Neuroscience

Accidental wounding of the peripheral nervous system leads to acute neural dysfunction. Normally, chronic deficits are overcome because peripheral nerves naturally regenerate. However, various genetic and metabolic defects can impair their natural regenerative capacity, which may be due to neuron-extrinsic mechanisms. Therefore, characterizing the behavior of multiple cells during nerve injury and repair in vivo is a pressing need in regenerative medicine. Here, we detail a method for precise wounding of sensory axons in zebrafish, followed by high-resolution in toto long-term quantitative videomicroscopy of neurons, Schwann cells, and macrophages. This protocol can be easily adapted to study the effects of targeted genetic or metabolic disruptions in zebrafish and other suitable organisms, as well as for screening pharmacological agents with therapeutic potential.

Graphical overview

The trafficking and sorting of proteins through the secretory-endolysosomal system is critical for the proper functioning of neurons. Defects in steps of these pathways are associated with neuronal toxicity in various neurodegenerative disorders. The prion protein (PrP) is a glycosylphosphatidylinositol (GPI)-anchored protein that follows the secretory pathway before reaching the cell surface. Following endocytosis from the cell surface, PrP sorts into endosomes and lysosomes for further recycling and degradation, respectively. A few detailed protocols using drug treatments and fluorescent dyes have previously allowed the tracking of PrP trafficking routes in real time in non-neuronal cells. Here, we present a protocol optimized for primary neurons that aims to monitor and/or manipulate the trafficking and sorting of PrP particles at several steps during their secretory-endolysosomal itineraries, including (a) ER export, (b) endocytosis, (c) lysosomal degradation, and (d) accumulation in axonal endolysosomes. These primary neuron live assays allow for the robust quantitation of accumulation and/or degradation of PrP or of other membrane-associated proteins that transition from the ER to the Golgi via the cell surface.

Graphical abstract

Zebrafish is an excellent model to study vertebrate neurobiology, but its synaptic components that mediate and regulate fast electrical synaptic transmission are largely unidentified. Here, we describe methods to solubilize and immunoprecipitate adult zebrafish brain homogenate under conditions to preserve electrical synapse protein complexes. The methods presented are well-suited to probe electrical synapse immunocomplexes, and potentially other brain-derived immunocomplexes, for candidate interactors from zebrafish brain.

Eukaryotic cells utilize sub-cellular compartmentalization to restrict reaction components within a defined localization to perform specified biological functions. One way to achieve this is via membrane enclosure; however, many compartments are not bounded with lipid membrane bilayers. In the past few years, it has been increasingly recognized that molecular components in non- or semi-membrane-bound compartments might form biological condensates autonomously (i.e., without requirement of energy input) once threshold concentrations are reached, via a physical chemistry process known as liquid–liquid phase separation. Molecular components within these compartments are stably maintained at high concentrations and separated from the surrounding diluted solution without the need for a physical barrier. Biochemical reconstitution using recombinantly purified proteins has served as an important tool for understanding organizational principles behind these biological condensates. Common techniques include turbidity measurement, fluorescence imaging of 3D droplets, and atomic force microscopy measurements of condensate droplets. Nevertheless, many molecular compartments are semi-membrane-bound with one side attached to the plasma membrane and the other side exposed to the cytoplasm and/or attached to the cytoskeleton; therefore, reconstitution in 3D solution cannot fully recapture their physiological configuration. Here, we utilize a postsynaptic density minimal system to demonstrate that biochemical reconstitution can be applied on supported lipid bilayer (SLB); we have also incorporated actin cytoskeleton into the reconstitution system to mimic the molecular organization in postsynaptic termini. The same system could be adapted to study other membrane-proximal, cytoskeleton-supported condensations.

The activity of numerous autophagy-related proteins depends on their phosphorylation status, which places importance on understanding the responsible kinases and phosphatases. Great progress has been made in identifying kinases regulating autophagy, but much less is known about the phosphatases counteracting their function. Genetic screens and modern proteomic approaches provide powerful tools to identify candidate phosphatases, but further experiments are required to assign direct roles for candidates. We have devised a novel protocol to test the role of purified phosphatases in dephosphorylating specific targets in situ. This approach has the potential to visualize context-specific differences in target dephosphorylation that are not easily detected by lysate-based approaches such as Western blots.

