Neuroscience

Research into nervous system injuries and regeneration has emerged as a crucial field of study. In many cases such as trauma or stroke, both axons and dendrites are equally damaged; however, studying injury and repair mechanisms in both neurite processes (axons and dendrites) of the same neuron has been challenging. Additionally, correlating the behavioral aspects of neuronal injury with anatomical regeneration is important for a better understanding of the functional rewiring process. Here, we describe protocols for injuring the dendrites and the axon of the PVD neuron of C. elegans using a two-photon infrared (IR) femtosecond laser system, and subsequent imaging of injured neurites during the course of regeneration. Additionally, we describe the protocols for the behavioral study concerning the PVD neuron and their analysis, which can offer valuable insights. These assays can be implemented to assess the function of the pathways that play specific roles in dendrite vs. axon regeneration.

Fluorescence lifetime imaging microscopy (FLIM) is a highly valuable technique in the fluorescence microscopy toolbox because it is essentially independent of indicator concentrations. Conventional fluorescence microscopy analyzes changes in emission intensity. In contrast, FLIM assesses the fluorescence lifetime, which is defined as the time a fluorophore remains in an excited state before emitting a photon. This principle is advantageous in experiments where fluorophore concentrations are expected to change, e.g., due to changes in cell volume. FLIM, however, requires collecting a substantial number of photons to accurately fit distribution plots, which constrains its ability for dynamic imaging. This limitation has recently been overcome by rapidFLIM, which utilizes ultra-low dead-time photodetectors in conjunction with sophisticated rapid electronics. The resulting reduction in dead-time to the picosecond range greatly enhances the potential for achieving high spatio-temporal resolution. Here, we demonstrate the use of multi-photon-based rapidFLIM with the sodium indicator ION NaTRIUM Green-2 (ING-2) for the quantitative, dynamic determination of Na⁺ concentrations in neurons in acute rodent brain tissue slices. We describe the loading of the dye into neurons and present a procedure for its calibration in situ. We show that rapidFLIM not only allows the unbiased determination of baseline Na⁺ concentrations but also allows dynamic imaging of changes in intracellular Na⁺, e.g., induced by inhibition of cellular ATP production. Overall, rapidFLIM, with its greatly improved signal-to-noise ratio and higher spatio-temporal resolution, will also facilitate dynamic measurements using other FLIM probes, particularly those with a low quantum yield.

Calcium-permeable AMPA receptors (CP-AMPARs) and kainate receptors (CP-KARs) play crucial roles in synaptic plasticity and are implicated in various neurological processes. Current methods for identifying neurons expressing these receptors, such as electrophysiological recordings and immunostaining, have limitations in throughput or inability to distinguish functional receptors. This protocol describes a novel approach for the vital identification of neurons containing CP-AMPARs and CP-KARs using calcium imaging. The method involves loading neurons with Fura-2 AM, a calcium-sensitive fluorescent probe, KCl application to identify all neurons, and further addition of specific AMPAR agonists (e.g., 5-fluorowillardiine) in the presence of voltage-gated calcium channel blockers and NMDAR/KAR antagonists to identify CP-AMPAR-containing neurons. CP-KAR-containing neurons are identified using domoic acid applications in the presence and absence of NASPM (a CP-AMPAR antagonist). This technique offers several advantages over existing methods, including the ability to assess large neuronal populations simultaneously, distinguish between different receptor types, and provide functional information about CP-AMPAR and CP-KAR expression in living neurons, making it a valuable tool for studying synaptic plasticity and neurological disorders.

The neuromuscular junction (NMJ) is an interface between motor neurons and skeletal muscle fibers, facilitating the transmission of signals that initiate muscle contraction. Its pivotal role lies in ensuring efficient communication between the nervous system and muscles, allowing for precise and coordinated movements essential for everyday activities and overall motor function. To provide insights into neuromuscular disease and development, understanding the physiology of NMJ is essential. We target acetylcholine receptors (AChR) by immunofluorescence assay (IFA) with α-bungarotoxin (BTX; snake venom neurotoxins binding to AChR) to visualize and quantify the NMJ. Changes in AChR distribution or structure can indicate alterations in receptor density, which may be associated with neuromuscular disorders or conditions that affect synaptic transmission. This protocol provides the methodology for isolating and longitudinally sectioning gastrocnemius muscle for AChR-targeted IFA for confocal microscopy and performing quantitative analysis of NMJs.

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

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

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

The pancreas is a heavily innervated organ, but pancreatic innervation can be challenging to comprehensively assess using conventional histological methods. However, recent advances in whole-mount tissue clearing and 3D rendering techniques have allowed detailed reconstructions of pancreatic innervation. Optical clearing is used to enhance tissue transparency and reduce light scattering, thus eliminating the need to section the tissue. Here, we describe a modified version of the optical tissue clearing protocol iDISCO+ (immunolabeling-enabled three-dimensional imaging of solvent-cleared organs) optimized for pancreatic innervation and endocrine markers. The protocol takes 13-19 days, depending on tissue size. In addition, we include protocols for imaging using light sheet and confocal microscopes and for 3D segmentation of pancreatic innervation and endocrine cells using Imaris.

Recently developed CLARITY (Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging/Immunostaining/In situ-hybridization-compatible Tis-sue-hYdrogel) technique renders the tissue transparent by removing lipids in the tissue, while preserving and stabilizing the cellular and subcellular structures. This provides effective penetration of diverse labeling probes, from primary and secondary antibodies to complementary DNA and RNA strands. Followed by high-resolution 3D imaging of neuronal cells and their projections in thick sections, tissue blocks, whole brains, or whole animals, CLARITY allows for superior quantitative analysis of neuronal tissue. Here, we provide our detailed protocol for PACT (Passive Clarity Technique) in brain tissue of diverse species, including human, non-human primate, rodents, and zebrafish. We describe the six principal steps: (1) Tissue fixation and preparation, (2) Passive lipid removal, (3) Immuno-labeling, (4) Optical clearing, (5) Imaging, (6) 3D visualization and quantification.