimized, resulting in the development of very tight and efficient vector systems for cDNA expression as used here.
Shortly after establishment of RNA interference (RNAi) for gene knockdown in mammalian cells4, vectors expressing short-hairpin RNAs (shRNAs) were described that function very similar to siRNAs5-11. However, these shRNA-mediated knockdown approaches have the same limitation as conventional knockout strategies, since stable depletion is not feasible when gene targets are essential for cellular survival. To overcome this limitation, van de Wetering et al.12 modified the shRNA expression vector pSUPER5 by inserting a TO in the promoter region, which enabled them to generate stable cell lines with tetracycline-inducible depletion of their target genes of interest.
Here, we describe a method to efficiently generate stable human Tet-on cell lines that reliably drive either inducible overexpression or depletion of the gene of interest. Using this method, we have successfully generated Tet-on cell lines which significantly facilitated the analysis of the MST/hMOB/NDR cascade in centrosome13,14 and apoptosis signaling15,16. In this report, we describe our vectors of choice, in addition to describing the two consecutive manipulation steps that are necessary to efficiently generate human Tet-on cell lines (Figure 2). Moreover, besides outlining a protocol for the generation of human Tet-on cell lines, we will discuss critical aspects regarding the technical procedures and the characterization of Tet-on cells.

Generation of Stable Human Cell Lines with Tetracycline-inducible Tet-on shRNA or cDNA Expression

A rapid and simple way to generate human cell lines with inducible and reversible cDNA overexpression or shRNA-mediated knock-down of the gene of interest. This method enables researchers to reliably and highly reproducibly manipulate cell lines that are difficult to alter by transient transfection methods or conventional knockdown/knockout strategies.

A major approach in  the field of mammalian cell biology is the manipulation of the expression of  genes of interest in selected cell lines, with the aim ...

Anaerobic Growth and Maintenance of Mammalian Cell Lines

Most mucosal surfaces along with the midpoints in tumors and stem cell niches are geographic areas of the body that are anoxic. Previous studies show that the incubation in normoxic (5% CO2 in air) or hypoxic (low oxygen levels) conditions followed by an anoxic incubation (an absence of free oxygen) results in limited viability (2&#8211;3 days). A novel methodology was developed that enables an anoxic cell cultivation (for at least 17 days; the maximum time tested). The most critical aspect of this methodology is to ensure that no oxygen is introduced into the system. This is obtained by the degassing of media, and by flushing tubes, dishes, flasks, and pipettes with an anaerobic gas mixture (H2, CO2, N2) followed by permitting the materials to equilibrate to the anoxic (non-oxygen) environment prior to usage.&#160;Additional care must be exercised when acquiring photomicrographs to ensure that the micrographs obtained do not contain artifacts. In the absence of oxygen, cell morphology is significantly altered. Two distinct morphotypes are noted for all anaerobically-grown cells. The ability to grow and maintain mammalian cells in the absence of oxygen can be applied to the analysis of cell physiology, polymicrobial interactions, and the characterization of biosynthetic pathways for novel cancer drug development.

Here, we describe a new method that enables the anaerobic long-term cultivation of established cell lines. The maximum survival time that was tested is 17 days. This method is suitable for the testing of cytotoxic agents and exploring the physiology of anoxically replicating cells.

Most mucosal surfaces along with the midpoints in tumors and stem cell niches are geographic areas of the body that are anoxic. Previous studies show that ...

