CRISPR Guide

Basic Mechanisms of CRISPR

The requirements for a CRISPR knockout assay are simple. You’ll need a Cas enzyme and a gRNA specific to the gene of interest. The genomic target of the gRNA can be any ~20 nucleotide sequence, provided it meets two conditions:

• The sequence is unique compared to the rest of the genome
• The target is present immediately adjacent to a Protospacer Adjacent Motif (PAM)

The PAM sequence (NGG) serves as a binding signal for Cas9, but the exact sequence depends on which Cas protein you use, and there are several to choose from. Many variations of Cas9 exist, along with other Cas enzymes like Cas12a, which are available for specialized applications. Check out our list of additional Cas proteins and PAM sequences.

Once you’ve selected your target, you’ll express your gRNA and Cas9 protein, which will form a ribonucleoprotein complex through interactions between the gRNA scaffold and surface-exposed positively-charged grooves on Cas9. gRNA binding induces a conformational change in Cas9, shifting it into an active, DNA-binding configuration, with the spacer region of the gRNA left free to interact with the target DNA.

Cas9 will only cleave a given locus if the gRNA spacer sequence shares sufficient homology with the target DNA. Once the Cas9-gRNA complex binds a putative DNA target, the seed sequence (8–10 bases at the 3′ end of the gRNA targeting sequence) will begin to anneal to the target DNA. If the seed and target DNA sequences match, the gRNA will continue to anneal to the target DNA in a 3′ to 5′ direction. The location of any potential mismatches matters. Mismatches between the target sequence in the 3′ seed sequence inhibit target cleavage, while mismatches toward the 5′ end distal to the PAM often permit target cleavage.

Generating a Knockout Using CRISPR

Cas9 undergoes a second conformational change upon target binding. In this formation, its nuclease domains, RuvC and HNH, cleave opposite strands of the target DNA. This results in a double-strand break (DSB) within the target DNA, located ∼3–4 nucleotides upstream of the PAM sequence.

The resulting DSB is then repaired by one of two general repair pathways: the efficient but error-prone non-homologous end joining (NHEJ) pathway and the less efficient but high-fidelity homology directed repair (HDR) pathway. HDR relies on copying DNA from a matching template to accurately repair or fill in the missing sequence, and is more commonly used for small precision edits. NHEJ directly ligates the break ends without the need for a homologous template. Both NHEJ and HDR have their uses, but we’ll focus on NHEJ first and come back to HDR.

NHEJ is a naturally occurring repair pathway and is the primary repair mechanism for DSBs. While this pathway is efficient in repairing DSB, it frequently causes small nucleotide insertions or deletions (indels) at the DSB site. Therefore, a population of cells expressing Cas9 and a gRNA will result in a diverse array of mutations (see: Validate genetic modification). In most cases, NHEJ gives rise to small indels in the target DNA that result in amino acid deletions, insertions, or frameshift mutations leading to premature stop codons within the open reading frame (ORF) of the targeted gene. The ideal result is a loss-of-function mutation within the targeted gene. However, the strength of the knockout phenotype for a given mutant cell must be validated experimentally.

Browse Plasmids: Double-Strand Break (Cut)

Multiplex Genome Engineering

Many CRISPR experiments involve editing more than one gene. In these cases, delivering multiple gRNAs using a single plasmid ensures that all gRNAs will be expressed in the same cell. This is called multiplexing, and increases the likelihood that any cell containing the CRISPR plasmid will have all desired genomic edits carried out by Cas9. Multiplexing applications include editing multiple genes at once; using dual nickases to generate a knockout or gene edit; or using Cas9 to generate large genomic deletions or inversions with two gRNA sites on the same chromosome.

Most multiplex systems enable researchers to target anywhere from 2–7 genetic loci by cloning multiple gRNAs into a single plasmid — though some have achieved targeting in the double digits. These multiplex gRNA vectors can be used to knock out, activate, or repress target genes when paired with the appropriate Cas enzyme. Specific enzymes, such as Cas12a or Cas13, can improve multiplexing efficiency and are discussed in more detail later. Read more about Cas9 multiplexing.

Browse Plasmids: Multiplex gRNA Vectors

Engineering the Cas9 Endonuclease

CRISPR specificity is determined by both the gRNA sequence and the Cas9 enzyme. Ideally, a gRNA targeting sequence will have perfect homology to the target DNA with no homology elsewhere in the genome. Realistically, most gRNA targeting sequences will have additional sites of partial homology throughout the genome, called off-targets, that can impact your experiment. There are many online tools available to help you select an optimized gRNA for your application (see: Plan Your Experiment).

