Neuroscience

The process of moving proteins and organelles along the axon is essential for neuronal survival and function, ensuring proper communication between the cell body and distant synapses. The efficient and precise delivery of proteins via axon transport is critical for processes ranging from synaptic plasticity and neurotransmission to neuronal growth and maintenance. However, the identities of all the transported proteins have only recently begun to be investigated. Retinal ganglion cells (RGCs) provide a unique opportunity for access to central nervous system (CNS) axons as the retina is located outside the brain in the eye, with long axonal projections (~1 cm in mouse) that innervate the brain. We have developed and optimized methods for unbiased in vivo protein labeling in rodent RGC somata with intravitreal N-hydroxysuccinimido (NHS)-biotin and subsequent visualization of transported proteins along the optic nerve using confocal microscopy. Here, we describe these procedures in detail.

An improved correlative light and electron microscopy (CLEM) method has recently been introduced and successfully employed to identify and analyze protein inclusions in cultured cells as well as pathological proteinaceous deposits in postmortem human brain tissues from individuals with diverse neurodegenerative diseases. This method significantly enhances antigen preservation and target registration by replacing conventional dehydration and embedding reagents. It achieves an optimal balance of sensitivity, accuracy, efficiency, and cost-effectiveness compared to other current CLEM approaches. However, due to space constraints, only a brief overview of this method was provided in the initial publication. To ensure reproducibility and facilitate widespread adoption, the author now presents a detailed, step-by-step protocol of this optimized CLEM technique. By enhancing usability and accessibility, this protocol aims to promote broader application of CLEM in neurodegenerative disease research.

Proper brain function depends on the integrity of the blood–brain barrier (BBB), which is formed by a specialized network of microvessels in the brain. Reliable isolation of these microvessels is crucial for studying BBB composition and function in both health and disease. Here, we describe a protocol for the mechanical dissociation and density-based separation of microvessels from fresh or frozen human and murine brain tissue. The isolated microvessels retain their molecular integrity and are compatible with downstream applications, including fluorescence imaging and biochemical analyses. This method enables direct comparisons across species and disease states using the same workflow, facilitating translational research on BBB biology.

Brain endothelial cells, which constitute the cerebrovasculature, form the first interface between the blood and brain and play essential roles in maintaining central nervous system (CNS) homeostasis. These cells exhibit strong apicobasal polarity, with distinct luminal and abluminal membrane compositions that crucially mediate compartmentalized functions of the vasculature. Existing transcriptomic and proteomic profiling techniques often lack the spatial resolution to discriminate between these membrane compartments, limiting insights into their distinct molecular compositions and functions. To overcome these limitations, we developed an in vivo proteomic strategy to selectively label and enrich luminal cerebrovascular proteins. In this approach, we perfuse a membrane-impermeable biotinylation reagent into the vasculature to covalently tag cell surface proteins exposed on the luminal side. This is followed by microvessel isolation and streptavidin-based enrichment of biotinylated proteins for downstream mass spectrometry analysis. Using this method, we robustly identified over 1,000 luminally localized proteins via standard liquid chromatography–tandem mass spectrometry (LC–MS/MS) techniques, achieving substantially improved enrichment of canonical luminal markers compared with conventional vascular proteomic approaches. Our method enables the generation of a high-confidence, compartment-resolved atlas of the luminal cerebrovascular proteome and offers a scalable platform for investigating endothelial surface biology in both healthy and disease contexts.

The global burden of stroke has increased in the past several decades, and post-stroke epilepsy (PSE) is a common complication. Contrasted with the advancement in knowledge of stroke pathophysiology, the exact pathogenesis of PSE is unclear. Various animal stroke models have been utilized to investigate the underlying mechanisms of PSE, but the success rate of PSE induction is low. To address this limitation, a novel PSE model was established in the rat by inducing status epilepticus using lithium-pilocarpine one week after photothrombotic stroke. Successful indication of status epilepticus and mortality rate at three days after status epilepticus were the main measurements. Potential usefulness of this model was also illustrated by preliminary results on locomotor activity, exploratory behavior, and anxiety level evaluated using the open-field test, as well as mossy fiber sprouting (MFS) in the hippocampal dentate granule cells using Zinc transporter 3 immunofluorescence staining at 8 weeks after PSE induction. This novel composite method of PSE induction may facilitate future studies on the pathogenesis and treatment of PSE.

