Neuroscience

Recent advancements in tissue-clearing techniques and volumetric imaging have greatly facilitated visualization and quantification of biomolecules, organelles, and cells in intact organs or even entire organisms. Generally, there are two types of clearing methods: hydrophobic and hydrophilic (i.e., clearing with organic or aqueous solvents, respectively). The popular iDISCO approach and its modifications are hydrophobic methods that involve dehydration, delipidation, decolorization (optional), decalcification (optional), and refractive-index (RI) matching steps. Cleared samples are often stored for a relatively long period of time and imaged repeatedly. However, cleared tissues can become opaque over time, which prevents accurate reimaging. We reasoned that the resurgent haziness is likely due to rehydration, residual lipids, and uneven RI deep inside those tissue samples. For rescue, we have developed a simple procedure based on iDISCO. Beginning with a methanol dehydration, samples are delipidated using dichloromethane, followed by RI matching with dibenzyl ether (DBE). This simple method effectively re-clears mouse brains that have turned opaque during months of storage, allowing the user to effectively image immunolabeled samples over longer periods of time.

Key features

• This simple protocol rescues previously cleared tissue that has turned opaque.

• The method does not cause detectable loss of immunofluorescence from previously stained samples.

Graphical overview

Structural and functional changes in vascular networks play a vital role during development, causing or contributing to the pathophysiology of injury and disease. Current methods to trace and image the vasculature in laboratory settings have proven inconsistent, inaccurate, and labor intensive, lacking the inherent three-dimensional structure of vasculature. Here, we provide a robust and highly reproducible method to image and quantify changes in vascular networks down to the capillary level. The method combines vasculature tracing, tissue clearing, and three-dimensional imaging techniques with vessel segmentation using AI-based convolutional reconstruction to rapidly process large, unsectioned tissue specimens throughout the body with high fidelity. The practicality and scalability of our protocol offer application across various fields of biomedical sciences. Obviating the need for sectioning of samples, this method will expedite qualitative and quantitative analyses of vascular networks. Preparation of the fluorescent gel perfusate takes < 30 min per study. Transcardiac perfusion and vasculature tracing takes approximately 20 min, while dissection of tissue samples ranges from 5 to 15 min depending on the tissue of interest. The tissue clearing protocol takes approximately 24–48 h per whole-tissue sample. Lastly, three-dimensional imaging and analysis can be completed in one day. The entire procedure can be carried out by a competent graduate student or experienced technician.

Key features

• This robust and highly reproducible method allows users to image and quantify changes in vascular networks down to the capillary level.

• Three-dimensional imaging techniques with vessel segmentation enable rapid processing of large, unsectioned tissue specimens throughout the body.

• It takes approximately 2–3 days for sample preparation, three-dimensional imaging, and analysis.

• The user-friendly pipeline can be completed by experienced and non-experienced users.

Graphical overview

Capillary density in skeletal muscles is key to estimate exercise capacity in healthy individuals, athletes, and those with muscle-related pathologies. Here, we present a step-by-step, high-throughput semi-automated method for quantifying capillary density from whole human skeletal muscle cross-sections, in areas of the muscle occupied by myofibers. We provide a detailed protocol for immunofluorescence staining, image acquisition, processing, and quantification. Image processing is performed in ImageJ, and data analysis is conducted in R. The provided protocol allows high-throughput quantification of capillary density.

Key features

• This protocol builds upon the method and results described in Abbassi-Daloii et al. (2023b).

• It includes step-by-step details on image acquisition and image processing of the entire muscle section.

• It enables high-throughput and semi-automated image quantification of capillary density.

• It provides a robust analysis for determining capillary density over the entire muscle cross section.

Graphical overview

Measuring the action potential (AP) propagation velocity in axons is critical for understanding neuronal computation. This protocol describes the measurement of propagation velocity using a combination of somatic whole cell and axonal loose patch recordings in brain slice preparations. The axons of neurons filled with fluorescent dye via somatic whole-cell pipette can be targeted under direct optical control using the fluorophore-filled pipette. The propagation delays between the soma and 5–7 axonal locations can be obtained by analyzing the ensemble averages of 500–600 sweeps of somatic APs aligned at times of maximal rate-of-rise (dV/dtmax) and axonal action currents from these locations. By plotting the propagation delays against the distance, the location of the AP initiation zone becomes evident as the site exhibiting the greatest delay relative to the soma. Performing linear fitting of the delays obtained from sites both proximal and distal from the trigger zone allows the determination of the velocities of AP backward and forward propagation, respectively.

Key features

• Ultra-thin axons in cortical slices are targeted under direct optical control using the SBFI-filled pipette.

