Beyond the Double-Stranded Cut: Evolution of CRISPR Methods

Base Editing is an advanced genome editing technique that precisely converts one base pair to another without introducing double-strand breaks in DNA. The process utilizes a catalytically inactive (dCas9) or nickase Cas protein fused to a DNA deaminase enzyme. These enzymes enable specific base conversions, such as C•G to T•A or A•T to G•C, via direct chemical modification of the DNA sequence. There are currently two types of base editing systems: cytidine deaminase base editors (CBEs) and adenine deaminase base editors (ABEs). Cytidine deaminases are naturally occurring enzymes isolated from bacteria that induce C to T substitutions in DNA. Adenine deaminases, however, do not occur naturally and were engineered from existing bacterial enzymes to induce A to G substitutions.

The majority of human genetic diseases are caused by single nucleotide polymorphisms, making base editors an ideal therapeutic option. Base editing is highly specific and minimizes undesired edits, off-target effects, or larger genomic rearrangements typically associated with double-strand breaks. However, challenges remain in the delivery of Base editing components to target cells, particularly in vivo, as well as in minimizing rare but potential off-target activity. Addressing these challenges is crucial for ensuring safe and effective use in therapeutic settings. Base Editing shows immense promise in treating genetic disorders caused by point mutations, including inherited blood disorders and metabolic diseases. It is being actively explored in preclinical studies and clinical trials, with the goal of translating its precision into impactful therapies for conditions that currently lack effective treatments.

For a deeper understanding of the mechanism, challenges, and its implications in cell and gene therapies, visit our Base Editing application page for full details.

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Prime editing systems allow for both substitution mutations and the insertion and deletion of small fragments of DNA without DSBs. Prime editors (PEs) use an engineered Cas9 nickase (nCas9), which cuts only one strand of DNA, fused to reverse-transcriptase (RT), a viral enzyme that allows for reverse transcription of a DNA template, effectively writing a new section of DNA into the genome. Unlike CRISPR-Cas9, which uses an sgRNA to direct editing and a DNA donor template for HDR, PE systems employ a single construct known as a prime editing guide RNA (pegRNA).

The engineered pegRNA construct contains both the primer binding site (PBS) to direct editing and a DNA sequence that contains the edit; the Cas9 nickase binds at the PBS and cuts one strand of DNA, and the RT uses the other part of the pegRNA sequence as a template for reverse transcription, adding the desired nucleotides to the nicked strand. The original DNA sequence is then removed by endonucleases, and an additional guide RNA directs nCas9 to nick the opposing (unedited) strand, causing the cell to repair the DNA using the edited strand as a template.

The original PE system allowed for insertions of up to 50 base pairs and deletions of up to 80 base pairs. A newer iteration, twin-prime editing (twinPE), uses two pegRNAs to simultaneously target opposing strands of DNA, allowing for the insertion and deletion of much larger sequences. PE systems show promise in the treatment of many genetic disorders, including heart disease, sickle cell disease, prion diseases, diabetes type II, and Alzheimer’s, while twinPE can potentially be used to treat diseases caused by multiple pathogenic mutations in large genes.

Dive deeper into the mechanisms, challenges, and clinical uses of prime editing on our Prime Editing application page.

The first CRISPR derivatives used to make genetic modifications without inducing DSBs were the CRISPRactivation (CRISPRa) and CRISPRinhibition (CRISPRi) systems. These technologies use a catalytically dead Cas9 (dCas9), which retains its search function, but has been inactivated so that it can no longer cleave DNA.

CRISPRi originally worked via directing dCas9 to the promoter of a target gene, where it will bind and prevent transcription of the gene, leading to gene silencing. This activity was later enhanced by fusing dCas9 with other effectors that help to repress transcription. In contrast, CRISPRa uses dCas9 fused to transcriptional activators, directing them to gene promoters or enhancers to increase transcription.

However, CRISPRi and CRISPRa can only regulate the expression of genes that are already present in the genome and are not capable of correcting disease-causing mutations. Using CRISPR to actually modify the underlying genetic sequence without creating DSBs was more challenging, and required the use of other types of proteins from bacteria and viruses to work.

