CRISPR: Protocol for Genomic Deletions in Mammalian Cell Lines
Addgene is proud to present a video reprint from the Journal of Visualized Experiments (JOVE). The video publication by Stuart Orkin's and Daniel Bauer's labs details the use of CRISPR/Cas9 to create genomic deletions in mammalian cell lines. Along with the video, you can find the protocol section from the original publication below. You can also download the full-text of the publication here.
Article CitationGeneration of Genomic Deletions in Mammalian Cell Lines via CRISPR/Cas9. Bauer DE, Canver MC, Orkin SH. J. Vis. Exp. (95), e52118, doi:10.3791/52118 (2015). PubMed
Full Access to the Paper & Protocol VideoDownload the manuscript here: Bauer et al, 2014 (322.2 KB)
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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.
For referenced Tables and Figures, please access the manuscript PDF posted at the top of the page.
- Design sgRNAs manually or using freely available online tools1. 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 (“protospacer sequence”) upstream of an “NGG” sequence (“protospacer adjacent motif” 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 alleles2, which are likely to cause frameshift mutations leading to nonsense mediated decay of the mRNA transcript (Figure 1B).
- Use the example guides for the intended deletion of Pim1 in mouse (Mus musculus; Table 1, Figure 2A).
NOTE: In this example, sgRNA-A’s protospacer sequence and PAM happen to fall on the bottom (Crick) strand while sgRNA-B’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.
- Determine the reverse complement of each guide sequence. Example reverse complement sequences of the Pim1 sgRNA from Table 1 are found in Table 2.
- 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 selection1. 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.
- Add “CACC” before the 20-mer guide sequence and “AAAC” before the guide’s reverse complement for cloning into the pX330 vector using BbsI restriction enzyme (Table 3).
- 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’ end of the reverse complement oligo (e.g., sgRNA-A in Table 4). The resultant oligos would be 25-mer oligos.
- 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).
Design Deletion Screening Primers
- Design one set of primers internal to the sequence to be deleted (“non-deletion band”) and another set of primers upstream and downstream of the sgRNA cleavage sites (“deletion band”; Figures 1 - 2). In the absence of deletion, the “deletion band” 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.
- 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.
- For small deletions (as the “deletion band” may still amplify), resolve 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.
- Anneal and phosphorylate oligos.
- Resuspend oligos at a concentration of 100 μM in ddH2O.
- Prepare a 10 μl reaction mix for each guide and its reverse complement: 1.0 μl sgRNA 24- or 25-mer oligo (100 μM; see step 1.4), 1.0 μl sgRNA 24- or 25-mer reverse complement oligo (100 μM; see step 1.4), 1.0 μl 10x T4 Ligation Buffer, 6.5 μl ddH2O, and 0.5 μl 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.
- Anneal in a thermocycler using the following parameters: 37 °C for 30 min; 95 °C for 5 min and then ramp down to 25 °C at 5 °C/min.
- Dilute oligos 1:10 in ddH2O (e.g., 1.0 μl annealed oligos + 9.0 μl ddH2O to yield a concentration of 1 μM).
- Ligate annealed oligos into pX330 using a Golden Gate assembly cloning strategy3.
- Prepare a 50 μl reaction mix: 100 ng circular pX330 vector, 1.0 μl annealed oligos (1 μM; see step 3.1.4), 5.0 μl restriction enzyme buffer (10x), 4.0 μl (20 U) BbsI restriction enzyme (5,000 U/ml), 5.0 μl ATP (10 mM), 0.25 μl (5 μg) BSA (20 mg/ml), 0.375 μl (750 U) T4 DNA ligase(2,000,000 U/ml), and H2O to final volume of 50 μl. This reaction may be scaled down to a smaller final volume if necessary.
- Run samples in a thermocycler using the following parameters: Cycles 1-20 (37 °C for 5 min, 20 °C for 5 min); Cycle 21 (80 °C for 20 min). These cycling conditions allow for digestion and ligation to occur in one reaction (see step 3.2).
- Transform 10 μl of DH5α E. coli cells with 1 μl of reaction (from 3.2.1 - 3.2.2).
- Plate onto a lysogeny broth (LB) agar plate with 100 μg/ml ampicillin and incubate O/N at 37 °C.
- Pick 2 - 3 colonies and inoculate into a mini-prep culture.
- 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.
- Choose a sequence-verified colony and inoculate into a maxi-prep culture. Prep size may be scaled based on the required DNA yield.
- Perform maxi-prep for each CRISPR/Cas9 construct.
