Avoiding double strand breaks on the CRISPR path to therapeutic discovery

Now a key pillar in the early stages of therapeutics development, the limitations of introducing a DNA double strand break mean that workflows are broadening away from CRISPR–Cas as a gene knockout technology and towards platforms that enable genetic modulation and safer genetic alterations. Nicola McCarthy, Horizon Discovery, explains.

Navigating from target identification to validation to investigational new drug (IND) status is a complex process, not least in finding the right assays and models to validate or refute a hypothesis ahead of spending vast amounts of money on clinical trials. The past 10 years has seen acknowledgement that many therapeutics entering clinical trials fail to meet expectations set by pre-clinical workflows and that we all need to up our game in terms of accurate assessment of biological impact in vitro and in vivo. One Nobel Prize winning innovation that has made a substantial impression in this space is CRISPR–Cas gene editing. With the many different flavours of the CRISPR–Cas system now available, choosing the right format to address current complex biological questions in the therapeutics space is key.

Target ID and validation

The CRISPR–Cas gene editing platform is routinely used to knock out genes across the genome to assess how loss of each specific gene impacts a particular biological response. This could be in the form of a pooled CRISPR–Cas9 forward genetic screen, where a library of single guide RNAs (sgRNAs) targeting the initial coding exons of all protein coding genes combined with the nuclease Cas9 leads to gene knockout1,2. Or one can use CRISPR–Cas9 to generate knockout cells to investigate the impact of gene loss using a variety of in vitro and in vivo assays3,4.  Trust in these types of screens and validation models has increased owing to concordance between CRISPR screening studies5-8 and studies showing that approved and pre-clinical drugs have different mechanisms of action to those ascribed to them, as they retain their biological effects when the genes encoding their purported protein targets are knocked out by CRISPR–Cas9.

Discovering and validating new targets for human therapy has always been challenging, so it is not surprising that new technologies, such as CRISPR–Cas, highlight past deficiencies. However, these findings are not an indication that CRISPR–Cas gene knockout approaches are the way forward for all biological questions that are relevant in drug discovery. Iterations of CRISPR–Cas in the lab are numerous and have, for example, enabled gene modulation (CRISPR inhibition (CRISPRi)10-15; CRISPR activation (CRISPRa)15-20); and gene and chromatin labelling and imaging21, which provide alternative routes for forward genetic screens and target validation. When altering DNA sequences, greater precision is now available in the form of base-pair modification (base editing)22- 24 and re-writing of short sequences of DNA (Prime editing)25. All these recent innovations have one thing in common: they no longer require the nuclease activity of the Cas enzyme. This affords several advantages for different areas of biological study where avoidance of a DNA double strand break (DSB) could be crucial.

Avoiding the nuclease

CRISPR–Cas9 systems are best known for gene editing applications, but nuclease dead versions of Cas9 (dCas9) are used to build systems that enable gene silencing or gene activation10-21. In CRISPRi and CRISPRa, dCas9 and an sgRNA allows disruption of gene transcription by binding to proximal sequences at the promoter region of the gene and recruiting either a transcriptional repressor or a transcriptional activator. For example, by covalently linking the KRAB repressor to dCas9, suppression of gene transcription is achieved10,15,26. Conversely, it is also possible to create a transcriptional activation tool by fusing dCas9 to activation domains such as VP64 and p6515,17,27.

Improvements to the efficiency of CRISPRi and CRISPRa have been achieved through studying the interaction of dCas9 and its guide RNA with DNA and how the presence of nucleosomes can inhibit sgRNA access28-31. Such improvements are important as they enable the substitution of CRISPRi for CRISPR knockout in studies where CRISPR knockout isn’t ideally suited. Examples include the study of hypomorphic phenotypes or essential genes, and in the latter stages of drug development one can argue that complete gene KO might not be the best model for druggability, as most drugs do not reduce the activity to their target 100%. In several types of cancer, important genes that influence cancer biology exist as multiple copies and these cannot be knocked out by CRISPR–Cas because the multiple DNA double strand breaks introduced lead to p53 activation and commonly cell death32,33.  CRISPRi is a useful tool in these situations where gene transcription can be silenced and the impact studied without the risk of collateral DNA damage confounding the results. CRISPRa provides the capacity to study phenotypes arising from increased gene expression, which are common in the aetiology of cancer, for example, and have been successfully employed to look for host factors that protect against viral infection34,35. CRISPRi and CRISPRa might also be more suitable for studying differential expression of long noncoding RNAs, as these genes have proven difficult to target effectively with the CRISPR knockout platform36.

