This feature on prime and base editing is adapted from content provided in the Cancer Genomics 2023 Report.
The introduction of CRISPR in 20121 was a groundbreaking discovery that irrevocably changed the way scientists approached human disease. Now, we have the ability to directly alter the genome itself, paving the way for tailored T cell therapies and gene editing methods. In addition to CRISPR, newer techniques such as base and prime genome editing are making waves in the gene editing world, providing novel ways to treat disease and overcome the limitations of CRISPR.
Defined as the alteration of a genome by inserting, deleting or substituting sections of DNA, genome editing is commonly used to fix mutations, modulate gene activity or add new elements into genes. In cancer treatments, genome editing can be the solution to a range of different issues, including correcting somatic mutations, silencing oncogenes, or even altering immune cells to better recognise cancer.
In this feature, we will explore the basic fundamentals of gene editing mechanisms, the development of base and prime editing, before finally delving into the exciting studies that demonstrate their impact on the field of precision oncology.
Gene editing: A simplified introduction
The basic mechanism behind genome editing is relatively simple; double strand breaks (DSBs) are introduced into the target genome at a specific location, which are then repaired by the cell to create the desired outcome. Typically, CRISPR uses an endonuclease known as SpCas9 to create DSBs at the target genomic loci, although other Cas9 variants may also be used depending on the CRISPR system. Then, the DSB is repaired by one of two main mechanisms – non-homologous end joining (NHEJ) or homology-directed repair (HDR) (see Figure 1).
Figure 1: Overview of homology directed repair and non-homologous end joining repair of double-strand breads generated by Cas9-mediated cutting. Sourced from van Kampen et al, 20192.
Researchers can manipulate cells into using the repair mechanism that best suits their needs. Repair mechanism selection is heavily influenced by cell type and DNA template availability. As stated previously, there are two main DBS repair mechanisms commonly used in gene editing techniques:
- Non-homologous end joining (NHEJ): This is the primary DNA repair pathway in mammals, where DSBs are repaired without a homologous template. Due to its error-prone nature, NHEJ is commonly used for gene activity reduction (commonly referred to as “knock-down” but can be referred to as “knock-out” if the gene activity is permanently prevented) through the production of loss-of-function mutations. However, in post-mitotic cells where HDR is downregulated (such as neurons), NHEJ can be used for the expression of exogeneous DNA inserts3.
- Homology directed repair (HDR): This is a more precise DNA repair pathway that requires a homologous template to guide repair. Although less likely to introduce errors, HDR is generally considered to be less efficient than NHEJ and only occurs at certain stages of the cell cycle. By introducing a homologous template that contains the desired DNA change, HDR-based CRISPR systems are an excellent choice for knock-in experiments in diving cells.
It is clear that CRISPR/Cas9 genome editing systems can be tailored to suit many biological needs. Although improvements to nuclease and delivery systems have made CRISPR more effective, low repair efficiencies and the generation of indels within the target genomic locus can be detrimental when producing gene editing therapies for clinical use. In recent years, several novel techniques that combat these challenges (such as base and prime editing) have risen to the forefront of the gene editing landscape, with several studies demonstrating their usefulness in the precision oncology field.
First established in 20164, base editing describes an elegant method to change single nucleotides without requiring donor DNA templates or DSB creation (see Figure 2). Instead, the base is chemically altered to generate point mutations at the desired genomic locus. Therefore, base editors can make small, specific changes to DNA in both dividing and post mitotic cells, without relying on HDR pathways or generating indels.
The first base editor, BE1, was designed to catalyse the reaction of cytosine to uracil (a C/G to T/A base pair conversion) using the activity of a cytidine deaminase enzyme fused to a dead SpCas9 variant (dCas9). This is an example of just one type of base editor, cytosine base editors (CBEs), although other base editors that change adenine into guanine (adenine base editors (ABEs)) have also been developed.
This highlights the limitation of the technique; only certain types of base modifications can be performed. Specifically, transversion mutations (i.e., the conversion of a purine base to pyrimidine base, or vice versa) are not possible. Additionally, this technique cannot be used to create indels, which can limit their functionality. However, the impressive efficiency and ability to create multiple edits within a single cell makes base editing an attractive tool for therapeutic gene editing.
