Neuroscience

Adaptive behaviors shaped by prior experience are essential for increasing animal survival. Aversive experiences play a pivotal role in memory formation and in updating subsequent learning rules. While the negative value of aversive signals, which are both necessary and sufficient to drive a conditioned response, is considered to be innately specified, it can also be subject to experience-dependent scaling. Previous reports demonstrated synaptic potentiation in nociceptive pathways following robust aversive learning. However, the neuronal basis of experience-dependent value updating remains largely unknown. Recently, we demonstrated that long-term potentiation (LTP) in the parabrachial-central amygdala (PB-CeA) pathway, an important circuit involved in pain processing and aversive learning, enhances the negative value and thereby updates future learning rules. Here, we present a protocol that combines behavioral analysis using pathway-specific optogenetic induction of in vivo LTP with mathematical modeling to examine value modification using Bayesian inference of the unconditioned stimulus value using the Rescorla–Wagner model. This protocol enables investigation of the mechanisms underlying experience-dependent value modulation and learning-rule changes in mice. Potentially, this protocol may provide a framework for understanding learning rules across a wide range of species and for the development of treatments for stress-related disorders.

Three-dimensional immunohistochemistry (3D-IHC) shows the organization of molecular assemblies in the context of tissue architecture. Deep and rapid antibody penetration into 3D tissues and highly sensitive detection are crucial for high-throughput analysis of 3D-IHC imaging. Here, we provide a detailed protocol for a nanobody (nAb)-based 3D-IHC technique, namely POD-nAb/FT-GO 3D-IHC, for high-speed and high-sensitivity detection of targets within 1-mm-thick mouse brain tissues. Peroxidase-fused nAb (POD-nAb) is a genetically encoded recombinant antibody, which consists of a camelid nAb and a variant of horseradish peroxidase, and fluorochromized tyramide-glucose oxidase (FT-GO) is a fluorescent tyramide signal amplification (TSA) system. POD-nAb/FT-GO 3D-IHC incorporates three main components: 1) tissue permeabilization, 2) POD-nAb binding, and 3) 3D-TSA reaction with FT-GO. POD-nAbs enhance signal penetration depth and allow for highly sensitive detection when combined with FT-GO signal amplification. By using the 3D-IHC protocol provided herein, we can visualize target molecules in mouse brain tissues of 1-mm thickness with drastic signal enhancement within three days. This protocol for POD-nAb/FT-GO 3D-IHC could facilitate structural and molecular interrogation of 3D tissues.

Animal and human stem cell–derived three-dimensional models to study physio-pathological brain functioning are becoming a gold standard for in vitro electrophysiology, as they enable the recapitulation of complex network properties by accounting for spatial architectural features that better reflect in vivo conditions than simpler 2D models. Standard planar multielectrode arrays (MEAs), typically providing tens of recording electrodes, are commonly used to record activity from 2D neuronal cultures. However, when adapted for use with 3D models, planar 2D MEAs showed limited effectiveness. The main issues are limited specimen adhesion to the chip, a low number of sensing elements, inability to retrieve signals from within the tissue, and reduced perfusion and vitality of the tissue in contact with sensors. To overcome these limitations, a new generation of microchip-based 3D high-density MEAs (3D HD-MEA) has been developed and validated in recent years. This technological advancement has improved the sensing capabilities and the vitality of 3D models, providing a tool tailored to maximize their potential. Here, we present an optimized protocol for neural network activity recordings in 3D models (including acute slices, brain spheroids, and organoids) from various brain regions using 3D HD-MEAs. First, we summarize the critical steps for 1) obtaining viable acute slices from the mouse cerebellum, cortico-hippocampal circuit, and prefrontal cortex, 2) establishing efficient coupling of the slices with the chip, and 3) performing recordings and analyses. We then describe the main procedures required to obtain human and animal brain spheroids and neural organoids, as well as standardized routines to perform effective recordings and analyses. For each section, we highlight the crucial steps, identify tips for specific applications, and propose troubleshooting procedures. For example, the same type of preparation (e.g., acute slices) requires different adjustments when working with different brain areas. The specific information provided here is intended to assist researchers in their daily efforts to obtain efficient and reproducible functional recordings from 3D models by using the cutting-edge technique of 3D HD-MEA.

