Clustered Regularly Interspaced Palindromic Repeats (CRISPR)/Cas9 adaptive immunity against pathogens in bacteria has been adapted for genome editing and applied in zebrafish (Danio rerio) to generate frameshift mutations in protein-coding genes. Although there are methods to detect, quantify and sequence CRISPR/Cas9-induced mutations, identifying mutations in F1 heterozygous fish remains challenging. Additionally, sequencing a mutation and assuming that it causes a frameshift does not prove causality because of possible alternative translation start sites and potential effects of mutations on splicing. This problem is compounded by the relatively few antibodies generated to zebrafish proteins limiting validation at the protein level. To address these issues, we developed a detailed protocol to screen F1 mutation carriers, and clone and sequence identified mutations. For verifying that mutations actually cause frameshifts, we created a fluorescent reporter system that can detect frameshift efficiency based on the cloning of wild-type and mutant cDNA fragments and their expression levels. As proof-of-principle, we applied this strategy to three mutations in pycr1a, chd7 and hace1 genes. In pycr1a gene involved in proline biosynthesis, we identified an insertion of 7 nucleotides that at the mRNA level caused exon skipping. The fluorescent reporter approach revealed effective frameshifting only for 2-nucleotide deletion in chd7, a chromatin remodelling factor gene, suggesting activity of alternative translation sites in the other two. Thus, in addition to providing a protocol for characterizing frameshift mutations in zebrafish, this approach highlights the importance of checking mutations at the mRNA level and verifying their effects on translation by fluorescent reporters when antibody detection of protein loss is not possible.