In addition to an optimized gRNA design, engineered Cas9s can enhance CRISPR specificity. SpCas9 (from Streptococcus pyogenes) is the most popular Cas endonuclease, with many modified versions engineered to increase specificity and nuclease activity available. Engineered Cas9s have also led to reduced dependency on specific PAM sequences and the creation of new tools.

Cas9 Orthologs

While SpCas9 is the most commonly used Cas enzyme, additional Cas9 orthologs are available. SaCas9 (from Staphylococcus aureus) has a coding sequence ~1 kb shorter than SpCas9 while retaining the same basic function. Its size allows it to be efficiently packaged into adeno-associated viruses (AAVs).

Other natural Cas orthologs include Hsp1Cas9 (from Helicobacter sp. MIT 11-5569) and Nme2Cas9 (from Neisseria meningitidis), which are about the same size as SaCas9, with higher fidelity than SpCas9. FrCas9 (from Faecalibaculum rodentium) has increased fidelity, and recognizes NNTA, a PAM sequence often found in plants, expanding the targeting range for plant genome engineering.

Read more about Cas9 variants.

Precise Modifications Using Homology Directed Repair (HDR)

HDR generates specific genomic edits by copying from a DNA donor template. The donor template contains the desired sequence and is delivered to the cell along with the gRNA(s) and Cas enzyme. The donor DNA contains the desired edit as well as additional DNA matching the sequence immediately upstream and downstream of the target site (termed left and right homology arms). The length of each homology arm depends on the size of the change being introduced; larger insertions require longer homology arms.

Depending on the application, the donor DNA may be a single-stranded oligonucleotide, double-stranded linear DNA, or a double-stranded DNA plasmid. Small ssDNA oligos often work well for small edits like a single-point mutation, while whole plasmid or linear dsDNA templates work better for large edits or insertions. When designing the donor template, incorporating mutations that prevent further Cas9 cleavage will improve the accuracy and efficiency of the desired edit. For example, you should alter the PAM site or target sequence in your donor DNA with a silent mutation that does not change the amino acid sequence.

HDR efficiency is generally low (< 10% of modified alleles). Selecting a target cut site as close as possible to the desired edit can increase efficiency. Synchronizing cells or modulating the cell cycle can help as HDR takes place during the S and G2 phases. Chemically or genetically inhibiting genes involved in NHEJ or upregulating those in the HDR pathway can also help. Additionally, constructs with Cas9 fused to proteins that modulate HDR or NHEJ components, like miCas9, Cas9-DN1S, or others, can increase HDR efficiency.

In most applications, a large portion of the Cas9-induced DSBs will still be repaired via NHEJ. The resulting population of cells will contain some combination of wild type alleles, NHEJ-repaired alleles, and the desired HDR-edited allele. Therefore, it is important to confirm the presence of the desired edit experimentally and to isolate clones containing the desired edit (see: Validate genetic modification).

Browse Plasmids: Endogenous Tagging

CRISPR Base Editing

To overcome low HDR efficiency, researchers have developed base editors to specifically convert one base to another within a target locus of about 8 bp, specified by a gRNA. There are two main types of base edits:

• Base transitions - edit to same classification of nucleotide base (purine-to-purine or pyrimidine-to-pyrimidine)
• Base transversions - edit to different classification of nucleotide base (purine-to-pyrimidine and vice versa)

The first classes of base editors were cytosine base editors (purine-to-purine) and adenine base editors (pyrimidine-to-pyrimidine), which catalyze base transitions. Cytosine base editors are created by fusing Cas9 nickase (Cas9n) or dCas9 to a cytidine deaminase like APOBEC. Within the region of single-stranded DNA created by Cas9 binding, the cytidine deaminase converts cytidine to uridine, which is read as thymidine during subsequent DNA repair, creating a C to T change. Modern base editors include several fused components that promote the intended repair pathway or inhibit unwanted ones and improve editing efficiency.

Likewise, adenine base editors have been engineered to convert adenosine to inosine, which is treated like guanosine by the cell, creating an A to G (or T to C) change. Adenosine DNA deaminases do not exist in nature, but have been created by directed evolution of the Escherichia coli TadA, a tRNA adenosine deaminase. Like cytosine base editors, the evolved TadA domain is fused to Cas9n or dCas9 to create the adenine base editor.

