Neuroscience

The centrosome governs many pan-cellular processes including cell division, migration, and cilium formation. However, very little is known about its cell type-specific protein composition and the sub-organellar domains where these protein interactions take place. Here, we outline a protocol for the spatial interrogation of the centrosome proteome in human cells, such as those differentiated from induced pluripotent stem cells (iPSCs), through co-immunoprecipitation of protein complexes around selected baits that are known to reside at different structural parts of the centrosome, followed by mass spectrometry. The protocol describes expansion and differentiation of human iPSCs to dorsal forebrain neural progenitors and cortical projection neurons, harvesting and lysis of cells for protein isolation, co-immunoprecipitation with antibodies against selected bait proteins, preparation for mass spectrometry, processing the mass spectrometry output files using MaxQuant software, and statistical analysis using Perseus software to identify the enriched proteins by each bait. Given the large number of cells needed for the isolation of centrosome proteins, this protocol can be scaled up or down by modifying the number of bait proteins and can also be carried out in batches. It can potentially be adapted for other cell types, organelles, and species as well.

Graphical overview

An overview of the protocol for analyzing the spatial protein composition of the centrosome in human induced pluripotent stem cell (iPSC)-derived neural cells. ① Human iPSCs are expanded, which serve as the starting cell population for the neural induction (Sections A, B, and C in Procedure). ② Neurons are induced and differentiated for 40 days (Section D in Procedure), in at least four biological replicates. ③ Total protein is isolated either at 15th or 40th day of differentiation, for neural stem cells and neurons, respectively (Sections E and F in Procedure). ④ Selected bait proteins are immunoprecipitated using the respective antibodies (Sections G and H in Procedure). ⑤ Co-immunoprecipitated samples are analyzed with mass spectrometry (Section I in Procedure). ⑥ Mass spectrometry output (.RAW) files are processed using MaxQuant software to calculate intensities (Section A in Data analysis). ⑦ The resulting data are pre-processed, filtered, and statistically analyzed using Perseus and R software (Sections B and C in Data analysis) ⑧ Further analysis is done using software or web tools such as Cytoscape or STRING to gain biological insights (Sections D and E in Data analysis).

Parkinson’s disease is a devastating neurodegenerative disorder affecting 2-3% of the population over 65 years of age. There is currently no disease-modifying treatment. One of the predominant pathological features of Parkinson’s disease is mitochondrial dysfunction, and much work has aimed to identify therapeutic compounds which can restore the disrupted mitochondrial physiology. However, modelling mitochondrial dysfunction in a disease-relevant model, suitable for screening large compound libraries for ameliorative effects, represents a considerable challenge. Primary patient derived cells, SHSY-5Y cells and in vivo models of Parkinson’s disease have been utilized extensively to study the contribution of mitochondrial dysfunction in Parkinson’s. Indeed many studies have utilized LUHMES cells to study Parkinson’s disease, however LUHMES cells have not been used as a compound screening model for PD-associated mitochondrial dysfunction previously, despite possessing several advantages compared to other frequently used models, such as rapid differentiation and high uniformity (e.g., in contrast to iPSC-derived neurons), and relevant physiology as human mesencephalic tissue capable of differentiating into dopaminergic-like neurons that highly express characteristic markers. After previously generating GFP⁺-LUHMES cells to model metabolic dysfunction, we report this protocol using GFP⁺-LUHMES cells for high-throughput compound screening in a restoration model of PD-associated mitochondrial dysfunction. This protocol describes the use of a robust and reproducible toxin-induced GFP⁺-LUHMES cell model for high throughput compound screening by assessing a range of mitochondrial and neuronal morphological parameters. We also provide detailed instructions for data and statistical analysis, including example calculations of Z’-score to assess statistical effect size across independent experiments.

CRISPR-Cas9 technology has transformed the ability to edit genomic sequences and control gene expression with unprecedented ease and scale. However, precise genomic insertions of coding sequences using this technology remain time-consuming and inefficient because they require introducing adjacent single-strand cuts through Cas9 nickase action and invoking the host-encoded homology-directed repair program through the concomitant introduction of large repair templates. Here, we present a system for the rapid study of any protein-of-interest in two neuronal cell models following its inducible expression from the human AAVS1 safe harbor locus. With lox-flanked foundation cassettes in the AAVS1 site and a tailor-made plasmid for accepting coding sequences-of-interest in place, the system allows investigators to produce their own neuronal cell models for the inducible expression of any coding sequence in less than a month. Due to the availability of preinserted enhanced green fluorescent protein (EGFP) coding sequences that can be fused to the protein-of-interest, the system facilitates functional investigations that track a protein-of-interest by live-cell microscopy as well as interactome analyses that capitalize on the availability of exquisitely efficient EGFP capture matrices.

