Q&A Report: How to Create CRISPR-Edited T Cells More Efficiently for Tomorrow’s Cell Therapies

Explore future gene-editing trends, CRISPR challenges, and solutions with experts from Artisan Bio and STEMCELL Technologies.

The answers to these questions have been provided by:

Steven Loo-Yong-Kee, MEng, Scientist II at Artisan Bio

STEMCELL Technologies team

What do you believe will be the most significant developments in the gene editing field in the next few years?

Steven Loo-Yong-Kee: The field of gene editing is expected to see significant progress in the coming years. One notable trend that will continue is the shift from lentivirus-based editing, used in the first CAR-T therapies, to more precise CRISPR-based techniques. This transition is driven by the growing demand for a deeper understanding of the final product and its critical quality attributes (CQAs).

Another key area of progress is in allogeneic therapies, where induced pluripotent stem cells (iPSCs) and progenitor cells are gaining popularity due to their potential for off-the-shelf products. By harnessing the differentiation potential of iPSCs and progenitor cells, scientists can produce a diverse range of cell types for therapeutic applications.

Furthermore, high-throughput technologies and genome-scale knock-in methods will enable efficient modification of multiple genes or genomic regions simultaneously. Combined with artificial intelligence (AI) and machine learning (ML), data interpretation and visualization will play a pivotal role in helping researchers gain insights into complex biological systems.

As the gene editing field progresses, a significant focus will be on the development life cycle and the utilization of Good Manufacturing Practice (GMP)-ready materials. Companies like Stemcell Technologies provide GMP-grade materials and reagents necessary for safe and scalable cell therapies. The early adoption of GMP-ready materials will de-risk the processes and ensure quality, safety, and regulatory compliance of cell therapies as they move toward clinical translation.

STEMCELL Technologies: We have seen trends towards the use of iPSCs as starting material for the next generation of cell therapies, versus primary cells, because of their availability, quantity, possibility of using universal lines, amenability to editing, and ability to differentiate into several tissue types. This means that editing methods will also need to work well in this cell type.

Another trend we see is the increase in multiplex editing. Many researchers are capitalizing on the self-renewing properties of iPSCs by incorporating numerous edits into the genome, with the goal of creating bespoke combinations of gene modifications that endow cells with advanced capabilities. These may include functions such as host immune evasion in allogeneic cell therapies or enhancing migration and tumor specificity in CAR-T therapies. However, the introduction of multiple edits carries inherent risks to genome stability due to the increased manipulation and selective pressure placed on these modified cells. Creating robust workflows and new editing technologies that mitigate these risks and maintain high-quality iPSCs throughout the process of multiple editing will be a key focus in the field.

What is the biggest challenge you've encountered when editing iPSCs using CRISPR?

Steven Loo-Yong-Kee: The biggest challenge encountered when editing iPSCs using CRISPR is the poor viability and recovery of cells after editing. This can be attributed not only to the inherent nature of CRISPR technology, including the induction of DNA breaks and potential off-target effects, but also to factors like the transfection delivery method, such as electroporation. Scientists are working on optimizing both the editing conditions and the transfection delivery methods to improve viability and recovery rates.

STEMCELL Technologies: Inserting extra-large cargos into the genome can be very tricky, in particular as size increases >12kb. Editing efficiency and cell viability both decrease, and the remaining clones require extensive characterization to ensure the full cargo has been inserted properly, and that there are no off-target integrations that enhance survivability by unintended or unwanted means. These undesirable consequences can be mitigated by using high-fidelity gene editing tools, together with well-optimized workflows, and high quality starting iPSCs.

How are double insertions determined when designing a CRISPR experiment? Are double insertions in two different genes used to reduce side effects?

Steven Loo-Yong-Kee: When designing a CRISPR experiment, double insertions can be achieved using various approaches. One method is to employ bicistronic dual inserts into the same loci, which allows for the introduction of two mutations simultaneously. While two insertions in two different genes may seem like a strategy to reduce side effects, they can actually complicate the process. Simultaneous edits in multiple genes increase the risk of translocations, which can have unintended consequences. Therefore, careful design and downstream analytics are necessary when attempting double insertions in different genes.

What approach do you take to ensure a probable absence of transformation-associated functional phenotypes (TafP) while keeping the process within rational bounds, considering factors such as cost and time, and taking into account critical cell quality attributes beyond knock-out (KO) and knock-in (KI) transfection efficiency? Additionally, how do you design guides to minimize off-target effects in gene editing?

Steven Loo-Yong-Kee: Eliminating transformation-associated functional phenotypes (TafP) in CRISPR-engineered cells compared to wild-type cells poses a significant challenge. To address this, we employ a risk-based approach to nominate guide RNA designs that minimize unintended transformations. We achieve this by leveraging both publicly available and proprietary in silico predictions, as well as in vitro techniques like Digenome-seq. These measures effectively reduce the risk of TafP and enhance the precision of our CRISPR engineering process. Once the cut sites are selected, we proceed with in-cell cut site confirmation using rhAmpSeq. Additionally, we conduct thorough analyses to detect chromosome aberrations and perform phenotypic characterization as needed. This approach carefully balances TafP prevention with cost and time considerations, ensuring the integrity of the engineered cells.

What is the impact of starting with fresh versus frozen samples on editing efficiency?