The TadA domain has also been further evolved to catalyze cytidine deamination instead of (or in addition to) adenosine deamination, producing cytosine base editors that are smaller and less likely to cause off-target mutations than those derived from naturally occurring cytidine deaminases.

More recently, researchers have developed base editors that can catalyze base transversions. These editors are based on cytosine base editors and similarly work by first converting cytidine to uridine. However, they promote the base excision repair pathway, which fully removes the uridine from the DNA and replaces it with another base. In E. coli, this strategy yields C to A base edits, whereas in mammalian cells, it produces C to G edits.

Adenine transversion editors soon followed, using an enzyme evolved to excise inosine to access the same repair pathway and convert A to any other base. In principle, the ability to induce base transversions means that any base-to-base conversion is possible with base editing.

While many base editors are designed to work in a very narrow window proximal to the PAM sequence, some base editing systems create a wide spectrum of single-nucleotide variants (somatic hypermutation) in a larger editing window. These systems are well-suited to directed evolution applications. Examples of these base editing systems include targeted AID-mediated mutagenesis (TAM) and CRISPR-X, in which Cas9 is fused to activation-induced cytidine deaminase (AID).

Base editors are available with multiple Cas9 variants, including high fidelity Cas9s, Cas9s with various PAM requirements, and other Cas enzymes like Cas13. Specialized variants of the editing enzymes have been developed to reduce off-target mutations in DNA, RNA, or both, and are recommended in contexts where such mutations would be problematic.

Browse Plasmids: Base Edit

CRISPR Transposases

Transposon systems like piggyBac or Tn7 allow for large scale edits by integrating segments of DNA flanked by specific repeat elements into the genome. CRISPR-associated transposases (CASTs, developed by the Feng Zhang lab) combine the large-scale capacity of transposases with the precise DNA-targeting of CRISPR-Cas systems. You may also see them referred to as INTEGRATE (Insertion of Transposable Elements by Guide RNA-Assisted Targeting), developed by Samuel H. Sternberg’s lab. Many of these systems are based on naturally occurring CRISPR-associated Tn7-like transposases found in diverse species of microbes. However, synthetic dCas9-transposase fusions have also been developed.

The two main classes of CAST systems are type I and type V-K. Type I systems use a type I Cas effector, composed of several separate Cas enzymes that form a complex called Cascade (see Cas3 for more details), while type V-K systems use a type V-K Cas effector, Cas12k. No matter the system, the Cas effectors lack nuclease activity. Each system also includes Tn7-like transposase components. Type I (generally) includes tnsA, tnsB, tnsC, and tniQ. In comparison, type V-K includes just tnsB, tnsC, and tniQ. The CRISPR-Cas complex recruits the transposase proteins to the target location in the genome, where the transposase then carries out integration of donor DNA. Both types of CASTs have been found to achieve donor DNA integration for sequences > 10 kb!

Currently, these systems perform best in bacterial species, but newer versions engineered for eukaryotic cells are available and further improvements are actively in progress. Type I CASTs can result in bidirectional insertions — meaning they may not be suitable for applications where orientation of the insert is important. Type V-K CASTs appear to exhibit almost unidirectional insertions but are more prone to off-target insertions. And in the case of both types, the transposon end sequences are integrated along with the sequence, so the edit is not considered “scarless”.

Browse Plasmids: CRISPR transposases

What Are Pooled Lentiviral CRISPR Libraries?

Pooled lentiviral CRISPR libraries are a heterogeneous population of lentiviral transfer vectors. In most cases, each plasmid contains an individual gRNA targeting a single gene in a given genome. Some more recent libraries use arrays of multiple gRNAs to target paralogs or lncRNAs.

Guide RNAs are designed in silico and synthesized (Figure 8A), then cloned in a pooled format into lentiviral transfer vectors (Figure 8B). Libraries come in 1-vector systems, in which Cas9 is included in the gRNA-containing plasmid, or 2-vector systems, in which Cas9 must be delivered separately. They can knockout, activate, or repress genes.

Each library typically contains ∼3–6 gRNAs per gene to ensure modification of every target gene, so CRISPR libraries contain thousands of unique gRNAs. gRNA design for CRISPR libraries is usually optimized to select for gRNAs with high on-target activity and low off-target activity. Knockout libraries often target 5′ constitutively expressed exons, while activation and repression libraries target promoter or enhancer regions. As each library is unique, be sure to check the library information and original publication to see if a library is suitable for your experiment.