One of the interesting discoveries when scientists started exploring other bacterial CRISPR systems was new Cas nucleases that could target RNA instead of DNA. The ability to edit RNA offers an interesting alternative to traditional CRISPR because RNA is transiently expressed; any changes to RNA inside a cell are not permanent or heritable. There are many RNA-mediated diseases in humans caused by defects in the splicing, editing, modification, or translation of RNA transcripts which can potentially be treated using RNA editing. Viruses with RNA genomes, including SARS-CoV-2, can also be targeted using RNA editing.

The first RNA-targeting Cas nucleases to be described were the Cas13 family, which includes many subtypes. However, Cas13 cannot be used for therapeutic editing because of its problematic ‘collateral activity’; once it recognizes and cuts its target transcript, it tends to destroy all other cellular transcripts, causing cell death. Cas13 is now primarily used for diagnostic purposes, including for the rapid detection of the SARS-CoV-2 virus and other RNA pathogens.

The more recent discovery of Cas7-11 has led to renewed interest in RNA editing for therapeutic purposes. Cas7-11 displays highly specific RNA editing with no collateral activity. Importantly, it is also more efficient than other RNA knockdown methods, such as short hairpin RNA (shRNA), and with less impact on cell viability. This new nuclease provides an avenue for the treatment of RNA-mediated diseases for which there are limited treatment options, including Huntington’s disease. It can also potentially be used for the therapeutic removal of RNA viruses from human cells.

Programmable Addition via Site-specific Targeting Elements (PASTE) is one of the latest additions to the CRISPR toolbox, harnessing the power of prime editors and building on them. PASTE uses bacteriophage serine integrase proteins to insert up to 36 kb of DNA into the genome. Cleverly, the PASTE system employs prime editors to integrate DNA sequences known as integrase landing sites (AttB), which the integrase protein attachment site (AttP) will locate and bind to for editing. Using PEs to integrate the AttB sites, PASTE can potentially be used for targeted large-scale gene knock-in anywhere in the human genome.

The PASTE method could have enormous therapeutic value in the treatment of genetic diseases caused by many pathogenic mutations in large genes. For example, X-linked agammaglobulinemia (XLA) is caused by a variety of mutations in the Bruton's tyrosine kinase (BTK) gene, and researchers have struggled to develop a CRISPR-Cas9 gene therapy for this disease based on the size of the required DNA donor template for repair. PASTE may offer an alternative to correct the mutated gene with relative ease, without inducing potentially harmful DSBs.

PASTE editing allows for programmable gene insertion without double-stranded breaks, independent of DNA repair pathways. Image from paper.

CRISPR-based epigenetic editing is another nascent method for therapeutics without DSBs. The epigenome controls how our genes are expressed through DNA methylation (5mC), histone modifications, and non-coding RNAs. Because 5mC and histone modifications are regulated by enzymes, such as DNA methyltransferases and histone acetyltransferases, dCas9 can be fused to epigenetic enzymes or their catalytic domains in order to alter the epigenome at a target site, switching gene expression on or off.

For example, if a gene is transcriptionally repressed by 5mC in the CpG islands of the gene promoter, this epigenetic mark can be oxidized by fusing dCas9 to the catalytic domain of a ten-eleven translocation dioxygenase, such as TET1CD, restoring expression. In contrast, if a gene is upregulated due to the addition of histone 3 lysine 27 acetylation (H3K27ac), dCas9 fused to histone deacetylase 3 (HDAC3) can be used for targeted deacetylation, reducing the expression of the gene.

This type of CRISPR-based epigenetic reprogramming offers the ability to switch genes on and off, with a range of potential therapeutic applications. Although there are no current trials, there have been many proof-of-concept and pre-clinical studies demonstrating the power of these methods. For example, epigenetic editing can upregulate fetal hemoglobin production in adults who suffer from sickle cell disease. Similarly, it can restore the expression of genes repressed by hypermethylation, as seen in the FMR1 gene which causes Fragile X syndrome.

Repressive epigenetic marks can also be removed in order to alleviate neuropsychiatric disorders, such as anxiety and alcohol use disorder, caused by excessive alcohol exposure during adolescence. Additionally, epigenetic editing has enormous potential in cancer therapeutics by correcting forms of epigenetic dysregulation that contribute to tumor growth.