We recently developed an approach for cell type–specific CRISPR/Cas9 editing and transgene expression using a single viral vector. Here, we present a protocol describing how to design and generate plasmids and adeno-associated viruses (AAVs) compatible with this single-vector gene editing approach. This protocol has four components: (1) guide RNA (gRNA) design to target specific genes of interest, (2) ligation and cloning of CRISPR-competent AAV vectors, (3) production of vector-containing AAVs, and (4) viral titer quantification. The resultant vectors are compatible for use with mouse lines expressing the Cas9 protein from Streptococcus pyogenes (SpCas9) and Cre recombinase to enable selective co-expression of standard neuroscience tools in edited cells. This protocol can produce AAVs of any serotype, and the resulting AAVs can be used in the central and peripheral nervous systems. This flexible approach could help identify and test the function of novel genes affecting synaptic transmission, circuit activity, or morphology with a single viral injection.

The fatal motor neuron (MN) disease amyotrophic lateral sclerosis (ALS) is characterized by progressive degeneration of the phrenic MNs (phMNs) controlling the activity of the diaphragm, leading to death by respiratory failure. Human experimental models to study phMNs are lacking, hindering the understanding of the mechanisms of phMN degeneration in ALS. Here, we describe a protocol to derive phrenic-like MNs from human induced pluripotent stem cells (hiPSC-phMNs) within 30 days. During spinal cord development, phMNs emerge from specific MN progenitors located in the dorsalmost MN progenitor (pMN) domain at cervical levels, under the control of a ventral-to-dorsal gradient of Sonic hedgehog (SHH) signaling and a rostro-caudal gradient of retinoic acid (RA). The method presented here uses optimized concentrations of RA and the SHH agonist purmorphamine, followed by fluorescence-activated cell sorting (FACS) of the resulting MN progenitor cells (MNPCs) based on a cell-surface protein (IGDCC3) enriched in hiPSC-phMNs. The resulting cultures are highly enriched in MNs expressing typical phMN markers. This protocol enables the generation of hiPSC-phMNs and is highly reproducible using several hiPSC lines, offering a disease-relevant system to study mechanisms of respiratory MN dysfunction. While the protocol has been validated in the context of ALS research, it can be adopted to study human phrenic MNs in other research fields where these neurons are of interest.

Over the lifespan of an individual, brain function requires adjustments in response to environmental changes and learning experiences. During early development, neurons overproduce neurite branches, and neuronal pruning removes the unnecessary neurite branches to make a more accurate neural circuit. Drosophila motoneurons prune their intermediate axon bundles rather than the terminal neuromuscular junction (NMJ) by degeneration, which provides a unique advantage for studying axon pruning. The pruning process of motor axon bundles can be directly analyzed by real-time imaging, and this protocol provides a straightforward method for monitoring the developmental process of Drosophila motor neurons using live cell imaging.

Since the discovery that astrocytes are characterized by Ca²⁺-based excitability, investigating the function of these glial cells within the brain requires Ca²⁺ imaging approaches. The technical evolution from chemical fluorescent Ca²⁺ probes with low cellular specificity to genetically encoded indicators (GECIs) has enabled detailed analysis of the spatial and temporal features of intracellular Ca²⁺ signal. Different imaging methodologies allow the extraction of distinct information on calcium signals in astrocytes from brain slices, with resolution ranging from cell populations to single cells up to subcellular domains.

Here, we describe 2-photon laser scanning microscopy (2PLSM) Ca²⁺ imaging in astrocytes from the somatosensory cortex (SSCx) of adult mice in ex vivo acute cortical slices, performed using two genetically encoded Ca²⁺ indicators, i.e., cytosolic GCaMP6f and endoplasmic reticulum-targeted G-CEPIA1er. The main advantage of the 2PLSM technique, compared to single-photon microscopy, is the possibility to go deeper in the tissue while avoiding photodamage, by limiting laser excitation to a single focal plane. The fluorescent signal of the indicator is analyzed offline in different compartments—soma, proximal processes, and microdomains—for GCaMP6f experiments and in the perinuclear, somatic area for G-CEPIA1er. The analysis of Ca²⁺ signal from different compartments, although not providing a value of absolute concentration, allows a critical comparison of the degree of astrocyte activation between different experimental conditions or mouse models. Moreover, the analysis of G-CEPIA1er signal, which reveals metabotropic receptor activation as a dynamic decrease in free Ca²⁺ in the endoplasmic reticulum (ER), can provide information on possible alterations in this critical second messenger pathway in astrocytes, including, for example, steady-state ER Ca²⁺ levels and kinetics of Ca²⁺ release.

In vivo two-photon imaging of the mouse brain is essential for understanding brain function in relation to neural structure; however, its application is limited by the size and mechanical stability of conventional cranial windows. Here, we present the procedure of a large-scale cranial window technique based on the nanosheet incorporated into light-curable resin (NIRE) method. This approach utilizes a biocompatible polyethylene-oxide-coated CYTOP (PEO-CYTOP) nanosheet combined with light-curable resin, allowing the window to conform to the brain’s curved surface. The protocol enables long-term, high-resolution, and multiscale imaging—from subcellular structures to large neuronal populations—in awake mice over several months.