English
Immunotherapy has reshaped cancer treatment by harnessing the ability of T cells to selectively recognize and kill malignant cells. Despite remarkable clinical successes, durable responses remain limited to a minority of patients, underscoring the need to develop novel and more effective therapies. Genome engineering offers new opportunities to enhance T cells therapies, and CRISPR-Cas9 screens have become an indispensable tool for discovering genetic determinants of T cell function. However, existing screening workflows rely on indirect readouts of T cell function, such as proliferation, expression of activation markers, or cytokine secretion, and are not suited to directly measure T cell killing capacity. To overcome this limitation, we developed a droplet-based microfluidic platform that links CRISPR gene editing to the direct assessment of T cell cytotoxicity at single-cell resolution. In this system, CRISPR edited human CD8+ memory T cells are singularly co-encapsulated with target cells in picoliter water-in-oil droplets, creating thousands of microreactors that enable rapid, functional readouts of cytotoxic activity. Droplets containing T cells that successfully killed their target are selectively isolated by fluorescent-activated droplet sorting, allowing retrieval of the corresponding sgRNAs for enrichment analysis by next-generation sequencing. Leveraging this platform, we identified both positive and negative regulators of T cell cytotoxicity, including known factors such as RASA2, PTEN and FOXO1, and the less characterized cytotoxicity regulators RPTOR and AFAP1L2, whose deletion strengthened early cytotoxic responses and improved antitumor efficacy against liquid and solid tumors. This work establishes a high-throughput platform for functional single-cell CRISPR screening of T cell cytotoxicity. In addition to uncovering genetic drivers of T cell cytotoxicity, the developed platform enables systematic investigation of non-autonomous phenotypes, thereby expanding the CRISPR screening toolbox.