Postnatal mouse retinal vascular development is a widely used model for studying retinal vascular diseases and evaluating candidate therapies. This is particularly relevant for inherited disorders such as familial exudative vitreoretinopathy (FEVR), in which impaired vascular growth and organization are central to disease pathogenesis. Numerous approaches have been used to assess retinal vasculature in mouse flat mounts, ranging from qualitative descriptions to limited quantitative measurements of vascular growth. However, phenotypic variability across genetic models, including different models of FEVR, complicates comparisons and underscores the need for standardized, comprehensive multi-parameter analyses that are suitable for rapid and cost-effective screening studies. We describe a standardized morphometric protocol using ImageJ software to quantitatively analyze mouse retinal vasculature in a reproducible manner. The protocol begins with measurement of areas of vascular disorganization (meshes) as well as total vascular and retinal area. Two defined regions in the peripheral and midperipheral retina are then selected to quantify cell clusters, followed by image processing, binarization, and skeletonization. From these processed images, vascular density, branch number, branch length and thickness, junction number, triple points, and box-counting fractal dimension and lacunarity are quantified. Overall, this protocol provides a rapid, cost-effective, and standardized framework for quantifying retinal vascular phenotypes across diverse mouse models. By capturing multiple structural features and accommodating phenotypic variability, it is well-suited for comparative studies and therapeutic screening in retinal vascular disease.

The mammalian central circadian clock resides in the suprachiasmatic nucleus (SCN) of the hypothalamus in the brain and is responsible for coordinating daily rhythms of biological processes spanning from gene expression to behavior. Light, the primary environmental zeitgeber, entrains the SCN via melanopsin-expressing intrinsically photosensitive retinal ganglion cells that project through the retino-hypothalamic tract. Altered circadian rhythms are common in individuals diagnosed with neurodevelopmental and neurodegenerative disorders, and often, associated with structural alterations of the SCN and impaired retinal input; importantly, these anomalies can be recapitulated in animal models. Here, we describe step-by-step protocols for quantitative histomorphometrical analysis of the SCN and the assessment of retinal–SCN connectivity, previously used in mouse models of neurodevelopmental and neurodegenerative disorders. These include measurement of the SCN area, perimeter, height and width using Nissl- or DAPI-stained coronal sections, as well as densitometric and plot profile analyses of cholera toxin β-subunit–labeled retinal projections using Axiovision or Fiji/ImageJ. The protocols incorporate standardized region-of-interest, measurements by masked observers, and consistent scaling procedures to enhance reproducibility. These methods provide a rigorous framework for detecting structural anomalies and connectivity defects in the circadian system and can be broadly applied to other experimental models of circadian dysfunction.

α-Synuclein (α-syn) aggregation has emerged as a key pathogenetic feature in several neurodegenerative disorders. The α-syn protein has various conformational strains, each with unique structural features that influence their cytotoxicity, propagation, and neuroinflammation. A post-translational modification known as O-GlcNAcylation has been found to influence the toxicity of α-syn and its propensity to aggregate. Difficulties in detecting and quantifying this modification are a major challenge to understanding its roles among the conformational forms of α-syn. We now describe a protocol for detecting O-GlcNAcylated α-syn that combines a click chemistry labeling approach and western blotting. This chemoenzymatic method involves the transfer of azido-modified galactose (GalNAz) from UDP-GalNAz to O-GlcNAcylated proteins, enabling their further functionalization with alkyne-containing polyethylene glycol of defined molecular weight. This protocol facilitates the determination of the glycosylation status of varying conformations of α-syn and their stoichiometric ratios.