Another advantage of CRISPRi and CRISPRa is that they do not result in the alteration of the DNA sequence so their impact on gene expression is reversible: this is not possible with CRISPR knockout systems as the DNA sequence is altered through the introduction of a DNA double strand break and its subsequent repair.

Screens in which alteration of a DNA sequence is required, but for which introduction of DSBs could be detrimental now have an alternative in the form of base editing. This novel technology is based on the use of an engineered Cas enzyme that nicks a single strand of the DNA and a deaminase enzyme that chemically alters a specific nucleotide in the DNA sequence of a gene at a precise location, specified by a guide RNA22-24. The conversion of bases, for example a C:G changed to a T:A, can be used to knockout gene transcription by introducing a stop codon (for example changing a CAA to TAA) early in the coding part of the gene37or to disrupt conserved splice sites thereby impairing RNA maturation38.

Three recently published screens39-41 using base editors examined the impact of clinical associated genetic variant single nucleotide polymorphisms (SNPs) on genes involved in the DNA damage response and the impact of SNPs on sensitivity and resistance to cancer therapeutics. SNPs are numerous within the genome and their association with disease aetiology and drug response are well established but, studying their impact on a genomic scale through precision editing was not possible before the development of base editors.

CRISPRi, CRISPRa and base editing platforms are providing new methods for screening for genetic targets that impact human disease and treatment response, but the targets identified in these types of screens still need to be validated.

Doubling up for target validation

Validation remains a painstaking process and faster routes to validated genetic targets for therapeutic development are still needed. One way to achieve this could be through combining CRISPR screening platforms. Screening with both CRISPR knockout and CRISPRi platforms, for example, can provide an initial validation of the hits identified using either approach42. Here, the use of CRISPRi might identify genes that in the CRISPR knockout screen appear to be essential genes (genes encoding proteins that when completely lost cause the death of the cell) but are in fact valid therapeutic targets for drug development. A similar approach could be achieved by combining CRISPRi screens with base editing screens to limit the introduction of DSBs into the screening and validation process. This could be particularly beneficial when looking to screen or validate targets in primary cells, such as immune cells, where DSBs are more detrimental to cell survival. Screening with more than one CRISPR modality can help to identify gene networks that regulate a biological process of interest. For example, running both pooled CRISPRi and CRISPRa whole genome screens identified components of the apoptotic pathway that are crucial for the cellular response to the BRAF inhibitor vemurafenib43.

Although combining pooled screens generates a rich set of genetic data based on endpoints such as survival, proliferation or expression of cell surface proteins, arrayed CRISPR screens allow for more complex endpoints to be explored on a gene-by-well basis.

Arrayed screens can use either lentiviral or synthetic guide RNAs. Lentiviral arrayed screens have successfully identified genes underlying disease aetiology44-46, but such libraries are challenging to construct and costly to manufacture when compared with synthetic guide RNAs.  Synthetic guide RNAs can be introduced into cells using reverse transfection or electroportation in 384 well plates, making these screens amenable to high throughput automation47,48 and phenotypic endpoints49. Furthermore, synthetic guide RNAs are well-suited for multiplexing, enabling multiple guides to be used per well to achieve high rates of gene knockout47 or in arrayed CRISPRa screens for multiple genes to be activated, enabling pathway analysis or gene network discovery. For example, an arrayed CRISPRa crRNA library was used to identify positive and negative regulators of interleukin 6 secretion48. Interleukin 6 is a key inflammatory mediator in rheumatoid arthritis and is a crucial component of a cytokine storm, which results from overactivation of the immune response. This has been seen with the wider use of immunotherapeutics, such as checkpoint inhibitors, for the treatment of patients with cancer and more recently in patients hospitalised with severe Covid-19.

Gene-by-well readouts, such as cytokine production, lend themselves well to arrayed screens but, the use of co-culture endpoints and imaging endpoints are also possible in an arrayed format49,50. For example, CRISPR–Cas synthetic guide RNA screens can be used in primary T cells or regulatory B cells and co-culture endpoints, such as a mixed lymphocyte reaction (MLR) assay, T cell lysis or T cell suppression, can be used to assess the impact of gene loss. The use of co-culture endpoints along with primary cells instead of cell lines brings investigators a step closer to the cells that exist in their patient population and should improve the accuracy of target identification and validation. Such screens are needed now more than ever, given the recent shift in focus to primary cells rather than cell lines for drug discovery.

