Neuroscience

In the peripheral nervous system, Schwann cells are the primary type of glia; their in vitro differentiation and dedifferentiation system has not been described in detail in the literature. Thus, an in vitro differentiation and dedifferentiation system of rat Schwann cells is described in this protocol. These cultures and systems may be used to investigate the morphological and biochemical effects of drug interventions or lentivirus-mediated gene transfer on Schwann cells during differentiation or dedifferentiation.

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The zebrafish retina is a canonical vertebrate retina. Since the past few years, with the continually growing genetic toolbox and imaging techniques, zebrafish plays a crucial role in retinal research. This protocol describes a method to quantitatively evaluate the expression of Arrestin3a (Arr3a) and G-protein receptor kinase7a (Grk7a) in the adult zebrafish retina at protein levels by infrared fluorescence western blot. Our protocol can be easily adapted to measure protein levels in additional zebrafish tissues.

The functions of sleep remain largely unclear, and even less is known about its role in development. A general strategy to tackle these questions is to disrupt sleep and measure the outcomes. However, some existing sleep deprivation methods may not be suitable for studying the effects of chronic sleep disruption, due to their lack of effectiveness and/or robustness, substantial stress caused by the deprivation method, or consuming a large quantity of time and manpower. More problems may be encountered when applying these existing protocols to young, developing animals, because of their likely heightened vulnerability to stressors, and difficulties in precisely monitoring sleep at young ages. Here, we report a protocol of automated sleep disruption in mice using a commercially available, shaking platform–based deprivation system. We show that this protocol effectively and robustly deprives both non-rapid-eye-movement (NREM) sleep and rapid-eye-movement (REM) sleep without causing a significant stress response, and does not require human supervision. This protocol uses adolescent mice, but the method also works with adult mice.

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Automated sleep deprivation system. The platform of the deprivation chamber was programmed to shake in a given frequency and intensity to keep the animal awake while its brain and muscle activities were continuously monitored by electroencephalography and electromyography.

Anorexia nervosa (AN) is a devastating neuropsychiatric disease with a prevalence rate of approximately 0.3%–1% among women and morbidity and mortality rates among the highest of all neuropsychiatric disorders. The disease etiology is complex but primarily characterized by reduced food intake and body weight, and intense anxiety and fear associated with gaining weight. Existing rodent models of AN are useful and capture features of the disease, but either require specialized genetic mouse models or are difficult to implement in mice. Here, we describe two simple mouse models of stress-induced anorexia that are easy to implement in basic research labs, and capture core features associated with AN, such as reduced food intake in the context of social/physical stress and increased anxiety-related behavior. These protocols provide reproducible and robust assays for stress-induced anorexia and may be implemented with additional assays to probe the neural circuitry mediating the effects of psychological stress on feeding in mice.

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Targeted protein degradation (TPD) facilitates the selective elimination of unwanted and pathological cellular cargoes via the proteasome or the lysosome, ranging from proteins to organelles and pathogens, both within and outside the cell. Currently, there are several in vitro and in vivo protocols that assess the degradative potency of a given degrader towards a myriad of targets, most notably soluble, monomeric oncoproteins. However, there is a clear deficiency of methodologies to assess the degradative potency of heterobifunctional chimeric degraders, especially those in the autophagy space, against pathological, mutant tau species, such as detergent-insoluble oligomers and high-molecular aggregates. The protocol below describes both in vitro and in vivo biochemical assays to induce tau aggregation, as well as to qualitatively and quantitatively measure the degradative potency of a given degrader towards said aggregates, with specific applications of the AUTOTAC (AUTOphagy-TArgeting Chimera) platform provided as an example. A well-defined set of methodologies to assess TPD-mediated degradation of pathological tau species will help expand the scope of the TPD technology to neurodegeneration and other proteinopathies, in both the lab and the clinic.

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Overview of assays observing elimination of tauP301L aggregates with AUTOTAC. (A) Description of the biological working mechanism of heterobifunctional chimeric AUTOTAC degraders. (B) Schematic illustration of assays described in this paper.