Steven Loo-Yong-Kee: Starting with fresh or frozen samples has not shown any significant differences in editing efficiency, based on our observations. Advancements in cryopreservation have greatly improved the viability and functionality of frozen samples. However, it is important to optimize the thawing and recovery processes for frozen samples to minimize potential negative effects on cell viability and editing efficiency.

STEMCELL Technologies: For hPSC cultures, editing efficiency immediately after thaw is slightly reduced compared to editing fresh cultures. However, starting from a frozen source and editing after one or two passages can promote standardization across experiments during the process development stage.

It is important to distinguish between transfection efficiency (ability to get cargo into the cells), editing efficiency (how many cells were edited), and plating efficiency (how many cells survived the process and are available to assay). All three efficiencies are important as a deficiency in one will impact the overall success of your experiment. It is unlikely that there would be a significant difference between fresh and frozen samples in terms of transfection or editing efficiency, the major difference would likely be observed in plating efficiency. In this case, it does not matter how many cells received the desired edit if the majority of cells die post-plating. This is why it is critical to have high-quality, healthy cells going into the transfection and improving plating efficiency using specialized reagents, including CloneR™2.

What is the maximum payload size for knock-in (KI) experiments, and how is knock-in performed?

Steven Loo-Yong-Kee: We utilize HDR templates to enable knock-in for precise genetic engineering. By leveraging the homology-directed repair (HDR) pathway, we utilize an HDR template as a donor DNA to introduce specific genetic sequences into the target genome. This approach allows us to achieve accurate and efficient genetic modifications. Our knock-in experiments have successfully incorporated payloads up to 10kb in size, including a bicistronic insert comprising a CAR and a second insert.

Can I purchase the STAR nuclease and guides for gene editing separately, and do I need a CRISPR license to use your process?

Steven Loo-Yong-Lee: Artisan offers partners a collaboration plus licensing business model. We can perform genome engineering R&D, including novel guide development, off-target validation data packages, and payload insertion optimization. Partners have the option to obtain the clinical and commercial STAR-CRISPR license, which allows for CDMO and GMP manufacturing. Artisan licenses provide foundational CRISPR IP, eliminating the need for external third-party licenses to utilize our STAR-CRISPR technology. For further details on our collaboration services and licensing terms, please contact bd@artisancells.com.

Did the transduction efficiency correlate with the functional data you obtained?

Steven Loo-Yong-Kee: We found a good correlation between on-target editing efficiency and our functional data. Our editing efficiency, determined by INDEL % from Amplicon Seq, showed a strong correlation with the knock-in and knock-out effects of our edits, assessed using flow cytometry.

Do you know if it’s possible to do gene editing in adult stem cells?

STEMCELL Technologies: Yes, it is possible to edit adult stem cells. There are many groups currently working on editing CD34+ HSPCs derived from donors for the purpose of developing cell therapies. Some groups are also conducting CRISPR-based knock-out screens in intestinal organoids, where loss-of-function mutations are introduced into intestinal stem cells. This helps aid in understanding  how the intestinal environment is maintained and identify mechanisms that may contribute to the development of tumors.

How many cells are available in a fresh leukopak?

STEMCELL Technologies: The cell count varies depending on the donor. As a result, the number of cells that can be isolated after processing the sample cannot be guaranteed. However, STEMCELL Technologies’ minimum cell counts* for product release at the time of shipping are:

  • Full size leukopak: 9 x 109
  • Half-size leukopak: 5 x 109
  • Quarter-size leukopak: 2.5 x 109
  • Tenth-size leukopak: 0.9 x 109

*These minimum cell counts are the acceptance criteria for product release at the time of shipping. Minimum cell counts are the same for frozen leukopaks; however, for frozen leukopaks, cells are counted at the time of freezing. The viability of these cells, as well as the fresh cells after washing, is typically above 90%.

What is the percentage of viable cells with frozen leukopak?

STEMCELL Technologies: Upon cryopreservation, cell viability in the frozen leukopak is greater than 90%. After thawing and the cleanup process, the viability is usually greater than 90%.

Are there any advantages of working with frozen leukopaks compared to fresh leukopaks?

STEMCELL Technologies: Both fresh and frozen leukopaks are ideal starting materials for downstream cell isolation when large numbers of cells are required, reducing the time and reagents needed to process cells of interest. However, frozen leukopaks provide additional convenience to start experiments as per your schedule. Additionally, cell viability may be better preserved in frozen leukopaks during global shipping. Frozen leukopaks can also provide you with access to samples from the same lot at a future date. See the table below for a comparison of fresh and frozen leukopak product features, to help you choose the ideal source of human primary cells for your downstream applications.

Factor Fresh Leukopaks Frozen Leukopaks
Cell Type and Functionality Optimal for experiments requiring high cell functionality or sensitive cell types such as NK cells, dendritic cells, etc. Isolated cells have higher viability as no cryopreservation step is involved Cryopreservation ensures long-term stability of cells with ≥ 90% viability. Cells cryopreserved on the same day as collection typically outperform cells cryopreserved after overnight shipment
Logistics and Availability Inherent risk of shipping delays can impact the quality of cells. Less readily available Eliminate the inherent risk of shipping delays, with no decrease in cell quality. More readily available and can be used for multiple experiments
Downstream Application Ideal for time-sensitive assays and assays requiring optimal cell functionality. Can be processed and cryopreserved upon receipt Suitable for assays requiring a large number of cells or experiments conducted over extended periods of time. Not recommended to re-cryopreserve as second freeze thaw will result in increased cell death

Table I. Factors to Consider When Choosing the Right Leukopak Product for Your Research