Journal of Visualized Experiments

We describe a standardized and reproducible procedure to generate human organotypic lung cultures from surgical lung resection for the study of respiratory infections. The protocol details tissue harvesting, biopsy punching, mechanical slicing, culture at the air-liquid interface. This technique enables robust ex vivo infections of human lung tissue with respiratory viruses, including Influenza A and Nipah. The described system can be used to study host-pathogen interactions, analyze innate immune responses, and evaluate antiviral candidates in physiologically relevant human lung tissue. For complete details on the use and execution of this protocol, please refer to Cezard et al1.

Optogenetic modulation of peripheral sensory nerve activity holds great potential for the treatment of sensory disorders. Here, we present a protocol for applying optogenetic techniques to peripheral sensory nerves using an adeno-associated virus (AAV) vector. We describe the procedure for gene transduction into dorsal root ganglion neurons via retrograde transport following intra-nerve AAV injection. We then outline a terminal, acute electrophysiological experiment to evaluate optogenetic effects at the level of the dorsal root. For complete details on the use and execution of this protocol, please refer to Kosugi et al1. Graphical abstract O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=195 SRC="FIGDIR/small/693516v1_ufig1.gif" ALT="Figure 1"> View larger version (33K): org.highwire.dtl.DTLVardef@5a2de4org.highwire.dtl.DTLVardef@7ad4f0org.highwire.dtl.DTLVardef@32113aorg.highwire.dtl.DTLVardef@10918c9_HPS_FORMAT_FIGEXP M_FIG C_FIG

The development of translational ovarian cancer models to investigate and overcome treatment resistance accounting for the impact of the tumor microenvironment is critical. Here, we present a protocol to establish a multicellular culture model that retains both genetic complexity and the microenvironment of patient tumors, is amenable for molecular and phenotypic analyses, and high throughput drug testing. We describe steps for culturing and characterizing stromal cells derived from cryopreserved and fresh samples and detail procedures for combining them with organoids. Graphical abstract O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=193 SRC="FIGDIR/small/686515v1_ufig1.gif" ALT="Figure 1"> View larger version (57K): org.highwire.dtl.DTLVardef@792c12org.highwire.dtl.DTLVardef@13c6680org.highwire.dtl.DTLVardef@6d73adorg.highwire.dtl.DTLVardef@1c56834_HPS_FORMAT_FIGEXP M_FIG C_FIG Before you beginThis protocol describes the in vitro generation of a complex 3D multicellular model (MC), mimicking relevant cellular and extracellular matrix (ECM) interactions in metastatic ovarian high-grade serous carcinoma (HGSC). First, cultures of stromal cells (cancer-associated fibroblasts (CAF), mesothelial cells and adipocytes) are generated. CAF and mesothelial cell cultures are established from fresh tumor tissues and/or from cryopreserved tissue digest and ascites fluid. Next, the identity of the stromal cells is evaluated using relevant markers, and the validated cultures are expanded and cryopreserved. Adipocytes are isolated from fresh tumor tissues and cultured in suspension for a short period before 3D embedding. Finally, previously established patient-derived cancer organoids1 are combined with relevant components of the tumor microenvironment (TME)2, including Type I collagen (main ECM protein in omental metastases) and stromal cells (Figure 1). The resulting MC model, which is viable for at least 14 days, can be used in various downstream applications. Here, we provide detailed protocols for two of them: high throughput drug sensitivity testing and single-cell RNA sequencing (scRNA-seq). O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=110 SRC="FIGDIR/small/686515v1_fig1.gif" ALT="Figure 1"> View larger version (30K): org.highwire.dtl.DTLVardef@13c3a1dorg.highwire.dtl.DTLVardef@10b56f6org.highwire.dtl.DTLVardef@17a6e8forg.highwire.dtl.DTLVardef@f09b27_HPS_FORMAT_FIGEXP M_FIG O_FLOATNOFigure 1.C_FLOATNO Overview of the samples and culture conditions used to establish stromal cell cultures, and their integration with patient-derived organoids to generate the MC model. C_FIG