Transfecting CRISPRs into Cells of Interest
- Ensure there are 2 x 106 cells per CRISPR pair. Resuspend 2 x 106 cells in 100 μl of electroporation solution and add to electroporation cuvette.
- Add 5 μg of each CRISPR/Cas9 construct (10 μg total). Add 0.5 μg of GFP expression construct.
- 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.
- 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.
- Incubate at 30 - 37 °C for 24 - 72 hr. 30 °C may enhance genome editing efficiency, but 37 °C is acceptable.
Fluorescence Activated Cell Sorting (FACS) of Transfected Cells
- Prepare cells for FACS by filtering them through a 50 μm filter into a FACS tube.
- FACS sort the top ~3% of GFP positive cells in order to enrich for cells that received high levels of the CRISPR/Cas9 constructs.
- 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.
- Include 100 μl per well of cell culture media.
- For the remaining sorted cells (“bulk”) 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.
- Allow the bulk cells to incubate at 37 °C for 3 - 7 days and allow the clones to incubate at 37 °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.
Primer Validation and Screening for CRISPR/Cas9-Mediated Deletion
- Isolate gDNA from parental and bulk sorted cells by resuspending parental and bulk cell pellets in 50 μl 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.
- Run sample in thermocycler and run the following program: 65 °C for 6 min, 98 °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.
- Assemble a 20 μl PCR with the following components: 10 μl 2x PCR mix, 0.5 μl forward primer (10 μM), 0.5 μl reverse primer (10 μM), 50-100 ng gDNA, and H2O up to 20 μl. Use the primers designed in step 2 above. Conduct PCR for “non-deletion band” and “deletion band” in separate reactions.
NOTE: Numerous polymerases may be used for step 6.3.
- Run samples in a thermocycler using the following parameters: 95 °C for 15 min, 35 cycles of (95 °C for 30 sec, 60 °C for 1 min, 72 °C for 1 min), and 72 °C for 10 min. Optimize PCR conditions for each primer pair designed based on testing the bulk sorted cells.
- Run samples on 2% agarose gel at 10 V/cm using 1x Tris-acetate-EDTA (TAE) buffer.
- Examine samples for the presence/absence of non-deletion and deletion bands (Figure 2). Consider multiplexing the “deletion” and “non-deletion” 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.
Screening CRISPR/Cas9 Clones for Deletions and Clone Selection
- For suspension cells, transfer all clones to a single 96-well plate that already contains 50 μl cell culture media per well for a final volume of 150 μl. This facilitates screening by allowing a multichannel pipette to be used for the remainder of the steps in step 7.
- Transfer 50 μl from each well (leaving 100 μl in each well) to a 96-well PCR plate using a multichannel pipette.
- Centrifuge PCR plate at 400 x g for 5 min and remove supernatant by flicking the PCR plate over a sink. Add 50 μl of DNA extraction solution per well and resuspend. Continue to step 7.3 for suspension cells.
- For adherent cells, aspirate media. Add 20 μl of 0.05% trypsin-EDTA to each well with a clone present.
- Resuspend cells in 200 μl of media. Pipette mix to detach cells.
- Plate 100 μl 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.
- Add an additional 100 μl to each well for a total volume of 200 μl. Wait 24 - 72 hr to allow cells to grow.
- Aspirate media. Add 50 μl DNA extraction solution per well, resuspend and transfer to 96-well PCR plate. Continue to step 7.3.
- Extract the gDNA from clones. Run sample in thermocycler: 65 °C for 6 min and 98 °C for 2 min to extract gDNA.
- Screen each clone using the same PCR primers and reaction conditions optimized on the bulk cells (see step 6).
- Select the clones identified with the desired deletion and move to larger plate or flask for growth.
Validation of Biallelic Deletion Clones
- In order to characterize obtained clones and validate a successful knockout, evaluate clones at the DNA as well as RNA and/or protein levels.
- 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 DH5α 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 allele5–7. Repeating the PCR test for deletion after the initial screen ensures that the correct clone was selected and reproducibility of results.
- To evaluate the RNA, perform RT-qPCR for gene expression of the relevant gene7,8.
- To evaluate the protein, perform an immunoblot using an antibody against the relevant protein9.
NOTE: This protocol involves the delivery of CRISPR/Cas9 plasmids by electroporation4. 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.
ReferencesThis protocol is excerpted from: Generation of Genomic Deletions in Mammalian Cell Lines via CRISPR/Cas9. Bauer DE, Canver MC, Orkin SH. J. Vis. Exp. (95), e52118, doi:10.3791/52118 (2015). PubMed
For the complete publication, including representative results and discussion, download the manuscript here: Bauer et al, 2014 (322.2 KB)
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