Precise and reproducible isolation of desired cell types or layers from heterogeneous tissues is crucial to analyze specific gene profiles and molecular interactions in vivo. Forebrain is the core site of higher functions, like cognition and memory consolidation. It is composed of heterogeneous and distinct cell types, interconnected to form functional neural circuits. Any alteration in the development or function often leads to brain disorders with profound consequences. Thus, precise molecular understanding of forebrain development in normal and diseased scenarios is important. For quantitative studies, most traditional analytical methods require pooling of large cell populations, that results in loss of in vivo tissue integrity and of spatial, molecular and cellular resolution. Laser capture microdissection (LCM) is a fast and extremely precise method of obtaining uncontaminated, homogeneous sets of specific cell types and layers. Our current procedure involves cryo-sectioning and laser microdissection of fresh-frozen mouse forebrains, that are genetically modified and treated with small-molecule therapeutics. Using LCM, specific regions of interest, such as neural layers, cells from adjacent yet distinct subregions within a tissue layer, are obtained under RNase-free conditions. These small cellular cohorts are further used for downstream, high-throughput genomic or transcriptomic assays. Here, we have introduced break-points at multiple stages throughout our protocol. This makes our method simpler and more user-friendly to follow, without compromising on the quality. The current protocol can easily be adapted for different brain regions, as well as for other model organisms/human tissue.

We developed a protocol for photoconversion of endocytic marker FM1-43 followed by electron microscopy analysis of synaptic boutons at the Drosophila neuromuscular junction. This protocol allows detection of stained synaptic vesicle even when release rates are very low, such as during the spontaneous release mode. The preparations are loaded with the FM1-43 dye, pre-fixed, treated and illuminated to photoconvert the dye, and then processed for conventional electron microscopy. This procedure enables clear identification of stained synaptic vesicles at electron micrographs.

In order to explore the role of a specific gene/protein in the specific segment of the spinal cord, the technique of intraspinal injection is particularly used to deliver viral vectors targeting the specific gene/protein. These viral vectors can knockdown or overexpress the specific gene/protein in specific cells (glial cells or neurons). In this protocol, lentivirus containing shRNA for CXCL13 were injected into the dorsal horn of the spinal lumbar enlargement segment (Jiang et al., 2016). This technique allows the study of the role of CXCL13 in the ipsilateral dorsal horn in neuropathic pain without affecting DRG or contralateral dorsal horn.

This protocol is useful to manipulate gene expression in the embryonic retina and compare the result with the contralateral non electroporated retina. In addition, the electroporation of a membrane or cytoplasmic tagged GFP allows to determine the effects of gene manipulation on the outgrowth of retinal ganglion cell axons (Garcia-Frigola et al., 2007) or simply to follow axon outgrowth in mutant embryos. DNA can be directed to different quadrants of the retina (ventral or dorsally) by modifying the position of the electrodes (Petros et al., 2009; Sánchez-Arrones et al., 2013). After the procedure, embryos are left developing to the desired stage, including postnatal stages.

Successful neural circuit formation relies on the accurate navigation of axons towards their targets during development. Axons are guided by a combination of short-range and long-range, attractive and repulsive cues. The commissural axons of the developing spinal cord have provided an informative in vivo model for the identification of multiple axon guidance molecules and mechanisms. These axons extend ventrally from the dorsal spinal cord and cross the midline at the floor plate, before making a sharp rostral turn towards the head. This simple trajectory has facilitated the identification of many axon guidance molecules, because perturbation of the stereotypical guidance decisions as a result of genetic manipulations can be easily identified. The open-book assay is a method to assess the trajectory of spinal commissural axons. The spinal cord is dissected out, opened at the roof plate and pinned flat. Punctate injections of the lipophilic fluorescent dye, DiI, are used to trace commissural axon trajectories prior to microscopy and analysis.

Prion diseases are transmissible, fatal, neurodegenerative diseases in human and animals. The molecular basis of neurodegeneration in prion diseases is largely unclear. Developing a cellular model capable of monitoring prion-induced cytotoxicity would be a promising approach for better understanding the prion pathogenesis. One candidate cellular assay is a model based on neurospheres, which contains neural stem cells (NSCs). Both undifferentiated and differentiated NSCs have been demonstrated to be permissive to prion infection, and prion-induced cytopathic changes in differentiated neruosphere cultures were reported (Iwamaru et al., 2013). This protocol describes the procedure to induce differentiation of NSCs from transgenic mice overexpressing prion protein (tga20 mice) into cultures susceptible for prion infection.

Neurosphere contains neural stem cells that are capable of self-renewal and multilineage differentiation including neurons, astrocytes, and oligodendrocytes (Gage, 2000). Cell culture model using differentiated neurosphere cultures are suggested to be a valuable tool for studying the pathogenesis of prion disease at the cellular level (Iwamaru et al., 2013). This protocol describes the procedure for a culture of whole brain-derived neurospheres from newborn mouse brains. Neurosphere formation steadily occurs within a week from the cultures of neonatal whole brains and these cells have stem cell properties.