From cell lines to cell therapy

Cell lines have been the workhorse for initial target identification and validation in the drug development process however, the increasing success of immune-system-based therapeutics in the past 20 years, particularly in oncology, has triggered a widespread move in the pharmaceutical industry to cell-based therapeutics.

In 2017, the FDA approved two CD19-targeted chimeric antigen receptor (CAR) T cell therapies for the treatment of B cell lymphoma (Yescarta) and B cell leukaemia (Kymriah) and more recently approved Tecartus and Breyanzi for the treatment of adult patients with relapsed or refractory mantle cell lymphoma and diffuse large B-cell lymphomas, respectively. These cell-based therapies are produced using the patient’s own cells, also referred to as autologous cell therapy, using a process that is challenging and logistically complex51. For example, each autologous therapy originates from the patient’s cells therefore, heterogeneity is large from batch to batch and crucial quality attributes are difficult to maintain, reducing the safety and efficacy of the treatment. Autologous manufacture of CAR T cells also requires two clinical procedures per dose (isolation and re-infusion) that must be well timed and have substantial risk mitigations: cells delivered to the manufacturing centre and then back to the clinic are at increased risk of damage, loss and cross contamination.

By contrast, allogeneic cell therapy (using cells from a donor who is not the patient) offers several advantages and aligns with procedures and operations of pharmaceutical companies52,53. Cells that originated from a healthy donor are genetically engineered, analysed and batch controlled and can be stored in a cell bank system and shipped to patients when needed. As such, the patient receives the cell treatment on demand straight from the manufacturing facility, which saves precious time, particularly in the context of rapid disease progression.

Allogeneic immunotherapies might have many logistical advantages, but there are still challenges to overcome: the possibility of graft versus host disease (GvHD), where the donor’s T cells attack the recipient’s cells, and rapid elimination of the donor cells before they can elicit their therapeutic action owing to the patient’s immune system seeing these cells as ‘foreign’ to the body. For CAR T cells, GvHD is relatively easy to avoid through deletion of the gene or genes encoding the T cell receptor. However, human leukocyte antigen mismatches between the donor and the patient (alloreactivity), which trigger elimination of the donor’s cells, is harder to address as multiple genes are involved54.

CRISPR–Cas9 gene editing can provide the multiplex editing needed to knock out several genes involved in leukocyte antigen expression. However, the risks of this approach are high as multiple DSBs in the genome increase the risk of substantial alterations in the DNA, such as chromosomal translocations, risking adverse events for the patients arising owing to rare, mutated cells within the edited population of CAR T cells. In this context, the use of base editors to knockout multiple genes while avoiding the introduction of DSBs is a preferential route for editing allogeneic cell therapies.

It is too early to predict which approach — allogeneic or autologous cell therapy — will dominate the market in the future. However, when analysing trends in cell therapy development, a greater percentage of pre-clinical publications and Phase I trials are in the allogeneic therapy space55. Similar trends appear when examining large deals and partnerships being won by allogeneic cell therapy companies as opposed to companies sat squarely in the autologous space. Ultimately, the safety and efficacy profile of the treatment for a particular indication is the most important consideration. An allogeneic cell therapy that will offer similar safety and efficiency to autologous cell therapy would make the latter obsolete for that indication based on current logistical practices.

From bench-to-bedside and back again, CRISPR in its current guises is having substantial influence on the development of the next generation of therapeutic products. Although now a key pillar in the early stages of therapeutics development, the limitations of introducing a DNA double strand break mean that workflows are broadening away from CRISPR–Cas as simply a gene knockout technology and moving towards platforms that enable genetic modulation and safer genetic alterations that enable the engineering of multiple genes in one application. This is an exciting time to be working in target ID and validation.

Volume 22, Issue 2 – Spring 2021

About the author

Nicola McCarthy is the Global Translational Science Liaison at Horizon Discovery, a PerkinElmer Company, where she has worked for the past six years. Her role involves working with scientific and commercial teams at Horizon with a view to understanding better the scientific needs of our customers. McCarthy studied human anatomy for her B.Sc. and programmed cell death for her Ph.D. at the University of Birmingham, UK.

Acknowledgements

Thank you to Jonathan Frampton, Jennifer Harbottle, Michael Anbar, Žaklina Strezoska, Steven Jarvis and James Goldmeyer for helpful comments and information during the drafting of this article.

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