Silencing the Spark: CRISPR/Cas9 Genome Editing in Weakly Electric Fish

Electroreception and electrogenesis have changed in the evolutionary history of vertebrates. There is a striking degree of convergence in these independently derived phenotypes, which share a common genetic architecture. This is perhaps best exemplified by the numerous convergent features of gymnotiforms and mormyrids, two species-rich teleost clades that produce and detect weak electric fields and are called weakly electric fish. In the 50 years since the discovery that weakly electric fish use electricity to sense their surroundings and communicate, a growing community of scientists has gained tremendous insights into evolution of development, systems and circuits neuroscience, cellular physiology, ecology, evolutionary biology, and behavior. More recently, there has been a proliferation of genomic resources for electric fish. Use of these resources has already facilitated important insights with regards to the connection between genotype and phenotype in these species. A major obstacle to integrating genomics data with phenotypic data of weakly electric fish is a present lack of functional genomics tools. We report here a full protocol for performing CRISPR/Cas9 mutagenesis that utilizes endogenous DNA repair mechanisms in weakly electric fish. We demonstrate that this protocol is equally effective in both the mormyrid species Brienomyrus brachyistius and the gymnotiform Brachyhypopomus gauderio by using CRISPR/Cas9 to target indels and point mutations in the first exon of the sodium channel gene scn4aa. Using this protocol, embryos from both species were obtained and genotyped to confirm that the predicted mutations in the first exon of the sodium channel scn4aa were present. The knock-out success phenotype was confirmed with recordings showing reduced electric organ discharge amplitudes when compared to uninjected size-matched controls.

Here, a protocol is presented to produce and rear CRISPR/Cas9 genome knockout electric fish. Outlined in detail are the required molecular biology, breeding, and husbandry requirements for both a gymnotiform and a mormyrid, and injection techniques to produce Cas9-induced indel F0 larvae.

Electroreception and electrogenesis have changed in the evolutionary history of vertebrates. There is a striking degree of convergence in these ...

Genome Engineering of Primary Human B Cells Using CRISPR/Cas9

B cells are lymphocytes derived from hematopoietic stem cells and are a key component of the humoral arm of the adaptive immune system. They make attractive candidates for cell-based therapies because of their ease of isolation from peripheral blood, their ability to expand in vitro, and their longevity in vivo. Additionally, their normal biological function&#8212;to produce large amounts of antibodies&#8212;can be utilized to express very large amounts of a therapeutic protein, such as a recombinant antibody to fight infection, or an enzyme for the treatment of enzymopathies. Here, we provide detailed methods for isolating primary human B cells from peripheral blood mononuclear cells (PBMCs) and activating/expanding isolated B cells in vitro. We then demonstrate the steps involved in using the CRISPR/Cas9 system for site-specific KO of endogenous genes in B cells. This method allows for efficient KO of various genes, which can be used to study the biological functions of genes of interest. We then demonstrate the steps for using the CRISPR/Cas9 system together with a recombinant, adeno-associated, viral (rAAV) vector for efficient site-specific integration of a transgene expression cassette in B cells. Together, this protocol provides a step-by-step engineering platform that can be used in primary human B cells to study biological functions of genes as well as for the development of B-cell therapeutics.

Here we provide a detailed, step-by-step protocol for CRISPR/Cas9-based genome engineering of primary human B cells for gene knockout (KO) and knock-in (KI) to study biological functions of genes in B cells and the development of B-cell therapeutics.

B cells are lymphocytes derived from hematopoietic stem cells and are a key component of the humoral arm of the adaptive immune system. They make ...

TGF-&#946;-mediated Endothelial to Mesenchymal Transition (EndMT) and the Functional Assessment of EndMT Effectors using CRISPR/Cas9 Gene Editing

In response to specific external cues and the activation of certain transcription factors, endothelial cells can differentiate into a mesenchymal-like phenotype, a process that is termed endothelial to mesenchymal transition (EndMT). Emerging results have suggested that EndMT is causally linked to multiple human diseases, such as fibrosis and cancer. In addition, endothelial-derived mesenchymal cells may be applied in tissue regeneration procedures, as they can be further differentiated into various cell types (e.g., osteoblasts and chondrocytes). Thus, the selective manipulation of EndMT may have clinical potential. Like epithelial-mesenchymal transition (EMT), EndMT can be strongly induced by the secreted cytokine transforming growth factor-beta (TGF-&#946;), which stimulates the expression of so-called EndMT transcription factors (EndMT-TFs), including Snail and Slug. These EndMT-TFs then up- and downregulate the levels of mesenchymal and endothelial proteins, respectively. Here, we describe methods to investigate TGF-&#946;-induced EndMT in vitro, including a protocol to study the role of particular TFs in TGF-&#946;-induced EndMT. Using these techniques, we provide evidence that TGF-&#946;2 stimulates EndMT in murine pancreatic microvascular endothelial cells (MS-1 cells), and that the genetic depletion of Snail using clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein 9 (Cas9)-mediated gene editing, abrogates this phenomenon. This approach may serve as a model to interrogate potential modulators of endothelial biology, and can be used to perform genetic or pharmacological screens in order to identify novel regulators of EndMT, with potential application in human disease.

