FGF receptors are required for proper axonal branch targeting in Drosophila.

Proper axonal branch growth and targeting are essential for establishing a hard-wired neural circuit. Here, we examined the role of Fibroblast Growth Factor Receptors (FGFRs) in axonal arbor development using loss of function and overexpression genetic analyses within single neurons. We used the invariant synaptic connectivity patterns of Drosophila mechanosensory neurons with their innate cleaning reflex responses as readouts for errors in synaptic targeting and circuit function. FGFR loss of function resulted in a decrease in axonal branch number and lengths, and overexpression of FGFRs resulted in ectopic branches and increased lengths. FGFR mutants produced stereotyped axonal targeting errors. Both loss of function and overexpression of FGFRs within the mechanosensory neuron decreased the animal's frequency of response to mechanosensory stimulation. Our results indicate that FGFRs promote axonal branch growth and proper branch targeting. Disrupting FGFRs results in miswiring and impaired neural circuit function.

Protein and RNA quantification of multiple genes in single cells.

Single-cell analysis overcomes the problems of cellular heterogeneity by revealing the individual differences between cells in tissue. The current tools used to profile gene expression at the single-cell level are arduous and often require specialized equipment. We have previously developed a technique to quantify protein expression levels in single living cells. Here, we combine quantification of protein expression with absolute measurement of mRNA amounts of the same gene in the same cell, to profile the expression of genes at the transcriptional and translational levels. We show that high heterogeneity exists at both the mRNA and protein levels for multiple genes, even among monoclonal cells. We demonstrate a rapid, straightforward approach to single-cell profiling of RNA and protein production.

Generating stable cell lines with quantifiable protein production using CRISPR/Cas9-mediated knock-in.

Cell lines expressing foreign genes have been widely used to produce a variety of recombinant proteins. However, generating recombinant protein-expressing cell lines is usually a lengthy process and the resulting protein expression levels are often inconsistent. Here, we describe an efficient method for making stable cell lines expressing any recombinant protein of interest in a controllable and quantifiable manner. We integrate transgenes into specific genomic loci using CRISPR/Cas9 such that transgene expression is driven by endogenous promoters to ensure consistent and predictable expression of the recombinant protein. Expression levels can be predetermined by selecting promoters from genes with the desired level of expression. To quantify recombinant protein expression, a protein quantitation reporter (PQR) is incorporated between the endogenous and foreign genes. The PQR allows equimolar production of the endogenous protein, the recombinant protein, and a fluorescent reporter. As a result, expression levels of both the endogenous and recombinant proteins can be continuously monitored using fluorescence.

Homology Directed Knockin of Point Mutations in the Zebrafish tardbp and fus Genes in ALS Using the CRISPR/Cas9 System.

The methodology for site-directed editing of single nucleotides in the vertebrate genome is of considerable interest for research in biology and medicine. The clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein 9 type II (Cas9) system has emerged as a simple and inexpensive tool for editing genomic loci of interest in a variety of animal models. In zebrafish, error-prone non-homologous end joining (NHEJ) has been used as a simple method to disrupt gene function. We sought to develop a method to easily create site-specific SNPs in the zebrafish genome. Here, we report simple methodologies for using CRISPR/Cas9-mediated homology directed repair using single-stranded oligodeoxynucleotide donor templates (ssODN) for site-directed single nucleotide editing, for the first time in two disease-related genes, tardbp and fus.

Quantification of Protein Levels in Single Living Cells.

Accurate measurement of the amount of specific protein a cell produces is important for investigating basic molecular processes. We have developed a technique that allows for quantitation of protein levels in single cells in vivo. This protein quantitation ratioing (PQR) technique uses a genetic tag that produces a stoichiometric ratio of a fluorescent protein reporter and the protein of interest during protein translation. The fluorescence intensity is proportional to the number of molecules produced of the protein of interest and is used to determine the relative amount of protein within the cell. We use PQR to quantify protein expression of different genes using quantitative imaging, electrophysiology, and phenotype. We use genome editing to insert Protein Quantitation Reporters into endogenous genomic loci in three different genomes for quantitation of endogenous protein levels. The PQR technique will allow for a wide range of quantitative experiments examining gene-to-phenotype relationships with greater accuracy.