Structural proteomics methods allow for the proteome-wide interrogation of protein structural differences between two different conditions. Limited proteolysis mass spectrometry (LiP-MS), as originally implemented by the Picotti lab, utilizes a promiscuous protease to cleave at solvent-exposed regions of a protein to encode structural information, which is then read out with mass spectrometry proteomics. Here, we present a protocol that details experimental steps and data analysis for a LiP-MS workflow. First, tissue is homogenized under native conditions and then subjected to limited proteolysis using proteinase K (PK). The samples are prepared for mass spectrometry, and data are acquired using either data-dependent acquisition (DDA) or data-independent acquisition (DIA). Raw data is processed using FragPipe, and raw ion abundances are processed in FragPipe Limited-Proteolysis Processor (FLiPPR). Proteins with structural changes between the two conditions are identified in a proteome-wide manner.

Despite substantial progress in preclinical cannabinoid research, translational studies on cannabis use disorders (CUD) are still insufficient due to the absence of robust, validated animal models that fully recapitulate the multifactorial clinical phenotype of human CUD. The complex nature of CUD and the incomplete understanding of its underlying neurobiological mechanisms contribute to the limited availability of effective treatments. To address this gap, we developed an operant conditioning–based mouse model that enables the identification of individual vulnerability or resilience to CUD development. This highly translational model is based on the Diagnostic and Statistical Manual of Mental Disorders, 5th Edition (DSM-5) criteria for substance use disorders. The model allows the assessment of addiction-like behaviors by evaluating three behavioral domains: 1) persistence of responding during periods of cannabinoid unavailability, 2) motivation for cannabinoid seeking measured using a progressive ratio schedule, and 3) compulsivity, assessed when cannabinoid reward is paired with an aversive consequence such as a mild electric foot shock. A major strength of this paradigm is its ability to quantify two phenotypic traits proposed as predisposing factors for addiction vulnerability and two parameters related to craving. In addition, the model is specifically designed to evaluate genetic and circuit-level manipulations using chemogenetic approaches, with minor modifications required by surgical viral-vector delivery. Using this protocol, we can determine whether altering the excitability of specific neural networks promotes resilience or vulnerability to developing cannabinoid addiction. Elucidating these mechanisms is expected to facilitate the identification of novel and more effective therapeutic interventions for CUD.

Organic solvent–based tissue clearing methods are widely used for whole-brain imaging but often compromise endogenous fluorescence. Existing protocols, such as iDISCO and fluorescence-preserving variants, have improved optical transparency but still present trade-offs between fluorescence retention, tissue stability, and workflow complexity. Here, we present MDISCO, a modified iDISCO-based clearing protocol designed to enhance preservation of endogenous fluorescence while maintaining high transparency and stable tissue morphology. MDISCO is directly compared with FDISCO+, an established fluorescence-preserving protocol, for the preservation of endogenous tdTomato and YFP. Performance across clearing steps is evaluated by measuring brain weight, anteroposterior and mediolateral dimensions, and optical transparency before and after solvent clearing and refractive index matching. Fluorescence preservation is assessed using whole-brain light-sheet microscopy with standardized imaging parameters to enable direct comparison. This protocol provides an accessible and high-throughput, reproducible workflow for solvent-based clearing with robust endogenous fluorescence preservation, offering clear advantages for whole-brain 3D imaging of genetically encoded fluorescent reporters.

Super-resolution imaging of synapses in intact brain tissue remains challenging because light scattering, photobleaching, and limited probe penetration, along with antigen accessibility within the densely packed postsynaptic densities (PSDs), constrain resolution and labeling efficiency. Here, we present a protocol utilizing thin brain cryosections and tau-stimulated emission depletion (STED) nanoscopy to visualize the intricate nano-architecture of excitatory synapses in situ. Slicing the brain into 6 μm sections allows for highly efficient and even penetration of probes throughout sections while ensuring that the resolution is not significantly impacted by the imaging depth of the tissue. We outline step-by-step instructions for labeling pre- and postsynaptic nano-architecture using antibodies and nanobodies, highlighting how fixative choice influences the labeling efficiency of synaptic proteins. While this protocol is compatible with both confocal and super-resolution imaging, when combined with rapid image acquisition times of tau-STED, it enables clear separation of key synaptic features in three dimensions with minimal photobleaching. Thus, this approach enables robust multiplex imaging of fluorescently labeled synaptic proteins in the brain, providing exceptional spatial resolution for visualization and quantification of synaptic nanoarchitecture in its native environment.