SHORT ABSTRACTWe describe a novel micro-drive design, surgical implantation procedure, and post-surgery recovery strategy that allows for chronic field and single-unit recordings from up to sixteen brain regions simultaneously in juvenile and adolescent mice across a critical developmental window from p20 to p60 and beyond. LONG ABSTRACTIn vivo electrophysiology provides unparalleled insight into sub-second-level circuit dynamics of the intact brain and represents a method of particular importance for studying mouse models of human neuro-psychiatric disorders. However, such methods often require large cranial implants which cannot be used in mice at early developmental timepoints. As such, virtually no studies of in vivo physiology have been performed in freely behaving infant or juvenile mice, despite the fact that a better understanding of neurological development in this critical window is likely to provide unique insights into age-dependent developmental disorders such as autism or schizophrenia. Here, we describe a novel micro-drive design, surgical implantation procedure, and post-surgery recovery strategy that allows for chronic field and single-unit recordings from up to sixteen brain regions simultaneously in mice as they age from postnatal day 20 (p20) to postnatal day 60 (p60) and beyond, a time window roughly corresponding to human ages 2-years-old through adult. The number of recording electrodes and final recording sites can be easily modified and expanded, allowing flexible experimental control of in vivo monitoring of behavior- or disease-relevant brain regions across development.

Single-molecule pull-down (SIM-Pull) combined with TIRF microscopy enables direct visualization of proteins and multi-protein complexes. Here, we present an extended SIM-Pull protocol for analyzing protein interactions at the active zone and their ability to recruit isolated synaptic vesicles (SV). SV recruitment mediated by STX1A-SNARE or RIM1-Rab3a interactions, respectively; can be directly visualized and quantified. This technique opens new avenues to examine the subcellular vesicle-associated protein-protein interactions at a molecular level in a near-native cellular context. HighlightsO_LIExtended SIM-Pull protocol combining biochemical isolation with TIRF microscopy to study synaptic protein complexes at near-native environment C_LIO_LIEnables direct quantification of synaptic vesicle recruitment at the surface (active zone) via protein-mediated vesicle tethering C_LIO_LIAdaptable platform for probing molecular interactions of protein complexes within different neuronal, cellular or subcellular compartments C_LI Graphical abstract O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=198 SRC="FIGDIR/small/671146v1_ufig1.gif" ALT="Figure 1"> View larger version (57K): org.highwire.dtl.DTLVardef@1c42949org.highwire.dtl.DTLVardef@1727b9aorg.highwire.dtl.DTLVardef@110ec6aorg.highwire.dtl.DTLVardef@114ec7_HPS_FORMAT_FIGEXP M_FIG C_FIG Institutional permissionsAnimals were handled and maintained according to the guidelines laid down by the Animal Welfare Body (AWB) (Instantie voor Dierenwelzijn IvD) in line with the animal experimentation policy within Radboud University and RadboudUMC; under the license/protocol numbers 2021-0040-001/002 to Dr. Anne-Sophie Hafner.

Acute spinal cord slice electrophysiology is a powerful technique used in preclinical basic science research to investigate sensory and motor neuron function and pathophysiology. A major barrier that stands between implementing these findings into effective clinical treatments is the translational gap between rodent models and human patients. To date, no methods or protocols describe how to prepare viable human spinal cord slices for acute electrophysiological recordings. To bridge this translational divide, we describe here a protocol for the extraction of spinal cord tissue from consenting human organ donors and the preparation and sectioning of this tissue for acute spinal cord slice electrophysiology. With the collaboration of a transplant service and licensed surgeon, tissue can be extracted in 30-50 minutes. Acute spinal cord slices can then be prepared in the laboratory by trained graduate students in 2.5-5 hours, depending on the amount of tissue and scope of experiments. Using a viability stain to confirm that spinal slices are of sufficient quality to proceed, slices can then be used for either patch-clamp recordings to study the excitability of individual neurons or for high-density multielectrode array recordings to study intact sensory circuits. Slices remain viable for 4 to 8 hours, providing ample time for investigating synaptic and circuit-level signalling dynamics, including the use of pharmacological agents to probe the roles of specific molecular targets. The approaches described here can be implemented to improve translational physiological research and as a human tissue-based preclinical drug target identification and validation assay.

