Though tumor targeting using chimeric receptors (CAR) engineered to bind tumor-associated surface proteins such as CD19 in B cells have demonstrated remarkable efficacy in cancer, this therapy has hit a glass ceiling w.r.t. initial and durable remissions. Inclusion of a costimulatory domain in the CAR design have solved much of limited clinical efficacy issues, some challenges remained. We have recently demonstrated that T cell-intrinsic mechanisms solved part of the glass ceiling conundrum in leukemia, we recently demonstrated that primary chronic lymphocytic leukemia cells elicit a blunted response of second generation CAR T cells, even when using normal donor CAR-engineered T cells. In my talk I will highlight our discoveries in T cell- and tumor-intrinsic determinants of anti-CD19 CAR T cell responses in this disease.

Learning Objectives:

1. Classify the Chimeric Antigen Receptor (CAR)-engineered T cells have dramatically altered the clinical outlook for many blood cancers

2. Identify that with the discovery of costimulation as a critical component of an effective T cell responses and the inclusion of such domains in CAR design we have overcome previous significant roadblocks, but encountered new challenges along the way

3. Recognize the part of what explains response and resistance to this therapy is T cell-intrinsic; however, even the most efficacious CAR showed ineffective against primary tumor cells

4. Summarize the highlights of some of our discoveries in elucidating T cell- and tumor-cell-intrinsic determinants of CAR T cell efficacy

Genome editing technologies have enabled the precise manipulation of DNA sequences at targeted sites to achieve therapeutic effects. Engineered endonucleases like CRISPR-Cas9 institute site-specific DNA breaks to either knock out a gene or enhance the homology directed repair (HDR) based gene correction with an intact donor template. High-fidelity editing, multiplexing capacity, and the ease of implementing the CRISPR-Cas9 system have expanded the scope of programmable genetic manipulations to include simultaneous deletions or insertions of multiple DNA sequences in a single round of mutagenesis. Gene editing has many applications in basic and biomedical research, including disease modeling. A promising cell therapy approach for hemoglobinopathies or cancer immunotherapy involves the ex vivo gene editing of autologous hematopoietic stem cells (HSCs) or T cells before administering patients. For hemoglobinopathies, gene editing tools like CRISPR-Cas9 can be used to install a double stranded break to either knock out a gene like BCL11A or enhance the knock-in with an intact donor DNA template to correct the mutation in the beta subunit of the hemoglobin gene (HBB). Cancer immunotherapy applications use CRISPR-Cas9 to knock out the TCR locus or checkpoint modulators to improve the efficacy and clinical outcomes of the living drug. Here, we describe the use of a clinically validated, regulatory compliant, scalable electroporation platform for the high efficiency, low toxicity gene editing of hCD34+ hematopoietic stem cells (HSCs), T cells, and iPSC cells at a clinical scale for preclinical evaluation and commercial production of cell therapy products for Sickle Cell Disease, TCR therapy or in disease modeling.

Learning Objectives:

1. llustrate therapeutic level of gene correction of the SCD mutation in patient derived CD34+ HSCs cells by electroporation of CRISPR-Cas9 gene editing tools. The gene edited cells with homology directed repair (HDR) engrafted and were detectable for a significant period of time in mouse xenograft models.

2. Paraphrase the optimization of the RNP mediated genome editing using CRISPR-Cas9 and a ssODN template to engineer iPSCs for treatment of diseases like DMD and Miyoshi Myopathy.

3. Describe a clinical trial where patients were infused with an autologous CRISPR-Cas9 genetically edited CD34+ HSC commercial product aimed knocking out the BCL11A enhancer, thereby reactivating the production of fetal hemoglobin to alleviate the debilitating symptoms of SCD and beta thalassemia.

Learning Objectives:

1. Define the background introduction to AAV gene therapy

2. Recall the novel methods to build AAV capsid libraries for in vivo screening through rational design and directed evolution

3. Identify the tissue specific promoter and enhancer engineering to optimize transgene expression

4. Recall the design of a novel AAV-microdystrophin vector for Duchenne Muscular Dystrophy gene therapy

Learning Objectives:

1. Summarize delivery vehicles for the Cas9 RNP

2. Identify Gene editing after a direct tissue injection

3. Indicate new linkers for conjugating proteins to polymers

Biocatalysis, protein engineering, and flow chemistry are key enabling technologies that can yield drastically shorter and greener chemical processes. We sought to leverage these technologies to realize an improved synthesis. A cheap and green chemical commodity was identified as a starting material that allowed the generation of a key intermediate in the synthesis in a stepwise reaction. As an initial proof of concept, an enzyme-catalyzed reaction was demonstrated with modest yields and selectivity. However, the initial enzyme and process would not be sufficient to support large scale synthesis. To improve the synthesis, we engaged in enzyme evolution, enzyme immobilization, and flow chemistry. Through several rounds of evolution the enzyme was evolved to reach high conversion, selectivity and stability in the presence of organic solvents.

Learning Objectives:

1. Know protein engineering is often necessary to improve desired properties of an enzyme to be used in commercial manufacturing routes.

