(H-291) Optimization of CRISPR/Cas-based programmable DNA methylation in human primary T cells for regulation of immunological exhaustion

Introduction:: CRISPR/Cas-based technologies have emerged as powerful methods to engineer the human epigenome. These rapidly maturing capabilities are now being harnessed to stimulate commercial opportunities as well as discoveries and innovations in basic and translational research. For instance, we and others have shown that engineered molecules consisting of deactivated Cas9 (dCas9) fused to human DNA methyltransferase or demethylase enzymes can be used to modulate methylation levels at endogenous human loci. Our team has recently shown that Tuberculosis (TB) patients display dampened immunity and generalized immunological exhaustion that is linked to DNA hypermethylation at immune signaling pathway genes. Despite these correlations, the molecular mechanisms by which DNA hypermethylation can elicit inhibited host immunity remain unclear. Here we developed an optimized strategy for lentiviral delivery of dCas9-based DNA methyltransferases and demethylases to human cell lines and primary T cells and to dissect the role that DNA methylation plays at key immune genes in human T cells and how this epigenetic regulatory mechanism can be engineered for therapeutic benefits.

Materials and Methods:: Lentiviral transduction of human cell lines and primary T cells was optimized and used to deliver multiplexed gRNAs and dCas9-based epigenome editing tools. RT-qPCR was used to evaluate modulation of targeted immune regulatory gene expression levels. Bisulfite PCR followed by deep sequencing allowed us to focus on how targeting of specific immune loci enabled the synthetic deposition or erasure of DNA methylation marks, and aided in establishing the causal relationship between DNA methylation and T cell phenotypes. Immune cell co-culture killing assays, flow cytometry, and ELISA will be used for functional benchmarks in this study. 

Results, Conclusions, and Discussions:: Our results show that certain design considerations are critical to the viral delivery of some engineered epigenetic effectors. By taking these considerations into account, we were able to improve the titers of viral vectors, and in turn, improve viral transduction efficiencies in primary T cells. Building upon this optimization and to further clarify the role of focal DNA hypermethylation in TB-induced immunological exhaustion, we applied an optimized epigenome editing strategy to target DNA methylation to multiple loci using multiplexed gRNAs in tandem with dCas9-DNMT3A3L at a critical testbed locus. Collectively, our studies show that engineered DNA methylation at key endogenous human genes results in decreased gene expression and suggest that DNA methylation is a critical driver of dampened immune responses in TB patients.

Tuberculosis (TB) infection results in 10 million infections and 1.5 million deaths a year, making TB the world’s leading infectious cause of mortality. Therefore, new strategies to address the pathological consequences of TB infection are urgently needed. Our work combines the innovative use of engineered CRISPR/Cas-based technologies and unique mechanistic insights from TB-patient derived cells. Our findings highlight the promise that epigenome editing in primary human cell types holds and increase our understanding of how aberrant DNA methylation controls the balance between health and disease cell states. Specifically, our data suggests that targeted DNA hypermethylation is achieved in different levels depending on the targeted loci and consequently it can produce different levels of gene silencing. Importantly, aberrant DNA methylation is linked with numerous human diseases and exhausted immunological states, thus while our study here is focused on the underlying epigenetic mechanisms of immune exhaustion in TB, our results set a foundation to better understand and engineer DNA methylation in a wide range of cell states and pathologies.

Acknowledgements (Optional): : This work was supported by the Cancer Prevention & Research Institute of Texas (CPRIT; RR170030 to I.B.H), the John S. Dunn Foundation (to I.B.H. and A.D.), the National Institute of General Medical Sciences (NIGMS; R35GM143532 to I.B.H.), and the National Cancer Institute (NCI) through a SPORE Award in Lymphoma Developmental Research Project (P50CA126752; PI Heslop).
The authors thank all members of the Hilton and DiNardo laboratories for helpful discussions and feedback.

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