Advances in single-particle cryogenic electron microscopy (cryoEM) now allow for routine structure determination of well-behaved biological specimens to high-resolution. Despite advances in the electron microscope, direct electron detectors, and data processing software, the preparation of high-quality grids with thin layers of vitreous ice containing the specimen of interest in random orientations remains a critical bottleneck for many projects. Although numerous efforts have been dedicated to overcoming hurdles frequently encountered during specimen vitrification using traditional blot-and-plunge specimen preparation techniques, the development of blot-free grid preparation devices provide a unique opportunity to carefully tune ice thickness, particle density, and specimen behavior during the vitrification process for improvements in image quality. Here, we describe critical steps of high-quality grid preparation using a SPT Labtech chameleon, evaluation of grid quality/ice thickness using the chameleon software, high-throughput imaging in the electron microscope, and recommend steps for troubleshooting grid preparation when standard parameters fail to yield suitable specimen. Video LinkContents of this manuscript are available as a video tutorial. This video can be found here

In Alzheimers disease, there is an imbalance in neurotransmitter release and altered neuronal activation. We present a novel approach to analyze neuronal activity by combining local field potential (LFP) recording with microdialysis within the same animal. This method measures glutamate and GABA levels following chronic hippocampal amyloid-beta oligomer (A{beta}o) injections in rats. We outline the design of our electrode and canula, the surgical procedure, and the steps for LFP recording, interstitial fluid collection, and A{beta}o injections simultaneously in living animal. Graphical abstract O_FIG O_LINKSMALLFIG WIDTH=182 HEIGHT=200 SRC="FIGDIR/small/614333v2_ufig1.gif" ALT="Figure 1"> View larger version (37K): org.highwire.dtl.DTLVardef@155f3c0org.highwire.dtl.DTLVardef@1f60289org.highwire.dtl.DTLVardef@207ba2org.highwire.dtl.DTLVardef@94eeb3_HPS_FORMAT_FIGEXP M_FIG C_FIG

Mice navigate an odor plume with a complex spatiotemporal structure in the dark to find the source of odorants. This article describes a protocol to monitor behavior and record Ca2+ transients in dorsal CA1 stratum pyramidale neurons in hippocampus (dCA1) in mice navigating an odor plume in a 50 cm x 50 cm x 25 cm odor arena. An epifluorescence miniscope focused through a GRIN lens imaged Ca2+ transients in dCA1 neurons expressing the calcium sensor GCaMP6f in Thy1-GCaMP6f mice. The paper describes the behavioral protocol to train the mice to perform this odor plume navigation task in an automated odor arena. The methods include a step-by-step procedure for the surgery for GRIN lens implantation and baseplate placement for imaging GCaMP6f in CA1. The article provides information on real-time tracking of the mouse position to automate the start of the trials and delivery of a sugar water reward. In addition, the protocol includes information on using of an interface board to synchronize metadata describing the automation of the odor navigation task and frame times for the miniscope and a digital camera tracking mouse position. Moreover, the methods delineate the pipeline used to process GCaMP6f fluorescence movies by motion correction using NorMCorre followed by identification of regions of interest with EXTRACT. Finally, the paper describes an artificial neural network approach to decode spatial paths from CA1 neural ensemble activity to predict mouse navigation of the odor plume. SUMMARYThis protocol describes how to investigate the brain-behavior relationship in hippocampal CA1 in mice navigating an odor plume. This article provides a step-by-step protocol, including the surgery to access imaging of the hippocampus, behavioral training, miniscope GCaMP6f recording and processing of the brain and behavioral data to decode the mouse position from ROI neural activity.