2. Recognize enzyme immobilization along with protein engineering can be effectively used to drive high product conversion in organic solvents without loosing enzymatic activity and without the need of enzymatic removal during reaction work up.  

AAV peptide display libraries allow for the generation of novel variants capable of high-level transduction. We previously described an AAV peptide display library combined with a sensitive transduction reporter, iTransduce, that enabled the discovery of potent capsids in the mouse brain after only two rounds of selection via intravenous injection. Analogous to drug repurposing, AAV capsids can be used for gene delivery applications other than originally intended. With that in mind, we tested two of the capsids, AAV-F and AAV-S, from our selection in mice to transduce non-human primate (NHP) spinal cord and cochlea, respectively. We found that intrathecally injected AAV-F mediated higher expression in motor neurons and interneurons compared to AAV9 in the thoracic and lumbar regions, despite lower numbers of genomes per cell in AAV-F-treated animals. AAV-S was tested in the inner ears of mice and cynomolgus macaques, and in both cases drove robust, high-level transduction throughout the cochlea. Further, AAV-S was able to rescue the deafness phenotype of a mouse model of Clarin-1 related Usher Syndrome. Overall, this work serves to highlight that AAV variants selected from our library system can translate effectively to clinically relevant large animals, and provides further evidence for the benefits of capsid repurposing.

Learning Objectives:

1. Recall that AAV libraries allow for the selection of potent new variants; however, most of these are only extensively tested in mouse

2. Recognize the testing of two previously selected variants, AAV-F and AAV-S, in cynomolgus macaques

3. Restate how AAV-F expressed well in the spinal cord even at low doses, with slightly higher levels of expression compared to AAV9 despite lower per-cell genome numbers.

4. Indicate that AAV-S drove high level expression in the inner ears of mice and macaques, and was able to robustly rescue deafness in a mouse model of Clarin-1 Usher syndrome

Conventional CRISPR–Cas systems maintain genomic integrity by leveraging guide RNAs for the nuclease-dependent degradation of mobile genetic elements, including plasmids and viruses. Here we describe a remarkable inversion of this paradigm, in which bacterial transposons have coopted nuclease-deficient CRISPR–Cas systems to catalyze RNA-guided integration of mobile genetic elements into the genome. Programmable transposition requires CRISPR- and transposon-associated molecular machineries, including a novel co-complex between Cascade and the transposition protein TniQ. Donor DNA integration occurs at a fixed distance downstream of target DNA sequences, accommodates variable length genetic payloads, and functions robustly in diverse bacterial species. Deep sequencing experiments reveal highly specific, genome-wide DNA integration across dozens of unique target sites. The discovery of a fully programmable, RNA-guided transposase lays the foundation for kilobase-scale genome engineering that obviates the requirements for DNA double-strand breaks and homologous recombination.

Learning Objectives:

1. Recall CRISPR-Cas systems are naturally repurposes for non-canonical functions other than cleaving RNA and DNA

2. Indicate CRISPR RNA-guided transposases integrate donor DNA without generating DNA double-strand breaks

3. Explain RNA-guided DNA integration proceeds with high-fidelity and readily accommodates kilobase-size genetic payloads

Genome Engineering allows the easy manipulation of genomes down to the nucleotide level. CRISPR is a revolutionizing tool for generating custom edited cell and pre-clinical animal models for research. Targeted deep sequencing enables the detection and quantification of low-frequency editing events. However, the large amounts of data generated by targeted deep sequencing can be difficult to interpret and quickly analyze. We have developed a Python-based computer program called CRIS.py that allows the easy analysis of multiple types of editing events. We will show examples of how rapid deep sequence analysis has guided experimental design leading to high-efficiency genome editing in a broad range of applications. Additionally, we will discuss best practices for creating knockin cell and animal models including large insertions using long ssDNA.

Learning Objectives:

1. Explain how NGS analysis enables detection and quantification of CRISPR editing outcomes on a diverse range of sample inputs including cell pools, clonal populations, and animal tissues

2. Recognize CRIS.py is a user friendly NGS analysis pipeline that is open source

3. Discuss best practices for creating knock in cell lines 4. Identify that NGS can be used to genotype both small and large insertion events

We have harnessed homologous recombination to modify the genome of human stem cells with high frequencies (>40%). By designing the recombination donor in different ways, we can achieve a wide range of targeted integration outcomes including creating single nucleotide changes, inserting one gene into another gene, simultaneously inserting two genes into the genome, or even replacing one gene with another. In this talk, I will discuss some of these applications to engineering human stem cells to treat patients.

Learning Objectives:

1. Discuss homologous recombination is a powerful and efficient mechanism to engineer cells using the principles of synthetic biology.

2. Review donor design for the recombination process is an essential part of harnessing the broad potential of homologous recombination based genome editing.

