Neuroscience

Research on brain disorders, particularly in the field of oncology, requires in vivo models to evaluate various therapeutic approaches, including intracerebral drug delivery. To meet this requirement, the implantation of intracerebral cannulas offers a reliable method for administering candidate therapeutics directly into the brain. This protocol describes a surgical technique for cannula implantation in mice, enabling repeated administration of therapeutic compounds in the context of glioblastoma treatment. The method was designed with an emphasis on using accessible, easy-to-handle, and sterilized tools to optimize surgical outcomes. Particular attention was also given to animal welfare, notably through refined procedures for asepsis, anesthesia, and postoperative care.

Oxygen tension is a key regulator of early human neurogenesis; however, quantifying intra-tissue O₂ in 3D models for an extended period remains difficult. Existing approaches, such as needle-type fiber microsensors and intensity-based oxygen probes or time-domain lifetime imaging, either perturb the organoids or require high excitation doses that limit the measurement period. Here, we present a step-by-step protocol to measure intra-organoid oxygen in human cerebral organoids (hCOs) using embedded ruthenium-based CPOx microbeads and widefield frequency-domain fluorescence lifetime imaging microscopy (FD-FLIM). The workflow covers dorsal/ventral cerebral organoid patterning, organoid fusion at day 12 with co-embedded CPOx beads, standardized FD-FLIM acquisition (470-nm external modulation, 16 phases at 50 kHz, dual-tap camera), automated bead detection and lifetime extraction in MATLAB, and session-matched Stern–Volmer calibration with Ru(dpp)₃(ClO₄)₂ to convert lifetimes to oxygen concentration. The protocol outputs per-bead oxygen maps and longitudinal patterns stratified by bead location (intra-organoid vs. gel) and sample state (healthy vs. abnormal), enabling direct linkage between developmental growth and oxygen dynamics.

Vestibulo-ocular reflexes (VORs) are compensatory ocular reflexes that maintain stable vision during head movements. In research, VORs encompass angular VOR (aVOR) and off-vertical axis rotation (OVAR) tests, which various groups have employed to assess vestibular function in mice. This protocol outlines the process for measuring VORs in mice, including eye rotation calibration, immobilizing the mouse with a noninvasive setup, configuring the aVOR and OVAR stimulus modes, and interpreting the obtained waveforms to derive VOR values. As technology advances, VORs are expected to yield more qualitative and quantitative insights into the function of the horizontal semicircular canal cristae (HSCC) and the otolith organs. This methodology can serve as a standard for evaluating common vestibular deficits in mice.

Cerebrospinal fluid-contacting neurons (CSF-cNs) are a specialized group of multifunctional neurons located around the central canal of the spinal cord. They play critical roles in motor regulation, postural maintenance, and spinal cord injury repair. However, the molecular mechanisms underlying the multifunctionality of CSF-cNs remain poorly understood, partly due to the lack of established in vitro methods for their efficient selection and purification, which significantly hinders mechanistic investigations. In this study, we describe a standardized method using a PKD2L1 promoter-driven lentiviral system, which enables effective enrichment and identification of CSF-cNs in vitro through GFP labeling and puromycin selection. This protocol includes key steps such as construction of the PKD2L1 promoter-driven lentiviral vector, spinal cord tissue collection and digestion from neonatal mice, lentiviral infection, antibiotic selection, and immunofluorescence-based identification of CSF-cNs. Our method provides a reliable platform for obtaining high-purity CSF-cNs (>99%), which facilitates their functional and mechanistic studies for regenerative approaches in vitro.

Nociception is critically shaped by descending modulation of spinal circuits, yet its cellular and synaptic mechanisms remain poorly defined. Elucidating these mechanisms is technically challenging, as it requires simultaneous activation of primary afferents and descending fibers while monitoring the functioning of individual spinal neurons. Here, we present a method to investigate the influence of the rostral ventromedial medulla (RVM), a principal supraspinal structure mediating descending modulation, on the activity of spinal lamina I neurons. Our approach combines electrophysiological recordings in ex vivo intact spinal cord preparation with optogenetics, granting several advantages. First, ex vivo preparation spares rostrocaudal and mediolateral spinal architecture, preserving lamina I as well as primary afferent and descending inputs. Second, virally mediated channelrhodopsin-2 (ChR2) expression enables selective photostimulation of RVM-originating fibers. When coupled with patch-clamp recordings, this photostimulation allows identifying postsynaptic inputs from RVM to spinal neurons and revealing RVM-dependent presynaptic inhibition of primary afferent inputs. Overall, our approach is well-suited for investigating both pre- and postsynaptic mechanisms of descending modulation in physiological and pathological pain conditions.

