神经科学

Transfecting neurons remains technically challenging due to their sensitivity. Conventional methods, such as Lipofectamine 2000 or Lipofectamine RNAiMAX, often result in significant cytotoxicity, which limits their utility. Although lentiviral transfection offers high efficiency, it is hindered by high costs and complex procedures. This experiment employs a small interfering RNA (siRNA)-specific transfection reagent from the Kermey company. This reagent is a novel nanoparticle-based lipid material designed for the efficient delivery of oligonucleotides, including siRNA, into a wide range of cell types. Its efficacy in achieving high transfection efficiency in neurons, however, has not yet been established. After several days of in vitro neuronal culture, researchers can perform a simple transfection procedure using this reagent to achieve robust transfection efficiency. Notably, the protocol does not require medium replacement 6–8 h post-transfection, streamlining the workflow and minimizing cellular stress.

To study gene function in regulating rodent retinal waves during development, an efficient method for gene delivery into whole-mount retinas is required while preserving circuit functionality for physiological studies. We present an optimized electroporation protocol for developing rodent retinal explants. The procedure includes the fabrication of horizontally aligned platinum electrodes and the placement of retinal explants between them to generate a uniform electric field for high transfection efficiency. The entire process—dissection and electroporation—can be completed within 1–2 h. Successful transfection is verified by fluorescence microscopy, and physiological assays such as patch-clamp recordings and live imaging can be performed within 1–4 days following electroporation. This rapid and reliable protocol enables functional analysis for a specific gene in regulating retinal waves and can be adapted to other organotypic slice cultures.

Autonomic regulation of heart and respiratory rates is essential for understanding brain–body interactions in health and disease. Preclinical cardiovascular recordings are often performed under anesthesia or via telemetry, both of which introduce physiological confounds such as stress or impaired recovery due to the need for acute or chronic implantation of sensors. Here, we present a minimally invasive protocol for simultaneous acquisition of high-quality electrocardiography and respiratory signals in awake mice. Using an in-house-modified physiological monitor in awake, head-fixed mice that were briefly habituated to experimental conditions, we ultimately enable stable, long-term physiological recordings alongside in vivo microscopy. This protocol provides a robust, low-stress method for acquiring physiological signals, enabling the simultaneous study of cardiovascular–cerebral dynamics in awake head-fixed mice, thereby enhancing the translational relevance of preclinical measurements.

Primary cilia are evolutionarily conserved organelles that play critical roles in brain development. In the developing cortex, neural progenitors extend their primary cilia into the ventricular surface, where the cilia act as key signaling hubs. However, visualizing these cilia in a systematic and intact manner has been challenging. The commonly used cryostat sectioning only provides a limited snapshot of cilia on individual sections, and this process often disrupts the ciliary morphology. By contrast, the previously established whole-mount technique has been shown to preserve ciliary architecture in the adult mouse cortex. Here, we adapt and optimize the whole-mount approach for embryonic and neonatal brain, allowing robust visualization of ciliary morphology at the ventricular surface during development. This protocol describes step-by-step procedures for whole-mounting and immunostaining delicate embryonic and neonatal mouse cortices, enabling direct visualization of cilia in neural progenitors in the developing brain.

Hair cells are the sensory receptors of the auditory and vestibular systems in the inner ears of all vertebrates. Hair cells also serve to detect water flow in the lateral line system in amphibians and fish. The zebrafish lateral line serves as a well-established model for investigating hair cell development and function, including research on genetic mutations associated with deafness and environmental factors that cause hair cell damage. Rheotaxis, the ability to orient and swim in response to water flow, is a behavior mediated by multiple sensory modalities, including the lateral line organ. In this protocol, we describe a rheotaxis assay in which station holding behavior, which employs positive rheotaxis to maintain position in oncoming water flow, serves as a sensitive measure of lateral line function in larval zebrafish. This assay provides a valuable tool for researchers assessing the functional consequences of genetic or environmental disruptions of the lateral line system.

Optogenetic stimulation of peripheral motor nerves is a promising technique for modulating neural activity via illumination of light-sensitive ion channels known as opsins. Stimulating muscle activity through this method offers many advantages, such as a physiological recruitment order of motor units, reduced fatigue, and target-specific stimulation, which make it a favorable option for use in many neuroscience and motor rehabilitation applications. To enable such optical stimulation, opsin expression in peripheral nerves can be achieved either with transgenic animal models or through injection of viral vectors. In this protocol, we describe a method for driving peripheral nerve opsin expression via intramuscular adeno-associated virus (AAV) injection with the goal of enhancing virus uptake by targeting injections to neuromuscular junctions with electrical stimulation. We also describe procedures for non-invasively assessing functional opsin expression over time with transdermal optical stimulation of opsin-labeled nerves and electromyography (EMG) recordings. The presence of time-locked EMG spikes 4–8 ms after each stimulation pulse demonstrates that functional opsin expression is present at a given assessment time point. Onset of functional optical sensitivity generally occurs 2–4 weeks following virus injection, and sensitivity generally peaks or plateaus between 6–10 weeks. Stimulation sequences such as light intensity, stimulation pulse width, and frequency sweeps provide further information on functional opsin expression at the testing timepoint. The methods presented here can be used for driving functional opsin expression with a standard AAV6 vector commonly used in similar experiments or as a protocol for assessing peripheral nerve opsin expression with novel viral vectors.

Developing preclinical animal models that faithfully mimic the progressive nature of Parkinson’s disease (PD) is crucial for advancing mechanistic insights as well as therapeutic discovery. While recombinant adeno-associated virus (rAAV)-driven α-synuclein overexpression is widely used, its reliance on high viral titers introduces nonspecific toxicity and limits physiological relevance. The SynFib model, which combines modest rAAV-driven α-synuclein expression (Syn) with α-synuclein preformed fibril (PFF) seeding (Fib), has shown promise in reproducing PD-like pathology. However, current implementations of this SynFib model have largely been confined to rats and require sequential surgeries, which increase animal distress and reduce reproducibility. Here, we present a streamlined protocol to generate a SynFib mouse model of PD that integrates rAAV-α-synuclein delivery and PFF injection into a single stereotaxic surgery. Using fine glass capillaries, this method prevents backflow of injected material, reduces injection-induced trauma, minimizes neuroinflammation, and ensures robust lesion development. This streamlined mouse model provides a reproducible and practical system to investigate α-synuclein-associated pathology and serves as a versatile platform for preclinical testing of potential therapeutics for PD.

Detecting the proliferation of cells with copper(I)-catalyzed azide-alkyne cycloaddition (click chemistry) and the thymidine analogue, 5-ethynyl-2’-deoxyuridine (EdU), is a simpler and more versatile method than traditional antibody-based approaches. Instead of the harsh series of steps typically used for 5-bromo-2’-deoxyuridine (BrdU) detection, detecting EdU does not require DNA denaturation and is suitable for use with other applications. This approach was implemented in an animal model of ischemic stroke. The following protocol details how to use EdU to label, track, and visualize leukocyte recruitment for flow cytometry and fluorescence microscopy, including the processes for EdU injection and blood and tissue sample preparation. Considerations for timing, dosing, and cell viability are also outlined to tailor the protocol to experimental needs. This method could be applied to various models that require extended tracking periods, as the signal from EdU can last several cell divisions, depending on cell type and condition.

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.