Although lentiviral libraries are the most popular method for CRISPR screening, they are not suitable for all cell types or experiments. Mammalian CRISPR libraries have also been created in AAV backbones for in vivo experiments and in a retroviral backbone for delivery to cells that are poorly transduced by lentivirus. Non-mammalian CRISPR libraries are also available, such as those for yeast or drosophila. Although CRISPR has been less widely used in bacteria due to technical challenges, bacterial CRISPR libraries have been developed for inhibition using dCas9.

Activation or Repression of Target Genes

Targeting dCas9 to transcription start sites is sufficient to repress transcription by blocking initiation. More robust transcriptional repression (CRISPR interference, or CRISPRi) or activation (CRISPRa) of downstream target genes can be achieved by fusing dCas9 to transcriptional repressors or activators and targeting promoter regions. You might sometimes see these referred to together as CRISPRi/a.

dCas9-mediated gene activation or repression is reversible, as it does not permanently modify the genomic DNA. dCas9 has also been used for genome-wide screens to activate or repress gene expression in mice and human cells.

The simplest dCas9-based activators and repressors consist of dCas9 fused directly to a single transcriptional activator (e.g. VP64) or repressor (e.g. KRAB; Figure 9A). Other examples of activation systems to target genes in mammalian cells include:

• SunTag system - co-expression of epitope-tagged dCas9 and antibody-activator effector proteins; long-term imaging of proteins in living cells (Figure 9B)
• dCas9-VPR - dCas9 fused to several different activation domains in series; a single fusion protein, instead of a two-component system (Figure 9C)
• SAM activators - co-expression of dCas9-VP64 with a modified scaffold gRNA and additional RNA-binding helper activators; highest levels of single-gene activation (Figure 9D)

In bacteria, activating gene expression is more difficult because of the absence of effective gene activators when fused with dCas9. Recently, synthetic CRISPR-Cas gene activators have been developed for bacteria by using a scaffold RNA that contains the gRNA and an RNA hairpin to recruit activation proteins.

CRISPRi has also been adapted for diverse bacterial species using a modular system called Mobile-CRISPRi. In this system, CRISPRi is introduced into bacteria using conjugation and stably integrated into the chromosome.

Browse Plasmids: Activate, Repress/Interfere

Epigenetic Modifications Using CRISPR

A variation of CRISPRi/a involves dCas9 fused to epigenetic modifiers. These programmable epigenome engineering tools alter gene expression by modifying the methylation state of cytosines in a gene’s promoter or by inducing histone acetylation or demethylation. The most common modifiers include:

• P300 - activation through increased H3K27 acetylation
• TET1 - activation through cytosine demethylation
• DNMT3A - repression through cytosine methylation
• MQ1 - repression through cytosine methylation with improved kinetics
• LSD1 - repression through targeted removal of H3K4 and H3K9 methylation

In certain cases, epigenetic modifiers may be a better choice than other activators/repressors due to their persistence and heritability. Not all modifications are stable across cell divisions, and some types of modifications are more frequently inherited by daughter cells. To maintain activation or repression, many CRISPR epigenetic tools require constitutive expression of the dCas9 fusion and/or expression of multiple fusions. The effects of CRISPRi/a may be reversible once the effector is inactivated or removed from the system.

CRISPRoff, developed by the Luke Gilbert and Jonathan Weissman labs, is an all-in-one dCas9 fusion with KRAB, DNMT3A, and DNMT3L. CRISPRoff maintains gene silencing across much of the human genome following transient expression of the dCas9 fusion, with the epigenetic modifications persisting through many generations and even cell differentiation. The system is reversible, with a companion CRISPRon to undo repression using a TET1-dCas9 fusion.

Browse Plasmids: Epigenetics

Visualizing Genomic Loci Using Fluorophores

CRISPR imaging has numerous advantages, including the simplicity of gRNA design, programmability for different genomic loci, and the capability to detect multiple targets simultaneously. CRISPR imaging can visualize chromatin and RNA dynamics in live cells.

Labeling can be done by fusing catalytically inactive dCas9 to a fluorescent marker like green fluorescent protein (GFP), creating a customizable DNA or RNA label for fluorescence microscopy. RNA aptamers, like MS2 or PP7, can also be fused to the gRNA to recruit fluorescently-tagged RNA-binding proteins (RBPs).

Multicolor CRISPR imaging can be achieved by using orthogonal dCas9s (e.g. S. pyogenes dCas9 and S. aureus dCas9) tagged with different fluorescent proteins or by fusing the gRNAs to orthogonal protein-interacting RNA aptamers, such as in the CRISPRainbow kit. Using multiple copies of the aptamer improves stability and amplifies the signal for better imaging of genomic loci, as in the CRISPR-Sirius.