Graphical abstract:

Mitochondrial dysfunction is associated with perturbations in the cellular oxidative status, changes in energy production and metabolic rate, and the onset of pathological processes. Classic methods of assessing mitochondrial dysfunction rely on indirect measures, such as evaluating mitochondrial DNA copy numbers, or direct but more costly and skilled techniques, such as electron microscopy. The protocol presented here was recently implemented to evaluate mitochondrial dysfunction in response to insecticide exposure in Drosophila melanogaster larvae, and it relies on the use of a previously established MitoTimer mutant strain. MitoTimer is a genetically engineered mitochondrial protein that shows green fluorescence when newly synthetized, irreversibly turning into red as mitochondria age. The protocol described here allows for the easy and direct assessment of shifts in mitochondrial turnover, with tissue-specific accuracy. This protocol can be adapted to assess changes in mitochondrial turnover in response to drugs, rearing conditions, and/or mutations in larva, pupa, or adult fruit flies.

The quantification of labeled cells in tissue sections is crucial to the advancement of biological knowledge. Traditionally, this was a tedious process, requiring hours of careful manual counting in small portions of a larger tissue section. To overcome this, many automated methods for cell analysis have been developed. Recent advances in whole slide scanning technologies have provided the means to image cells in entire tissue sections. However, common automated analysis tools do not have the capacity to deal with the large image files produced. Herein, we present a protocol for the quantification of two fluorescently labeled cell populations, namely pericytes and microglia, in whole brain tissue sections. This protocol uses custom-made scripts within the open source software QuPath to provide a framework for the careful optimization and validation of automated cell detection parameters. Images obtained from a whole-slide scanner are first loaded into a QuPath project. Manual counts are performed on small sample regions to optimize cell detection parameters prior to automated quantification of cells across entire brain regions. Even though we have quantified pericytes and microglia, any fluorescently labeled cell with clear labeling in and around the nucleus can be analyzed using these methods. This protocol provides a user-friendly and cost-effective framework for the automated analysis of whole tissue sections.

Mammalian tissues are highly heterogenous and complex, posing a challenge in understanding the molecular mechanisms regulating protein expression within various tissues. Recent studies have shown that translation at the level of the ribosome is highly regulated, and can vary independently of gene expression observed at a transcriptome level, as well as between cell populations, contributing to the diversity of mammalian tissues. Earlier methods that analyzed gene expression at the level of translation, such as polysomal- or ribosomal-profiling, required large amounts of starting material to isolate enough RNA for analysis by microarray or RNA-sequencing. Thus, rare or less abundant cell types within tissues were not able to be properly studied with these methods. Translating ribosome affinity purification (TRAP) utilizes the incorporation of an eGFP-affinity tag on the large ribosome subunit, driven by expression of cell-type specific Cre-lox promoters, to allow for identification and capture of transcripts from actively translating ribosomes in a cell-specific manner. As a result, TRAP offers a unique opportunity to evaluate the entire mRNA translation profile within a specific cell type, and increase our understanding regarding the cellular complexity of mammalian tissues.

Graphical abstract:

Schematic demonstrating TRAP protocol for identifying ribosome-bound transcripts specifically within cerebellar Purkinje cells.

G-protein coupled signaling pathways are organized into multi-protein complexes called signalosomes that are located within and on cellular membranes. We describe the use of silica nanoparticles coated with a unilamellar phospholipid bilayer (lipobeads) to reconstitute the activated photoreceptor G-protein α-subunit (Gtα*) with its cognate effector (phosphodiesterase-6; PDE6) for biochemical and structural studies of the activation mechanism regulating this GPCR signaling pathway. Lipobeads are prepared by resuspending dried-down phospholipid mixtures with monodisperse 70 nm silica particles, followed by extrusion through a 100 nm membrane filter. This uniform and supported liposomal preparation is easily sedimented, permitting the separation of soluble from membrane-associated proteins. Upon loading lipobeads with Gtα* and PDE6, we find that activation of PDE6 catalysis by Gtα* occurs much more efficiently than in the absence of membranes. Chemical cross-linking of membrane-confined proteins allows detection of changes in protein-protein interactions, resulting from G-protein activation of PDE6. The advantages of using lipobeads over partially purified membrane preparations or traditional liposomal preparations are generally applicable to the study of other membrane-confined signal transduction pathways.