Chimeric antigen receptor engineered T cell (CAR-T) immunotherapy has shown efficacy against a subset of hematological malignancies, yet its autologous nature and ineffectiveness against epithelial and solid cancers limit widespread application. To overcome these limitations, targeted nucleases have been used to disrupt checkpoint inhibitors and genes involved in alloreactivity. However, the production of allogeneic, off-the-shelf T cells with enhanced function requires multiplex genome editing strategies that risk off-target effects, chromosomal rearrangements, and genotoxicity due to simultaneous double-strand break (DSB) induction at multiple loci. Moreover, it has been well documented that DSBs are toxic lesions that can drive genetic instability. Alternatively, CRISPR/Cas9 base editors afford programmable enzymatic nucleotide conversion at targeted loci without induction of DSBs. We reasoned this technology could be used to knockout gene function in human T cells while minimizing safety concerns associated with current nuclease platforms. Through systematic reagent and dose optimization, we demonstrate highly efficient multiplex base editing and consequent protein knockout in primary human T cells at loci relevant to the generation of allogeneic CAR-T cells including the T cell receptor alpha constant (TRAC) locus, beta-2 microglobulin (B2M), and programmed cell death 1 (PDCD1). Multiplex base edited T cells equipped with a CD19 CAR killed target cells more efficiently; and importantly, both DSB induction and translocation frequency were greatly reduced compared to cells engineered with Cas9 nuclease. Collectively, our results establish a novel multiplex gene editing platform to enhance both the safety and efficacy of engineered T cell-based immunotherapies.