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Technology Networks Explores the CRISPR Revolution: An Interview With Dr Amy Butler

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In this week's instalment of Technology Networks Explores the CRISPR Revolution, we're focusing on the technical elements of CRISPR technology.

How have the experimental approaches to CRISPR research studies changed over recent years? Some researchers opt to use a CRISPR kit for their research studies, whereas others may decide to use a "DIY" approach. What are the benefits and drawbacks of each, and why might a researcher choose to adopt one over the other?

Helping us to answer these questions is Amy Butler, Ph.D., President of the Biosciences Division at Thermo Fisher Scientific.

MC: Please can you briefly outline how the CRISPR-Cas mechanism can be programmed for gene editing? Why might a researcher choose to utilize CRISPR tools for gene-editing rather than alternative available methods?

Amy Butler (AB):
At a high level, genome editing is the process of introducing a specific DNA sequence change at a specific site within the genome of a cell. To accomplish this, scientists harness the cell’s own DNA repair processes to make these changes happen. All of the current genome editing technologies in their native forms, whether it be CRISPR, TALENS, or Zinc finger nucleases, are used to initiate this process by introducing a double stranded break (DSB) in the genomic DNA at or near the site of the desired edit.

Depending on the type of edit desired, researchers rely on the cell to use one of two broad categories of DNA repair processes to accomplish the desired change. If a researcher is creating a targeted knockout of a gene, then they are largely going to rely on the cell to fix the DSB using a process called Non-Homologous End Joining (NHEJ). NEHJ is error prone resulting in the insertion or deletion of base pairs at the repair site. This results in a functional knockout of the target gene.

The second broad category of edits are what are typically referred to as knock-in edits. These are the more elegant edits that create the precise changes to the genome that we think about when we think about the promise of genome editing technologies. This can involve single base pair changes up to the insertion of large pieces of exogenous DNA. Here again, scientists rely on the core nucleases (CRISPR, TALENS, and ZFN’s) to introduce the DSB at the site of the desired edit. For Knock-in edits, not only do we deliver the genome editing nuclease, but we also provide the cell with what we call a donor DNA molecule to assist with the repair. Within the donor DNA molecule, we encode the DNA sequence changes that we want to introduce. When a donor is available it drives the cell towards the use of a repair process called Homology Directed Repair (HDR). During HDR the cell utilizes the donor DNA molecule to repair the DSB resulting the incorporation of the desired sequence change at the target site.

Any of the targeted nucleases can be used in this process but CRISPR has been adopted as the go-to genome editing nuclease in the market at present because it has been the quickest and easiest of the genome editing technologies to apply. Targeting a new region of the genome simply requires the user to create a new guide RNA (gRNA). However, Thermo Fisher Scientific and others continue to develop the other technologies alongside CRISPR. We realize that, although CRISPR is a versatile technology, there are some limitations which can be addressed using the other technologies (ie. TALENS and Zinc-Fingers). Developments across the field are contributing to a complete toolbox of editing tools that will allow us to explore a wide range of research questions in nearly any cell type.

MC: How have the experimental approaches to CRISPR research studies changed and advanced over recent years?

The technology was initially developed around the use of DNA plasmids to deliver the nuclease and the gRNAs. While this approach remains popular, there has been a significant shift to the direct delivery of Cas9 protein and synthetic gRNAs into the cell. This approach has removed multiple inefficient steps in the assembly of the tools and delivery to the cell, resulting in a significant increase in editing efficiency (often reaching 70% or higher) while at the same time significantly reducing off-target activity. The result is a reduction in the cost and effort required to identify and expand edited clones, and those isolated clones become a more reliable research tool. We developed Thermo Fisher’s TrueCut Cas9 protein and TrueGuide Synthetic gRNA for this space and they continue to be excellent workhorse editing tools for our customers.