Reporter-expressing recombinant virus represents an excellent option and a powerful tool to investigate, among others, viral infection, pathogenicity, and transmission, as well as to identify therapeutic compounds that inhibit viral infection and prophylactic vaccines. To combat the still ongoing coronavirus disease 2019 (COVID-19) pandemic, we have established a robust bacterial artificial chromosome (BAC)-based reverse genetics (RG) system to rapidly generate recombinant severe acute respiratory syndrome coronavirus 2 (rSARS-CoV-2) to study the contribution of viral proteins in viral pathogenesis. In addition, we have also engineered reporter-expressing recombinant viruses in which we place the reporter genes upstream of the viral nucleocapsid (N) gene to promote high levels of reporter gene expression that facilitates the study of SARS-CoV-2 in vitro and in vivo. Although successful, the genetic manipulation of the BAC containing the entire SARS-CoV-2 genome of [~]30,000 nucleotides, is challenging. Herein, we depict the technical details to engineer rSARS-CoV-2 expressing reporter genes using the BAC-based RG approach. We describe i) assembly of the full-length (FL) SARS-CoV-2 genome sequences into the empty pBeloBAC, ii) verification of the pBeloBAC-FL, iii) cloning of a Venus reporter gene into the pBeloBAC-FL, and iv) recovery of the Venus-expressing rSARS-CoV-2. By following this protocol, researchers with basic molecular biology and gene engineering techniques knowledge will be able to generate wild-type and reporter-expressing rSARS-CoV-2.

Breast cancer (BC) is the second leading cause of cancer-related death among women in the U.S. Organoid models of solid tumors have been shown to faithfully recapitulate aspects of cancer progression such as proliferation and invasion. Although patient-derived organoids (PDOs) and patient-derived xenograft organoids (PDXOs) are pathophysiologically relevant, they are costly to propagate, difficult to manipulate and comprised primarily of the most proliferative cell types within the tumor microenvironment (TME). These limitations prevent their use for elucidating cellular mechanisms of disease progression that depend upon tumor-associated stromal cells which are found within the TME and known to contribute to metastasis and therapy resistance. Here, we report on methods for cultivating epithelial-stromal multicellular 3D cultures. Advantages of these methods include a cost-effective system for rapidly generating organoid-like 3D cultures within scaffold-free environments that can be used to track invasion at single-cell resolution within hydrogel scaffolds. Specifically, we demonstrate how to generate these hetero-multicellular 3D cultures using BT-474 breast cancer cells in combination with fibroblasts (BJ-5ta), monocyte-like cells(THP-1) and/or endothelial cells (EA.hy926). Additionally, differential fluorescent labeling of cell populations enables time-lapse microscopy to define 3D culture assembly and invasion dynamics. Notably, the addition of any two stromal cell combinations to 3D cultures of BT-474 cells significantly reduces circularity of the 3D cultures, consistent of the presence of organoid-like or secondary spheroid structures. In tracker dye experiments, fibroblasts and endothelial cells co-localize in the peripheral organoid-like protrusions and are spatially segregated from the primary BT-474 spheroid. Finally, hetero-multicellular 3D cultures of BT-474 cells have increased hydrogel invasion capacity. Since we observed these protrusive structures in hetero-multicellular 3D cultures of both non-tumorigenic and tumorigenic breast epithelial cells, this work provides an efficient and reproducible method for generating organoid-like 3D cultures in a scaffold-free environment for subsequent analyses of phenotypes associated with solid tumor progression. SUMMARYThere is a critical need for 3D cancer models that capture hetero-cellular crosstalk to study cancer metastasis. Our study presents the generation of hetero-multicellular stromal-epithelial in a scaffold and scaffold free environment that can be used to study invasion and cellular spatial distributions.

