In recent years, Setaria viridis has been developed as a model plant to better understand the C4 photosynthetic pathway in major crops. With the increasing availability of genomic resources for S. viridis research, highly efficient genome editing technologies are needed to create genetic variation resources for functional genomics. Here, we developed a protoplast assay to rapidly optimize the multiplexed CRISPR/Cas9 system in S. viridis. Targeted mutagenesis efficiency was further improved by an average of 1.4-fold with the exonuclease, Trex2. Distinctive mutation profiles were found in the Cas9_Trex2 samples with 94% of deletions larger than 10 bp, and essentially no insertions at all tested target sites. Further analyses indicated that 52.2% of deletions induced by Cas9_Trex2, as opposed to 3.5% by Cas9 alone, were repaired through microhomology-mediated end joining (MMEJ) rather than the canonical NHEJ DNA repair pathway. Combined with a robust Agrobacterium-mediated transformation method with more than 90% efficiency, the multiplex CRISPR/Cas9_Trex2 system was demonstrated to induce targeted mutations in two tightly linked genes, svDrm1a and svDrm1b, at the frequency ranging from 73% to 100% in T0 plants. These mutations were transmitted to at least 60% of the transgene-free T1 plants with 33% of them containing bi-allelic or homozygous mutations in both genes. This highly efficient multiplex CRISPR/Cas9_Trex2 system makes it possible to create a large mutant resource for S. viridis in a rapid and high throughput manner, and has the potential to be widely applicable in achieving more predictable and deletion-only MMEJ-mediated mutations in many plant species.