Neuroscience

Small fiber neuropathy (SFN) is an underdiagnosed condition characterized by sensory and autonomic dysfunction due to impairment of small nerve fibers in skin, blood vessels, and internal organs. Various underlying disorders are associated with SFN, and the pathophysiology of nerve fiber damage and functional impairment is the subject of extensive research. Diagnosis of SFN is challenging as standard electrodiagnostic techniques assess large fiber function and therefore are normal in SFN patients. The current gold standard for SFN diagnosis in humans is a skin biopsy, commonly obtained from the distal leg, hairy skin region, with evaluation of intraepidermal nerve fiber density (IENFD) using protein gene product 9.5 (PGP9.5) immunolabeling. While well-established in clinical practice, equivalent standardized, reproducible methods for assessing IENFD in experimental mouse models are lacking, which limits translational research in this field. Previous work in mice has relied on diverse antibodies, variable tissue sampling, and the use of confocal microscopy to trace nerve fibers. Other approaches have used chromogenic precipitate-based staining, which limits the ability to co-label multiple proteins. Here, we present a detailed, simple, and reproducible protocol for IENFD quantification of small nerves in the distal glabrous skin of the mouse hind paw. This protocol uses the two distal footpads, ensuring consistent sampling across animals. Prior to sectioning, the tissue is fixed and cryoprotected. Serial 20-μm sections are mounted on glass slides, dried, permeabilized, blocked, and immunostained with an anti-PGP9.5 monoclonal antibody, and then detected by binding secondary fluorescent-labeled antibodies. Although murine hairy skin analysis may apparently show a higher translational value, as it better reflects human biopsy sites, it is compromised by dense hair shafts and follicles, which interrupt epidermis continuity and thus interfere with sampling consistency. Polyneuropathy sensory symptoms, in fact, begin at the most distal sensory site, which is the glabrous skin of the toes. Thus, evaluation of this anatomical location best represents the clinical realm and may have the best sensitivity for identifying early axonal changes. In this protocol, we focused on IENFD quantification as done in human samples. Mechanoreceptors such as Meissner corpuscles are detectable and quantifiable by this method, and represent additional value since pressure-evoked pain, transmitted by these, is often reported by affected individuals. This immunolabeling protocol can be completed within one day [involving a small number of animals, where all three stages can be performed during a long working day (approximately 12 h)], while the entire workflow, including fixation and cryoprotection, is completed in up to 72 h. Importantly, the dermal and epidermal small fibers can be visualized using a standard fluorescence microscope, thereby avoiding the need for confocal imaging while maintaining high reproducibility. Preliminary validation in several animal models of inflammatory neuropathy and pain demonstrated a reproducible approximately 50% reduction in IENFD compared to controls, reaching statistical significance with n = 4 per group. This method supports SFN research and preclinical evaluation of novel therapeutics.

Peripheral nerve injuries (PNIs) often result in incomplete functional recovery due to insufficient or misdirected axonal regeneration. Balanced regeneration of myelinated A-fibers and unmyelinated C-fibers is essential for functional recovery, making it crucial to understand their differential regeneration patterns to improve PNI treatment outcomes. However, immunochemical staining does not clearly differentiate between A- and C-fiber axons in whole-mount nerve preparations. To overcome this limitation, we developed a modified protocol by optimizing the immunostaining to restrict the antibody access to myelinated axons. This enables visualization of A-fibers by myelin sheath labeling, while allowing selective staining of unmyelinated C-fiber axons. As a result, A- and C-fibers can be reliably distinguished, facilitating accurate analysis of their regeneration in both normal and post-injury conditions. Combined with confocal microscopy, this approach supports efficient screening of whole-mount nerve preparations to evaluate fiber density, spatial distribution, axonal sprouting, and morphological characteristics. The refined technique provides a robust tool for advancing PNI research and may contribute to the development of more effective therapeutic strategies for nerve repair.

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.

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.

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.