Telomere length maintenance is strongly linked to cellular aging, as telomeres progressively shorten with each cell division. This phenomenon is well-documented in mitotic, or dividing, cells. However, neurons are post-mitotic and do not undergo mitosis, meaning they lack the classical mechanisms through which telomere shortening occurs. Despite this, neurons retain telomeres that protect chromosomal ends. The role of telomeres in neurons has gained interest, particularly in the context of neurodegenerative diseases such as amyotrophic lateral sclerosis (ALS), where aging is a major risk factor. This has sparked interest in investigating telomere maintenance mechanisms in post-mitotic neurons. Nevertheless, most existing telomere analysis techniques were developed for and optimized using mitotic cells, posing challenges for studying telomeres in non-dividing neuronal cells. Thus, this protocol adapts an already established technique, the combined immunofluorescence and telomere fluorescent in situ hybridization (IF-FISH) on mitotic cells to study the processes occurring at telomeres in cortical neurons of the mouse ALS transgenic model, TDP-43 rNLS. Specifically, it determines the occurrence of DNA damage and the alternative lengthening of telomeres (ALT) mechanism through simultaneous labeling of the DNA damage marker, γH2AX, or the ALT marker, promyelocytic leukemia (PML) protein, together with telomeres. Therefore, the protocol enables the visualization of DNA damage (γH2AX) or the ALT marker (PML) concurrently with telomeres. This technique can be successfully applied to brain tissue and enables the investigation of telomeres specifically in cortical neurons, rather than in bulk tissue, offering a significant advantage over Southern blot or qPCR-based techniques.

Three-dimensional (3D) human brain tissue models derived from induced pluripotent stem cells (iPSCs) have transformed the study of neural development and disease in vitro. While cerebral organoids offer high structural complexity, their large size often leads to necrotic core formation, limiting reproducibility and challenging the integration of microglia. Here, we present a detailed, reproducible protocol for generating multi-cell type 3D neurospheres that incorporate neurons, astrocytes, and optionally microglia, all derived from the same iPSCs. While neurons and astrocytes differentiate spontaneously from neural precursor cells, generated by dual SMAD-inhibition (blocking BMP and TGF-b signaling), microglia are generated in parallel and can infiltrate the mature neurosphere tissue after plating neurospheres into 48-well plates. The system supports a range of downstream applications, including functional confocal live imaging of GCaMP6f after adeno-associated virus (AAV) transduction of neurospheres or immunofluorescence staining after fixation. Our approach has been successfully implemented across multiple laboratories, demonstrating its robustness and translational potential for studying neuron–glia interactions and modeling neurodegenerative processes.

Accurate labeling of excitatory postsynaptic sites remains a major challenge for high-resolution imaging due to the dense and sterically restricted environment of the postsynaptic density (PSD). Here, we present a protocol utilizing Sylites, 3 kDa synthetic peptide probes that bind with nanomolar affinity to key postsynaptic markers, PSD-95 and Gephyrin. eSylites (excitatory Sylites) specifically target the PDZ1 and PDZ2 domains of PSD-95, enabling precise and efficient labeling of excitatory postsynaptic density (ePSD). In contrast, iSylites (inhibitory Sylites) bind to the dimerizing E-domain of the Gephyrin C-terminus, allowing selective visualization of inhibitory postsynaptic density (iPSD). Their small size reduces linkage error and enhances accessibility compared to conventional antibodies, enabling clear separation of PSD-95 nanodomains in super-resolution microscopy. The protocol is compatible with co-labeling using standard antibodies and integrates seamlessly into multichannel immunocytochemistry workflows for primary neurons and brain tissue. This method enables robust, reproducible labeling of excitatory synapses with enhanced spatial resolution and can be readily adapted for expansion microscopy or live-cell applications.

Human induced pluripotent stem cell (hiPSC)-derived motor neurons (MNs) provide a critical source for the study of motor neuron diseases (MNDs), which has been hindered by the lack of appropriate disease models for many years. Although many spinal MN differentiation protocols have been established by mimicking in vivo neurogenesis using extrinsic signaling molecules, substantial variations in the duration and efficiency persist due to inconsistencies in concentrations, timing, and delivery methods of these molecules. Here, we present an efficient monolayer culture differentiation strategy that enables the generation of enriched CHAT⁺ spinal MNs (sMNs) in 18 days and functional sMNs exhibiting extensive network activities, as confirmed by multielectrode array (MEA), within 28 days. Therefore, this optimized MN differentiation protocol facilitates the production of mature sMNs for MND research, high-throughput drug screening, and potential cell replacement therapies.

Changes in learning and memory are important behavioral readouts of brain function across multiple species. In mice, a multitude of behavioral tasks exist to study learning and memory, including those influenced by extrinsic and intrinsic forces such as stress (e.g., escape from danger, hunger, or thirst) or natural curiosity and exploratory drive. The novel object recognition (NOR) test is a widely used behavioral paradigm to study memory and learning under various conditions, including age, sex, motivational state, and neural circuit dynamics. Although mice are nocturnal, many behavioral tests are performed during their inactive period (light phase, subjective night) for the convenience of the diurnal experimenters. However, learning and memory are strongly associated with the animal’s sleep-wake and circadian cycles, stressing the need to test these behaviors during the animals’ active period (dark phase, subjective day). Here, we develop a protocol to perform the NOR task during both light (subjective night) and dark (subjective day) phases in adult mice (4 months old) and provide a flexible framework to test the learning and memory components of this task at distinct times of day and associated activity periods. We also highlight methodological details critical for obtaining the expected behavioral responses.