TGF-&amp;#946;-mediated Endothelial to Mesenchymal Transition EndMT and the Functional Assessment of EndMT Effectors using CRISPR/Cas9 Gene Editing

We describe methods to investigate TGF-&#946;2-induced EndMT in endothelial cells by observing cell morphology changes and examining the expression EndMT-related marker changes using immunofluorescence staining. CRISPR/Cas9 gene editing was described and used to deplete the gene encoding Snail to investigate its role in TGF-&#946;2-induced EndMT.

In response to specific external cues and the activation of certain transcription factors, endothelial cells can differentiate into a mesenchymal-like ...

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 ...

CRISPR/Cas9 Gene Editing of Hematopoietic Stem and Progenitor Cells for Gene Therapy Applications

CRISPR/Cas9 is a highly versatile and efficient gene-editing tool adopted widely to correct various genetic mutations. The feasibility of gene manipulation of hematopoietic stem and progenitor cells (HSPCs) in vitro makes HSPCs an ideal target cell for gene therapy. However, HSPCs moderately lose their engraftment and multilineage repopulation potential in ex vivo culture. In the present study, ideal culture conditions are described that improves HSPC engraftment and generate an increased number of gene-modified cells in vivo. The current report displays optimized in vitro culture conditions, including the type of culture media, unique small molecule cocktail supplementation, cytokine concentration, cell culture plates, and culture density. In addition to that, an optimized HSPC gene-editing procedure, along with the validation of the gene-editing events, are provided. For in vivo validation, the gene-edited HSPCs infusion and post-engraftment analysis in mouse recipients are displayed. The results demonstrated that the culture system increased the frequency of functional HSCs in vitro, resulting in robust engraftment of gene-edited cells in vivo.

The present protocol describes an optimized hematopoietic stem and progenitor cell (HSPC) culture procedure for the robust engraftment of gene-edited cells in vivo.

CRISPR/Cas9 is a highly versatile and efficient gene-editing tool adopted widely to correct various genetic mutations. The feasibility of gene ...

CRISPR-Cas9-Mediated Precise Knock-In Edits in Zebrafish Hearts

Clustered regularly interspaced short palindromic repeats (CRISPR) in animal models enable precise genetic manipulation for the study of physiological phenomena. Zebrafish have been used as an effective genetic model to study numerous questions related to heritable disease, development, and toxicology at the whole-organ and -organism level. Due to the well-annotated and mapped zebrafish genome, numerous tools for gene editing have been developed. However, the efficacy of generating and ease of detecting precise knock-in edits using CRISPR is a limiting factor. Described here is a CRISPR-Cas9-based knock-in approach with the simple detection of precise edits in a gene responsible for cardiac repolarization and associated with the electrical disorder, Long QT Syndrome (LQTS). This two-single-guide RNA (sgRNA) approach excises and replaces the target sequence and links a genetically encoded reporter gene. The utility of this approach is demonstrated by describing non-invasive phenotypic measurements of cardiac electrical function in wild-type and gene-edited zebrafish larvae. This approach enables the efficient study of disease-associated variants in a whole organism. Furthermore, this strategy offers possibilities for the insertion of exogenous sequences of choice, such as reporter genes, orthologs, or gene editors.

This protocol describes an approach to facilitate precise knock-in edits in zebrafish embryos using CRISPR-Cas9 technology. A phenotyping pipeline is presented to demonstrate the applicability of these techniques to model a Long QT Syndrome-associated gene variant.

Clustered regularly interspaced short palindromic repeats (CRISPR) in animal models enable precise genetic manipulation for the study of physiological ...