Active sampling, such as respiration, is known to play a major role in modulating how sensory information is perceived and encoded in the field of olfaction. Hence, monitoring respiration is crucial for understanding olfactory-guided behavior and physiology. Several methods used to measure respiration, such as infrared cameras, piezoelectric sensors, video monitoring, temperature probes, intubation, and intranasal cannula, require the animal or at least its head to be fixed. However, telemetry-based sensors can be used wirelessly, allowing animals to move freely. Here, we describe the surgical protocol to implant a telemetry pressure sensor in the internal jugular vein to detect changes in thoracic pressure. The sensor can thus help in monitoring respiration by transmitting the signal wirelessly. We describe a way of inserting the probe into the right jugular vein aseptically while housing the transmitter underneath the skin on the back of the animal. Next, based on the optimal spot for the best signal, we secure the position of the probe and suture the skin. The animal then undergoes regular post-operative care with painkillers and soft diets for up to a week. The method offers two main advantages; first, it uses a strategy similar to the jugular vein catheterization, which is widely established in rodents. Second, it minimizes the need for extensive post-operative care, including not having to shift to a liquid diet post-recovery. This makes the animals fit for most behavioral experiments requiring water or food restrictions.

The neuromuscular junction (NMJ) is critical for muscle function, and its dysfunction underlies conditions such as sarcopenia and motor neuron diseases. Current protocols for assessing NMJ function often lack standardized stimulation parameters, limiting reproducibility. This study presents an optimized ex vivo method to evaluate skeletal muscle and NMJ function using the Aurora Scientific system, incorporating validated stimulation protocols for both nerve and muscle to ensure consistency. Key steps include tissue preparation in a low-calcium, high-magnesium solution to preserve NMJ integrity, determination of optimal muscle length, and sequential stimulation protocols to quantify neurotransmission failure and intratetanic fatigue. By integrating rigorous standardization, this approach enhances reproducibility and precision, providing a robust framework for investigating NMJ pathophysiology in aging and disease models.

The neuromuscular junction (NMJ) is a peripheral synaptic connection between a lower motor neuron and skeletal muscle fibre that enables muscle contraction in response to neuronal stimulation. NMJ dysfunction and morphological abnormalities are commonly observed in neurological conditions, including amyotrophic lateral sclerosis, Charcot–Marie–Tooth disease, and spinal muscular atrophy. Employing precise and reproducible techniques to visualise NMJs in mouse models of neuromuscular disorders is crucial for uncovering aspects of neuropathology, revealing disease mechanisms, and evaluating therapeutic approaches. Here, we present a method for dissecting the deep lumbrical and flexor digitorum brevis (FDB) muscles of the mouse hind paw and describe the process of whole-mount immunofluorescent staining for morphological analysis of NMJs. Similar whole-mount techniques have been applied to other muscles, such as the diaphragm; however, dense connective tissue in adult samples often impedes antibody penetration. Moreover, large hind limb muscles, including the gastrocnemius and tibialis anterior, are commonly used to examine NMJs but require embedding and cryosectioning. These additional steps increase the complexity and duration of the protocol and can introduce sectioning artefacts, including transection of NMJs and disruption of morphology. Using small hind paw muscles enables whole-mounting, which completely eliminates the requirement for embedding and cryosectioning. As a result, the entire neuromuscular innervation pattern can be visualised, allowing a more accurate assessment of NMJ development, denervation, and regeneration in mouse models of neurological disease and nerve injury, which can be applied across all postnatal ages.

Reconstructing single-neuron projectomes is essential for mapping the mesoscopic connectome and eventually for understanding brain-wide connectivity and diverse brain functions. The combination of sparse labeling techniques and large-scale and high-resolution optical imaging technologies has been revolutionizing the brain-wide reconstruction of single-neuron morphologies, as exemplified by the dataset for over 10,100 single-neuron projectomes of hippocampal neurons. Here, we illustrate a comprehensive protocol for large-scale single-neuron reconstruction in the mouse brain. This includes key steps and examples in imaging data preprocessing, neurite tracing, and registration into a template brain. These procedures enable efficient and accurate large-scale morphological reconstruction of single neurons in the mouse brain.