Though AsCas12a fills a crucial gap in the current genome editing toolbox, it exhibits relatively poor editing efficiency, restricting its overall utility. In collaboration with Integrated DNA Technologies, we showed that this engineered variant we refer to as AsCas12a Ultra, increased editing efficiency to nearly 100% at all sites examined in HSPCs, iPSCs, T cells, and NK cells. We showed that AsCas12a Ultra maintains high on-target specificity thereby mitigating the risk for off-target editing and making it ideal for complex therapeutic genome editing applications. We achieved simultaneous targeting of three clinically relevant genes in T cells at >90% efficiency and demonstrated transgene knock-in efficiencies of up to 60%. We demonstrate site-specific knock-in of a CAR in NK cells, which afforded enhanced anti-tumor NK cell recognition, potentially enabling the next generation of allogeneic cell-based therapies in oncology. AsCas12a Ultra is an advanced CRISPR nuclease with significant advantages in basic research and in the production of gene edited cell medicines.

Learning Objectives:

1. Identify the engineered AsCas12a Ultra nuclease is commercially available to researchers from Integrated DNA Technologies

2. Recall this engineered AsCas12a nuclease forms the basis for EDIT-301, an experimental treatment for sickle cell disease which is currently being tested in the clinic

3. Recall this engineered AsCas12a nuclease breaks the observed paradigm where most natural and engineered CRISPR nuclease variants that achieve very high specificity come with a cost to on-target editing activity in most clinically relevant cell types

Gene editing technologies such as CRISPR/Cas9 have greater flexibility and high efficiency. CRISPR technology enables targeted insertion of transgenes at the desired locus and has the ability to multiplex knocking out of multiple genes. These features provide a great opportunity to use this technology to develop next-generation CAR T cells. The popular method for the generation of CART cell drug product is delivery of CAR and its integration into the T cell genome using viral vectors. Lentivirus or gamma retrovirus-based CAR delivery was used in multiple CART cell productions. CART cell production using viral vectors to deliver the transgene results in the random insertion of CAR sequence across the whole genome. In addition to random CAR insertion viral vectors also deliver partial viral sequences into the cell genome. These things together could disrupt essential gene functions and affect surrounding functional gene expression. CRISPR/Cas9 gene-editing technology can achieve targeted insertion of CAR construct sequence in the desired locus such as T cell receptor locus (TRAC) or any genomic safe harbor sites. The transition from viral vectors to CRISPR/Cas9 based gene editing requires multiple reagents that are GMP compatible. A lot of progress was achieved in the commercial availability of CRISPR/Cas9 reagents in recent years for the generation of cGMP CART drug products. However, there are multiple issues that require careful attention in process development to achieve a reliable cGMP process for CRISPR/Cas9 based cell therapy drug product. These include gene knockout and knock-in efficiency, off target and on target effects, and the ability to achieve the desired quantity of the drug product. A study on the feasibility of using CRISPR/cas9 gene-editing technology for autologous CART drug production and the relevance of this technology for the generation of safe drugs for the patients will be discussed.

Learning Objectives:

1. Describe CART cell production process

2. Explain CRISPR/Cas9 gene editing

Learning Objectives:

1. Recall engineered CRISPR-Cas variants overcome natural limitations of wild-type enzymes

2. Know the genome is now nearly completely editable

3. Recognize minimal PAM CRISPR-Cas enzymes enable previously impossible edits

T-cells targeting shared oncogenic mutations can induce durable tumor regression in epithelial cancer patients. Such T cells can be detected in tumor-infiltrating lymphocytes and peripheral blood of patients with the common metastatic epithelial cancers. In recent years, we and others have worked toward developing libraries of T-cell receptors targeting oncogenic mutations such as KRAS and TP53. Such T-cell receptor libraries offer an opportunity to treat patients with autologous or allogeneic T-cells genetically engineered to express selected T-cell receptors. Although attractive, such clinical application of T-cell receptor libraries should be accompanied by innovative genetic engineering platforms allowing cheaper, faster, and more flexible manufacturing solutions. This talk will discuss the methods to isolate, clone, and validate T-cell receptors targeting oncogenic mutations, the therapeutic potential of such libraries, and the genetic engineering platforms currently used for clinical manufacturing. We will also address the genetic engineering tools needed to allow such an approach to become widely and commercially applicable.

Learning Objectives:

1. Discuss methods to isolate, clone, and validate T-cell receptors targeting oncogenic mutations.

2. Define the therapeutic potential of TCR libraries targeting oncogenic mutations.

3. Indicate the current and future genetic engineering platforms needed to support clinical applications using off-the-shelf TCR libraries.

Human T cells are critical effectors of immune protection from infections, autoimmune pathology, and cancer immunotherapy. We use CRISPR-mediated gene editing in primary human T cells to systematically identify genetic targets that modulate the functions of T cells in contexts ranging from immunosuppression to cancer killing. By developing and applying CRISPR based methodologies such as pooled knock-in screening, CRISPR activation, and CRISPR interference, we are pinpointing the regulatory networks controlling T cell phenotypes as well as synthetic genetic programs that can be engineered into T cells to improve their utility as cell-based therapies for disease. We are working towards a range of genetically engineered cell therapies for cancer, autoimmunity, infections and other diseases.

Learning Objectives:

1. Discuss current approaches for CRISPR genome editing in primary human T cells

2. Review progress toward clinical genome editing with non-viral knockin strategies

3. Discuss application of genome-wide screens as discovery platform for enhanced cellular therapies