Fluorescent CRISPR systems have been used for dynamic tracking of repetitive and non-repetitive genomic loci, chromosome painting, and RNA tracking in living cells. Visualizing a specific genomic locus requires recruitment of many copies of labeled proteins to the site. Chromosome-specific repetitive loci can be efficiently visualized in living cells using a single gRNA that has multiple target sequences in close proximity. A non-repetitive genomic locus can also be labeled by co-delivering multiple gRNAs that tile the locus. Chromosome painting requires delivery of hundreds of gRNAs with target sites throughout the chromosome. Using modified dCas9, Csm, or Cas13 family proteins that target RNA instead of DNA enables tracking of RNA in live cells, including orthogonal RNA/DNA labeling in the same cell.

Browse Plasmids: Visualize

Efficient Editing and Multiplexing with Cas12a

One of the most popular Cas9 alternatives is Cas12a (Cpf1). There are many Cas12a orthologs, but the most common is AsCas12a (from Acidaminococcus sp.). The popularity can be attributed to Cas12a’s many benefits.

Cas12a exhibits increased efficiency with larger edits using HDR. Cas12a-mediated DNA cleavage creates DSBs with a short 3′ overhang, allowing for directional gene transfer, which may increase the efficiency of gene editing. Cas12a also has more condensed multiplexing capabilities, as it can process multiple gRNAs under a single promoter, and can cleave ssDNA non-specifically.

Cas12a also has an expanded targeting range to AT-rich regions, as it recognizes TTTV (V = A, C, or G) PAM sites. Engineered versions of Cas12a recognize different PAM variants, with one of the most popular being EnCas12a, an engineered version of AsCas12a that recognizes TYCV and TATV PAM sites.

The DETECTR platform, developed by the Jennifer Doudna lab, can identify specific strains of pathogens or genetic mutations. In DETECTR, Cas12a is targeted to a specific genomic location, such as a viral genome, and a ssDNA-fluorescently quenched reporter is added. As this reporter is degraded by Cas12a, it will release a quantifiable signal that indicates the presence of your target DNA sequence.

Browse Plasmids: Cas12a

RNA Targeting and Editing using Cas13

Cas13 is a type VI CRISPR enzyme that recognizes ssRNA. Type VI enzymes, which include Cas13a, Cas13b, and Cas13d, can process crRNA precursors to mature crRNAs. Upon ssRNA recognition by the crRNA, the target RNA is degraded. In bacteria, Cas13 enzymes can also cleave RNAs non-specifically after the initial crRNA-guided cleavage, which slows bacterial cell growth and may further protect bacteria from viral pathogens. This promiscuity is unique to bacterial cells and does not occur in mammalian cells.

SHERLOCK, developed by the Feng Zhang lab, is a Cas13a-based molecular detection platform. Based on this non-specific cleavage, a Cas13a-based molecular detection platform, termed SHERLOCK, can identify target sequences using a fluorescent reporter. In SHERLOCK, a quenched ssRNA fluorescent reporter is added to the reaction with the Cas13a and gRNA. When Cas13a binds to the target sequence, it promiscuously cleaves the reporter as well, and a quantifiable signal is produced, indicating the presence of that specific target. SHERLOCK (and SHERLOCKv2) allows for greater amplification and quantification and can be used on lateral flow strips. Other Cas enzymes can be added to SHERLOCKv2 to detect multiple targets. For example, Cas12 can be added to detect the presence of infectious organisms (like SARS-CoV-2) and genetic mutations.

Similar to Cas9 and Cas12, Cas13 can be converted to an RNA-binding protein through mutations in its catalytic domain. REPAIR, developed by the Feng Zhang lab, used a hyperactive adenosine deaminase that acts on RNA, ADAR2(E488Q) fused to catalytically dead Cas13b to create a programmable RNA base editor that converts adenosine to inosine in RNA. Since inosine is functionally equivalent to guanosine, the result is an A to G change in RNA. The catalytically inactive Cas13b ortholog from Prevotella sp., dPspCas13b, does not require a specific sequence adjacent to the RNA target, making this a very flexible editing system. Additional editors have been developed, including editors based on a second ADAR variant that displays improved specificity, and editors carrying an ADAR truncation that retain RNA editing capabilities but are small enough to be packaged in AAV particles.

Cas13 fusions have also been used in in vivo RNA imaging or to control alternative splicing.

Browse Plasmids: RNA Targeting, RNA Editing