Figure 2: Basic overview of the base editing system. Sourced from Zhao et al, 20235.
A new hope for CAR T cell therapy
In cancer therapy, DNA base editors can introduce single point mutations into immune cells to bioengineer better cancer immunotherapies. Often, this is done by creating CAR T cells, which are modified to express receptors that interact with specific cancer cell antigens. A major development in CAR T cell therapy came in December 2022, where base-edited T cells were used in a world-first clinical trial to treat acute lymphoblastic leukaemia (ALL)6, resulting in patient remission just 28 days post-infusion.
Similarly, in June 2023, a landmark study published in the New England Journal of Medicine detailed the use of base editing to treat aggressive T cell ALL in three children7. T cells from healthy donors were transduced to express a chimeric antigen receptor (CAR) with specificity for CD7 (CAR7), a protein that is expressed in ALL T-cells. Subsequently, base editing was used to introduce mutations into three genes encoding receptors – CD52, CD7 and the β chain of the αβ T-cell receptor – which inactivated the genes. This resulted in T cells with a reduced probability of CAR7 T-cell fratricide and graft-versus-host disease, making it a safer treatment for ALL patients.
The initial data from this study looks promising. Out of the three children that received the modified CARs, two entered remission within 28 days, whilst the third child died from unrelated complications. Building on this work, a new US-based trial (BEAM-201) was launched in September 2023, this time using T cells containing four base-edited genes8. The study utilises similar edits as the previous trial, with an additional edit in the PDCD1 gene to extend the lifespan of the engineered cells within the patient.
In contrast to base editing, the newly developed prime editing technique9 allows both base conversions (transition and transversion) and also the generation of small insertions and deletions. Occasionally called a “search and replace” technology, prime editing has the major benefit of not introducing DSBs whilst still delivering precise DNA modifications, although the efficiency of the process still requires optimisation5. Given its diverse functionality, prime editing could hypothetically correct up to 89% of known genetic variants associated with human disease10.
Prime editing works by creating single-stranded breaks (SSBs) into the genome, which are then repaired in an error-free manner (see Figure 3). This requires a fusion protein (a Cas9 nickase conjugated with a reverse transcriptase enzyme) that recognises the genomic target site and introduces a SSB into the non-target strand. Subsequently, the released 3’ DNA strand binds to the 3’ terminal region of the prime-editing guide RNA – containing the desired edit – and is reverse transcribed by the reverse transcriptase enzyme. After flap cleavage, the DNA is then ligated, and the edited DNA has been successfully incorporated into the genome.
Clearly, prime editing could have a significant influence on the precision oncology field. To this end, a number of prime editing-based methods have been developed that apply this technology to detecting and curing diseases11. Although the technology and early proof-of-concept studies seem promising, prime editing has yet to enter clinical trials5.
Figure 3: Basic overview of the prime editing system. Sourced from Zhao et al, 20235.
Making mouse models more accessible
As with many areas of medical research, animal models are a useful tool for determining the mechanisms that drive tumour initiation and disease progression. In cancer, genetically engineered mouse models (GEMMs) are a staple experimental tool for deciphering the effect of driver mutations in vivo, but despite their benefits, they remain expensive and time-consuming to produce. However, with the introduction of prime editing methods into the precision oncology world, this may all be about to change.
Published in May 2023, Ely et al. revealed the details of their in vivo prime edited mouse technique12, used to deliver a wide variety of cancer-related mutations into the mouse genome. By inserting a Cre-inducible prime editor into the mouse germline, rapid, tissue-specific expression of the desired mutation can be achieved. Notably, this approach used a lipid nanoparticle delivery system – thus avoiding the challenges of exogenous prime editor protein delivery, such as their large size and possible generation of anti-tumour responses within the mouse.
Using this technique, the authors also illustrated the feasibility of prime editing in mice by creating new models for lung and pancreatic cancer. Selecting clinically relevant KRAS mutations, the authors successfully introduced these variants into genetically engineered mouse strains and detected tumour formation. Overall, this study highlights the potential applications of prime editing in GEMMs, not just as a helpful tool for functional studies of cancer mutations, but also in clinical investigations of targeted therapeutics.
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