Optimization of CRISPR Base Editing Efficiency in Primary Human T Cells

Abstract

Cytosine- and adenine-base editors offer a route to installing precise point mutations in human T cells without the
double-strand breaks that drive translocations and large deletions during conventional Cas9 editing, but their
efficiency in primary, unmodified T cells has lagged well behind what is routinely achieved in cell lines. We set out
to identify which steps of a ribonucleoprotein-free, mRNA-based adenine base editing workflow most limit editing
rate in primary CD3+ T cells, and to build a protocol that closes that gap without sacrificing viability or the resting,
minimally activated state that many downstream cell-therapy applications require. Using ABE8e messenger RNA
co-electroporated with a chemically modified single guide RNA targeting a splice-donor site in PDCD1, we
systematically varied activation time before electroporation, mRNA and sgRNA dose, cuvette electroporation
programme, and post-electroporation recovery conditions across twelve donors. Editing efficiency, measured by
amplicon deep sequencing, rose from a baseline of 21.4% under a standard 48-hour activation protocol to 68.7%
when activation was shortened to 20 hours and mRNA dose was raised to 3 micrograms per million cells, without
a corresponding drop in seven-day viability (86.3% versus 84.1% for baseline). Flow cytometry confirmed loss of
surface PD-1 expression in edited populations proportional to the sequencing-based editing rate, and edited cells
retained normal proliferative capacity across three rounds of anti-CD3/CD28 restimulation. Off-target analysis by
CIRCLE-seq-nominated site sequencing found no editing above the sequencing error floor at any of eighteen
candidate loci. These results identify activation timing as an underappreciated but highly actionable variable in
primary T cell base editing and provide a practical, donor-generalisable protocol for researchers developing
base-edited cellular therapies.