Neuroscience

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.

The Morris water maze (MWM) is one of the most widely used procedures to assess hippocampus-dependent spatial learning and memory in rodents. By varying test protocols, researchers can test several different domains of learning and memory. Over multiple testing days, animals learn to swim to a platform hidden just under the water surface by using the spatial relationship between distal cues and the platform. Probe trials, where the platform is rendered unavailable, measure rodents’ spatial bias for the area where the platform was previously located. The ability of researchers to control the availability of the platform “on-demand” offers both practical and methodological advantages. Despite MWM’s prominence in the field of behavioral neuroscience, the high cost of purchasing a commercial MWM package is often prohibitively expensive for many research labs, especially on-demand platforms. Here, we describe a low-cost strategy for a build-your-own MWM that includes a remote-controlled on-demand platform (~530 USD) and tank (~550 USD). It is our hope that disseminating low-cost strategies aimed at expanding access to high-quality research tools at underfunded research institutions will accelerate biomedical discovery and foster further innovation.

Understanding the nanoscale organization and molecular rearrangement of synaptic components is critical for elucidating the mechanisms of synaptic transmission and plasticity. Traditional synaptosome isolation protocols involve multiple centrifugation and resuspension steps, which may cause structural damage or alter the synaptosomal fraction, compromising their suitability for cryo-electron tomography (cryo-ET). Here, we present an ultrafast isolation method optimized for cryo-ET that yields two types of synaptosomal fractions: synaptosomes and synaptoneurosomes. This streamlined protocol preserves intact postsynaptic membranes apposed to presynaptic active zones and produces thin, high-quality samples suitable for in situ structural studies. The entire procedure, from tissue homogenization to vitrification, takes less than 15 min, offering a significant advantage for high-resolution cryo-ET analysis of synaptic architecture.

The phototransduction cascade allows photoreceptors to detect light across a wide range of intensities without saturation, with cGMP serving as the second messenger and calcium feedback as the key regulatory mechanism. While experimental evidence suggests that cAMP may also play a role in modulating this cascade, such regulation would necessitate rapid changes in cAMP levels on a timescale of seconds. However, data on the dynamics of intracellular cAMP changes in photoreceptors remain scarce, primarily due to the limitations of conventional fluorescence-based methods in this specialized sensory system. To address this gap, we developed a methodology combining rapid cryofixation of retinal samples following light stimulation with the isolation of outer segment preparations. The rapid cryofixation setup comprises six computer-controlled sections, each with a high-speed stepper motor-driven lever that rapidly moves the specimen in a 180° arc within ~80 ms to press it against a liquid nitrogen-cooled copper cylinder for fixation. Using highly sensitive metabolomics techniques, we measured cAMP levels in these samples. This approach enables the investigation of rapid cAMP dynamics and its potential regulatory role in phototransduction, providing a foundation for understanding the interplay between cAMP and PKA signaling in photoreceptor function.

High-throughput sequencing has created a tremendous amount of information about the genes expressed in various cell types and tissues throughout the body. As such, there is a need for a quick and effective method to knock down genes of interest in order to investigate their roles. While there are many approaches for this in mammalian models, there are limited ways to knock down genes of interest in adult zebrafish. Unlike mammals, zebrafish have the natural ability to regenerate their neurons after injury or disease is detected, making them a staple in regenerative studies. Unfortunately, current approaches for gene knockdown in the retina of adult zebrafish are costly and provide a barrier for many scientists. We provide two cost-effective approaches for targeted gene knockdowns in adult zebrafish retinas. We describe this approach through the use of Vivo-morpholinos and lipid-encapsulated siRNAs that target the expression of the proliferating cell nuclear antigen (PCNA) gene in adult zebrafish. We also describe how to collect and process retina samples for downstream immunohistochemistry, imaging, and quantification. Overall, this protocol will provide researchers with a straightforward, cheap, and effective method to perform targeted gene knockdowns in adult zebrafish retinas.

Cognitive flexibility is a process that involves dynamically adapting behavior to obtain a desired series of outcomes during continuously changing stimulus-response-reward associations. Attentional set-shifting is a multi-modal decision-making paradigm that tests cognitive flexibility, which can be useful for investigating neural circuitry that is disrupted in multiple neuropsychiatric disorders, including schizophrenia, Alzheimer’s disease, depression, and addiction. The canonical human attentional set-shifting paradigm for measuring cognitive flexibility is the Wisconsin Card Sorting Test, in which subjects sort cards based on rules that change periodically and adapt their behavior by utilizing feedback in the form of correct and incorrect outcomes. To transfer these tests to rodent models, previous techniques involved attentional set-shifting paradigms in free-roaming test chambers, in which animals associated cues with reward locations. The protocol presented here involves an analogous attentional set-shifting paradigm, in which water-restricted mice are head-restrained on a platform and must keep track of periodically switching stimulus-response-outcome associations. The mice must learn to associate odor or whisker vibration cues with a binary directional lick response that triggers water delivery. The mice are trained to respond by licking one of two spouts, in which the correct decision is dependent on the current stimulus rule. This protocol allows for behaviorally measuring cognitive flexibility alongside neural activity by pairing the head-restrained paradigm with 2-photon calcium imaging, optogenetics, and extracellular and intracellular physiology.

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.