• Dual somatic whole cell and axonal loose patch recordings from 5–7 axonal locations.

• Ensemble averaging of 500–600 sweeps of somatic APs and axonal action currents.

• Plotting the propagation delays against the distance enables the determination of the trigger zone's position and velocities of AP backward and forward propagation.

Whole-brain clearing and imaging methods are becoming more common in mice but have yet to become standard in rats, at least partially due to inadequate clearing from most available protocols. Here, we build on recent mouse-tissue clearing and light-sheet imaging methods and develop and adapt them to rats. We first used cleared rat brains to create an open-source, 3D rat atlas at 25 μ resolution. We then registered and imported other existing labeled volumes and made all of the code and data available for the community (https://github.com/emilyjanedennis/PRA) to further enable modern, whole-brain neuroscience in the rat.

Key features

• This protocol adapts iDISCO (Renier et al., 2014) and uDISCO (Pan et al., 2016) tissue-clearing techniques to consistently clear rat brains.

• This protocol also decreases the number of working hours per day to fit in an 8 workday.

Graphical overview

Enhancing axon regeneration is a major focus of peripheral nerve injury research. Although peripheral axons possess a limited ability to regenerate, their functional recovery is very poor. Various activity-based therapies like exercise, optical stimulation, and electrical stimulation as well as pharmacologic treatments can enhance spontaneous axon regeneration. In this protocol, we use a custom-built cuff to electrically stimulate the whole sciatic nerve for an hour prior to transection and repair. We used a Thy-1-YFP-H mouse to visualize regenerating axon profiles. We compared the regeneration of axons from nerves that were electrically stimulated to nerves that were not stimulated (untreated). Electrically stimulated nerves had longer axon growth than the untreated nerves. We detail how variations of this method can be used to measure acute axon growth.

A fascinating question in neuroscience is how sensory stimuli evoke calcium dynamics in neurons. Caenorhabditis elegans is one of the most suitable models for optically recording high-throughput calcium spikes at single-cell resolution. However, calcium imaging in C. elegans is challenging due to the difficulties associated with immobilizing the organism. Currently, methods for immobilizing worms include entrapment in a microfluidic channel, anesthesia, or adhesion to a glass slide. We have developed a new method to immobilize worms by trapping them in sodium alginate gel. The sodium alginate solution (5%), polymerized with divalent ions, effectively immobilizes worms in the gel. This technique is especially useful for imaging neuronal calcium dynamics during olfactory stimulation. The highly porous and transparent nature of alginate gel allows the optical recording of cellular calcium oscillations in neurons when briefly exposed to odor stimulation.

Sleep is a conserved biological process in the animal kingdom. Understanding the neural mechanisms underlying sleep state transitions is a fundamental goal of neurobiology, important for the development of new treatments for insomnia and other sleep-related disorders. Yet, brain circuits controlling this process remain poorly understood. A key technique in sleep research is to monitor in vivo neuronal activity in sleep-related brain regions across different sleep states. These sleep-related regions are usually located deeply in the brain. Here, we describe technical details and protocols for in vivo calcium imaging in the brainstem of sleeping mice. In this system, sleep-related neuronal activity in the ventrolateral medulla (VLM) is measured using simultaneous microendoscopic calcium imaging and electroencephalogram (EEG) recording. By aligning calcium and EEG signals, we demonstrate that VLM glutamatergic neurons display increased activity during the transition from wakefulness to non-rapid eye movement (NREM) sleep. The protocol described here can be applied to study neuronal activity in other deep brain regions involved in REM or NREM sleep.

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.

Graphical abstract

Actin filaments are essential for various biological activities in eukaryotic cellular processes. Available in vitro experimental data on these systems often lack details and information on sample preparation protocols and experimental techniques, leading to unreproducible results. Additionally, different experimental techniques and polymerization buffers provide different, sometimes contradictory results on the properties of these systems, making it substantially difficult to gather meaningful data and conclusive information from them. This article presents a robust, accurate, detailed polymerization protocol to prepare high-quality actin filament samples for light scattering experiments. It has been shown to provide unicity and consistency in preparing stable, dispersed, aggregates-free, homogenous actin filament samples that could benefit many other scientific research groups currently working in the field. To develop the protocol, we used conventional actin buffers in physiological conditions. However, it can easily be adapted to prepare samples using other buffers and biological fluids. This protocol yielded reproducible results on essential actin filament parameters such as the translational diffusion coefficient and electrophoretic mobility. Overall, suitable modifications of the proposed experimental method could generate accurate, reproducible light scattering results on other highly charged anionic filaments commonly found in biological cells (e.g., microtubules, DNAs, RNAs, or filamentous viruses).

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