The second area that CRISPR has made significant advances is in the field of functional genomics screening. High throughput screening has been a core approach to identifying the genes involved in specific biological processes and diseases. Traditional approaches have relied on RNAi technologies which have known shortcomings, including incomplete knockdown of the target gene(s) and relatively high off-target activity. CRISPR, however, has the benefit of being able to produce a complete and permanent knock-out of the target gene and reduced off-target effects compared to RNAi. This in turn has the potential to result in higher quality hits emerging from screens and to accelerate research programs by helping to identify critical gene targets faster and more reliably. Thermo Fisher has developed our LentiArray and LentiPool CRISPR libraries to support this type of screening.

MC: There is now a huge diversity in the range of CRISPR-Cas tools available. Briefly, how do each of the Cas systems differ?

Various CRISPR-Cas systems have been described in hundreds of bacteria species. Some of these variants such as Cas12/CPF-1 and strep. aureus Cas9 have different properties and targeting requirements when compared with Cas9 from strep. pyogenes (considered WT cas9 and the workhorse CRISPR-Cas9 system used in most labs). At present the requirements for these alternative nucleases generally result in a more restricted design space within the genome and in general their editing efficiencies tend to be much lower than Cas9 from strep. pyogenes. It will be interesting to see how this space develops over the next several years. The real excitement in the expanding space of CRISPR is in new variants of strep. pyogenes Cas9 which enable reduced off-target effects, high fidelity Cas9 variants which allow for manipulation of gene expression without making permanent edits to the genome, and CRISPRi and CRISPRa base editors which allow direct change of single base pairs without the need to cut and introduce a donor DNA molecule, and variants that can alter epigenetic regulation.

MC: Some researchers may opt to use a CRISPR kit for their experiment, whilst others may choose a "DIY" approach. What are the benefits and drawbacks of each? Why might a researcher choose to adopt one over the other?

We consider the plasmid-based approaches as the “DIY” approach. In general, this is the approach that many scientists are first exposed to because it was what was first reported in the literature and the plasmids are readily available. The issues with this approach are that it tends to have much lower efficiency and much higher rates of off-target editing when compared to using “kitted” Cas9 protein and synthetic gRNAs solutions, such as TrueCut Cas9 protein and TrueGuide Synthetic gRNAs.

In fact, the use of Cas9 protein, as opposed to the plasmid, results in such significant increases in editing efficiency that researchers can now often conduct their experiment in an edited pool of cells without isolating clones. This opens up a whole host of possibilities for using the tool in primary and non-dividing cells. Importantly, this jump in efficiency and reduction in off-target effects from using the Cas9 protein and synthetic gRNA approach is one of the key developments enabling CRISPR to rapidly move from the research bench to the clinic in cell therapy applications.

MC: How can we continue to develop CRISPR tools that are applicable in a wide variety of cell types?

When discussing application to a wide variety of cells, the limitations of the current tools revolve around efficient delivery of the editing tools to the cells. This is why Thermo Fisher has focused on the development of the delivery systems alongside the delivery of the editing tools themselves. We have shown that coupling our editing reagents with the Neon electroporation system enables us to apply the tools to a wide variety of cells from workhorse cell lines to iPSC cell and primary cells. In most cases we can find a protocol to drive high-efficiency editing, even in challenging cells such as human primary T-Cells where we can routinely drive 90% or better editing efficiency.

MC: What limitations do researchers still face in CRISPR-Cas experiments? How can we look to overcome these limitations?

We are overcoming limitations and making the systems easier to use by taking a holistic approach to the entire process. We continue to focus on developing tools across the entire workflow from Gibco media systems, delivery platforms such as the Neon and the Lipofectamine CRISPRMAX reagent, the editing tools including TrueCut Cas9 and TrueGuide Synthetic gRNAs and downstream analysis tools to validate that the desired edit has been achieved and that we haven’t introduced any undesired off-target edits. By taking a holistic approach, we have been able to tackle limitations in different cell types such as iPSC or primary T-Cells and drive high level editing efficiency, which in turn is having a significant impact on the ability of researchers to rapidly move their research programs forward.

Amy Butler, President of the Biosciences Division at Thermo Fisher Scientific, was speaking with Molly Campbell, Science Writer, Technology Networks.

Catch up on the previous instalment of Technolology Networks Explores the CRISPR Revolution, an interview with Professor Glen Cohenn, here.