The neuromuscular junction (NMJ) is a specialized synapse that connects the terminal end of a motor neuron and a skeletal muscle fiber. Defects in NMJ cause abnormalities of neuromuscular transmission, leading to NMJ disorders. The mammalian diaphragm muscle is essential for respiration and has been widely used to study NMJ formation. Here, we provide a simple and straightforward protocol for preparing diaphragms from embryonic, neonatal, and adult mice and for subsequent NMJ staining.

This protocol aims to visualize the filamentous actin network in Chlamydomonas reinhardtii. We improved fixed-cell labeling conditions using the F-actin probe, phalloidin. We created a Chlamydomonas-optimized protocol by halving the phalloidin incubation time, electing for optimal fixation conditions, and selecting for a healthy cell population. This phalloidin protocol is quick, effective, and is the only labeling method to date that allows for reliable actin filament detection in fixed vegetative Chlamydomonas cells. This method reveals previously unidentified actin structures in Chlamydomonas and novel insights into cytoskeletal dynamics.

Quantitative analysis of proteins secreted from the cells poses a challenge due to their low abundance and the interfering presence of a large amount of bovine serum albumin (BSA) in the cell culture media. We established assays for detection of mutant huntingtin (mHtt) secreted from Neuro2A cell line stably expressing mHtt and rat primary cortical neurons by Western blotting. Our protocol is based on reducing the amounts of BSA in the media while maintaining cell viability and secretory potential, and concentrating the media prior to analysis by means of ultrafiltration.

According to the World Health Organization (WHO), at least 50% of emerging viruses endowed with pathogenicity in humans can infect the Central Nervous System (CNS) with induction of encephalitis and other neurologic diseases (Taylor et al., 2001; Olival and Daszak, 2005). While neurological diseases are progressively documented, the underlying cellular and molecular mechanisms involved in virus infection and dissemination within the CNS are still poorly understood (Swanson and McGavern, 2015; Ludlow et al., 2016). For example, measles virus (MeV) can infect neural cells, and cause a persistent brain infections leading to lethal encephalitis from several months to years after primary infection with no available treatment (Reuter and Schneider-Schaulies, 2010; Laksono et al., 2016). The Organotypic Brain Culture (OBC) is a suitable model for the virology field to better understand the CNS infections. Indeed, it allows not only studying the infection and the dissemination of neurotropic viruses within the CNS but it could also serve as screening model of innovative antiviral strategies or molecules, such as our recently published studies about fusion inhibitory peptides and the HSP90 chaperone activity inhibitor, 17-DMAG (Welsch et al., 2013; Bloyet et al., 2016). Based on our previous work, we propose here an optimized method to prepare OBC of hippocampi and cerebellums which are suitable for small rodent models based virus studies, including mice, rats as well as hamsters at a post-natal stage, between P6 to P10. We notably took into account the stress of the slice procedure on the tissue and the subsequent cellular reactions, which is essential to fully characterize the model prior to any use in infectious conditions. With this knowledge, we propose a protocol highlighting the requirements, including potential trouble shootings of the slicing parameters, to consider the variations we observed according to the structure and animal studied. This framework should facilitate the use of OBC for better conclusive studies of neurotropic viruses.

This protocol aims at standardizing the procedure to obtain primary cultures of hippocampal and cerebrocortical neurons for in vitro experiments. Cultures should be prepared from cells isolated during embryonic development when neuronal precursor cells are not yet fully differentiated. This helps increasing the quality and quantity of cells, while offering minimal cell death that often occurs during dissociation of differentiated neurons. Cells plated under the appropriate conditions, either in Petri-dishes or in multi-well plates, will develop and establish synaptic contacts over time since the neuronal culture medium provides the nutrients and trophic factors required for differentiation. In this protocol we describe the methodology for the preparation of both cortical and hippocampal neuronal cultures.

Action potential conduction velocity is the speed at which an action potential (AP) propagates along an axon. Measuring AP conduction velocity is instrumental in determining neuron health, function, and computational capability, as well as in determining short-term dynamics of neuronal communication and AP initiation (Ballo and Bucher, 2009; Bullock, 1951; Meeks and Mennerick, 2007; Rosenthal and Bezanilla, 2000; Städele and Stein, 2016; Swadlow and Waxman, 1976). Conduction velocity can be measured using extracellular recordings along the nerve through which the axon projects. Depending on the number of axons in the nerve, AP velocities of individual or many axons can be detected.

This protocol outlines how to measure AP conduction velocity of (A) stimulated APs and (B) spontaneously generated APs by using two spatially distant extracellular electrodes. Although an invertebrate nervous system is used here, the principles of this technique are universal and can be easily adjusted to other nervous system preparations (including vertebrates).

Pathological conditions such as amyotrophic lateral sclerosis, spinal cord injury and chronic pain are characterized by activation of astrocytes and microglia in spinal cord and have been modeled in rodents. In vivo imaging at cellular level in these animal models is limited due to the spinal cord’s highly myelinated funiculi. The preparation of acute slices may offer an alternative and valuable strategy to collect structural and functional information in vitro from dorsal, lateral and ventral regions of spinal cord. Here, we describe a procedure for preparing acute slices from mouse spinal cord (Garré et al., 2016). This preparation should allow further understanding of how glial cells in spinal cord respond acutely to various inflammatory challenges.

In these culture models, the normal cytoarchitecture and local neuronal circuits of the spinal cord are preserved, offering a compromise between dissociated cell cultures and complete animal studies. The addition of 5-ethynyl-2’-deoxyuridine (EdU) to the culture medium allows for the detection of proliferating cells.

This protocol details beta-amyloid (Aβ) extraction from transgenic murine brain homogenates. Specifically, mechanical homogenization of brain tissue and sequential extraction of both soluble and insoluble proteins are detailed. DEA extracts soluble proteins, such as Aβ isoforms and APP. Formic acid enables extraction of insoluble protein aggregates, such as Aβ isoforms associated with plaques. This procedure produces soluble and insoluble extracts that are amenable to analysis of Aβ species via western blotting and/or enzyme-linked immunosorbent assays (ELISAs), and these results help assess amyloidogenic burden in animals.

Inositol triphosphate (IP₃) is an important second messenger that participates in signal transduction pathways in diverse cell types including hippocampal neurons. Stimulation of phospholipase C in response to various stimuli (hormones, growth factors, neurotransmitters, neurotrophins, neuromodulators, odorants, light, etc.) results in hydrolysis of phosphatidylinositol 4, 5-bisphosphate (PIP₂), a phospholipid that is located in the plasma membrane, and leads to the production of IP₃ and diacylglycerol. Binding of IP₃ to the IP₃ receptor (IP₃R) induces Ca²⁺ release from intracellular stores and enables the initiation of intracellular Ca²⁺-dependent signaling. Here we describe a procedure for the measurement of cellular IP₃ levels in tissue homogenates prepared from rat hippocampal slices.