Lysosomes play a central role in signaling, nutrient sensing, response to stress, and the degradation and recycling of cellular content. Defects in lysosomal digestive enzymes or structural components can impair lysosomal function with dire consequences to the cell, such as neurodegeneration. A number of methods exist to assess lysosomal stress in the model Drosophila, such as specific driver and reporter strains, transmission electron microscopy, and the investigation of gene expression. These methods, however, can be time consuming and, in some cases, costly. The procedure described here provides a quick, reliable, and low-cost approach to measure lysosomal stress in the Drosophila brain. Using fluorescence confocal microscopy and the LysoTracker staining, this protocol allows for the direct measurement of lysosome size and number. This method can be used to assess lysosomal stress under a number of different genetic and environmental scenarios in the Drosophila brain.

Molecular characterization of different cell types in rodent brains is a widely used and important approach in neuroscience. Fluorescent detection of transcripts using RNAscope (ACDBio) has quickly became a standard in situ hybridization (ISH) approach. Its sensitivity and specificity allow for the simultaneous detection of between three and forty-eight low abundance mRNAs in single cells (i.e., multiplexing or hiplexing), and, in contrast to other ISH techniques, it is performed in a shorter amount of time. Manual quantification of transcripts is a laborious and time-consuming task even for small portions of a larger tissue section. Herein, we present a protocol for creating high-quality images for quantification of RNAscope-labeled neurons in the rat brain. This protocol uses custom-made scripts within the open-source software QuPath to create an automated workflow for the careful optimization and validation of cell detection parameters. Moreover, we describe a method to derive mRNA signal thresholds using negative controls. This protocol and automated workflow may help scientists to reliably and reproducibly prepare and analyze rodent brain tissue for cell type characterization using RNAscope.

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In nature, parasitoid wasp infections are a major cause of insect mortality. Parasitoid wasps attack a vast range of insect species to lay their eggs. As a defense, insects evolved survival strategies to protect themselves from parasitoid infection. While a growing number of studies reported both host defensive tactics and parasitoid counter-offensives, we emphasize that this parasite–host relationship presents a unique ecological and evolutionary relevant model that is often challenging to replicate in a laboratory. Although maintaining parasitoid wasp cultures in the laboratory requires meticulous planning and can be labor intensive, a diverse set of wasp species that target many different insect types can be maintained in similar culture conditions. Here, we describe the protocol for culturing parasitoid wasp species on Drosophila larvae and pupae in laboratory conditions. We also detail an egg-laying assay to assess the reproductive modification of Drosophila females in response to parasitoid wasps. This behavioral study is relatively simple and easily adaptable to study environmental or genetic influences on egg-laying, a readout for female germline development. Neither the parasitoid culture conditions or the behavioral assay require special supplies or equipment, making them a powerful and versatile approach in research or teaching laboratory settings.

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Pathogen invasion of the central nervous system (CNS) is an important cause of infection-related mortality worldwide and can lead to severe neurological sequelae. To gain access to the CNS cells, pathogens have to overcome the blood–brain barrier (BBB), a protective fence from blood-borne factors. To study host–pathogen interactions, a number of cell culture and animal models were developed. However, in vitro models do not recapitulate the 3D architecture of the BBB and CNS tissue, and in vivo mammalian models present cellular and technical complexities as well as ethical issues, rendering systematic and genetic approaches difficult. Here, we present a two-pronged methodology allowing and validating the use of Drosophila larvae as a model system to decipher the mechanisms of infection in a developing CNS. First, an ex vivo protocol based on whole CNS explants serves as a fast and versatile screening platform, permitting the investigation of molecular and cellular mechanisms contributing to the crossing of the BBB and consequences of infection on the CNS. Then, an in vivo CNS infection protocol through direct pathogen microinjection into the fly circulatory system evaluates the impact of systemic parameters, including the contribution of circulating immune cells to CNS infection, and assesses infection pathogenicity at the whole host level. These combined complementary approaches identify mechanisms of BBB crossing and responses of a diversity of CNS cells contributing to infection, as well as novel virulence factors of the pathogen.

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Procedures flowchart.
Mammalian neurotropic pathogens could be tested in two Drosophila central nervous system (CNS) infection setups (ex vivo and in vivo) for their ability to: (1) invade the CNS (pathogen quantifications), (2) disturb blood–brain barrier permeability, (3) affect CNS host cell behaviour (gene expression), and (4) alter host viability.