SUMMARY A scalable and high-throughput method to identify precise subcellular localization of endogenous proteins is essential for integrative understanding of a cell at the molecular level. Here, we developed a simple and generalizable technique to image endogenous proteins with high specificity, resolution and contrast in single cells in mammalian brain tissue. The technique, termed SLENDR, uses in vivo genome editing to insert a sequence encoding an epitope tag or a fluorescent protein to a gene of interest by CRISPR-Cas9-mediated homology-directed repair (HDR). Single-cell, HDR-mediated genome editing was achieved by delivering the editing machinery to dividing neuronal progenitors through in utero electroporation. We demonstrate that SLENDR allows rapid determination of the localization and dynamics of many endogenous proteins in various cell types, regions and ages in the brain. Thus, SLENDR provides a high-throughput platform to map the subcellular localization of endogenous proteins with the resolution of micro- to nanometers in the brain.

SUMMARY Precise genome editing via homology-directed repair (HDR) in targeted cells, particularly in vivo, provides an invaluable tool for biomedical research. However, HDR has been considered to be largely restricted to dividing cells, making it challenging to apply the technique in postmitotic neurons. Here we show that precise genome editing via HDR is possible in mature postmitotic neurons as well as mitotic cells in mice brain by combining CRISPR-Cas9-mediated DNA cleavage and the efficient delivery of donor template with adeno-associated virus (AAV). Using this strategy, we achieved efficient tagging of endogenous proteins in primary and organotypic cultures in vitro and developing, adult, aged, and pathological brains in vivo. Thus, AAV- and CRISPR-Cas9-mediated HDR will be broadly useful for precise genome editing in basic and translational neuroscience.

Neural circuits are shaped by elimination of early-formed redundant synapses during postnatal development. Retrograde signaling from postsynaptic cells regulates synapse elimination. In this work, we identified semaphorins, a family of versatile cell recognition molecules, as retrograde signals for elimination of redundant climbing fiber to Purkinje cell synapses in developing mouse cerebellum. Knockdown of Sema3A, a secreted semaphorin, in Purkinje cells or its receptor in climbing fibers accelerated synapse elimination during postnatal day 8 (P8) to P18. Conversely, knockdown of Sema7A, a membrane-anchored semaphorin, in Purkinje cells or either of its two receptors in climbing fibers impaired synapse elimination after P15. The effect of Sema7A involves signaling by metabotropic glutamate receptor 1, a canonical pathway for climbing fiber synapse elimination. These findings define how semaphorins retrogradely regulate multiple processes of synapse elimination.

Neural circuits are shaped by activity-dependent elimination of redundant synapses during postnatal development. In many systems, postsynaptic activity is known to be crucial, but the precise mechanisms remain elusive. Here, we report that the immediate early gene Arc/Arg3.1 mediates elimination of surplus climbing fiber (CF) to Purkinje cell (PC) synapses in the developing cerebellum. CF synapse elimination was accelerated when activity of channelrhodopsin-2-expressing PCs was elevated by 2-day photostimulation. This acceleration was suppressed by PC-specific knockdown of either the P/Q-type voltage-dependent Ca(2+) channels (VDCCs) or Arc. PC-specific Arc knockdown had no appreciable effect until around postnatal day 11 but significantly impaired CF synapse elimination thereafter, leaving redundant CF terminals on PC somata. The effect of Arc knockdown was occluded by simultaneous knockdown of P/Q-type VDCCs in PCs. We conclude that Arc mediates the final stage of CF synapse elimination downstream of P/Q-type VDCCs by removing CF synapses from PC somata.