Actin cytoskeleton-based materials are widely investigated as model cellular materials to elucidate physical mechanisms of cell mechanics, such as shape regulation and force production, as well as intriguing soft polymeric materials. In this method, we detail creating actin-based assemblies in vitro using purified protein for fluorescence microscopy studies. We polymerize long actin filaments in a sample chamber and use a polymer depletant to crowd filaments into a 2D-entangled network against a surface passivated with a surfactant layer. Adding skeletal muscle myosin II filaments in the presence of ATP induces contraction of the actin network. By bundling actin filaments with cross-linker, we tune the contractility of the assembly, transitioning from a material that buckles to a material that slides at the microscale. By reducing the length of the actin filaments through co-polymerizing actin in the presence of capping protein, we tune the material from being a 2D network to a liquid crystal. Cross-linking of dispersed short actin filaments results in 3D liquid crystal droplet formation.

Congenital vascular malformations are commonly caused by aberrant, genetic activation of class I phosphoinositide 3-kinase (PI3K) signalling. Advances in mechanistic understanding and therapeutic targeting of these disorders will be accelerated by high-fidelity, human disease models. Building on a previously optimised differentiation strategy, we present a validated workflow for PI3K inhibitor-free generation of arterial- and venous-like endothelial cells from human induced pluripotent stem cells (iPSCs) under defined, xeno-free conditions. We further report experimental analyses of endothelial maturation under flow, culture duration-dependent stability, and downstream molecular and phenotypic characterisation. By providing a reproducible human system for mechanistic and translational studies, this platform will enable disease-relevant modelling of PI3K-driven vascular malformations, including PIK3CA-related overgrowth spectrum (PROS) and PTEN hamartoma tumour syndrome (PHTS).

Stem cell differentiation with controlled geometry results in reproducible pattern formation. In contrast to constraining differentiating cells on micropatterned surfaces, we initialise colony formation using elastomeric stencils that adhere to culture dishes and create microwells with defined sizes and shapes. After colony formation, stencils are removed to allow colony growth and cell migration. Stencil fabrication involves mould production by photolithography followed by replica-moulding polydimethylsiloxane (PDMS). This approach produces reproducible two-dimensional organoids tailored for quantitative studies of growth control and pattern formation.

The mammalian heart relies on high rates of mitochondrial oxidative phosphorylation to meet its energy demand, with fatty acids serving as the primary fuel source in healthy adult hearts. While metabolic flexibility, the ability to switch between metabolic fuel substrates, is known to change during development and cardiac diseases, standardized methods for assessing substrate usage in intact, living cardiac tissue remain limited. Here, we present a protocol that adapts the Seahorse Mito Fuel Flex Test for use in living organotypic cardiac slices. This method enables the quantification of fuel dependency and capacity for fatty acids (FA), glucose (GLC), and glutamine (GLN) by sequentially inhibiting their respective mitochondrial oxidative phosphorylation pathways with the inhibitors etomoxir, UK5099, and BPTES. First, we validated the protocol by comparing results from organotypic cardiac slices to the standard published protocol using isolated adult mouse primary cardiomyocytes. Next, we demonstrated the sensitivity of this assay by modulating metabolism with AICAR, an AMPK activator, at varying concentrations, to demonstrate improved metabolism and then metabolic suppression at higher toxic doses. Finally, we applied this protocol to organotypic cardiac slices from different chambers of human donor hearts. This protocol provides a high-throughput, physiologically relevant platform for investigating cardiac metabolism, applicable across species and adaptable to other tissue types. It enables the study of metabolic remodeling in development and disease while overcoming the limitations of traditional cell-based assays by preserving native tissue architecture, physiology, and multicellular heterogeneity.

Cerebrospinal fluid (CSF) is a clear fluid surrounding and nourishing the brain and spinal cord. Molecular profiling of the CSF is a common diagnostic approach for central nervous system (CNS) diseases, including infectious diseases, autoimmune disorders, brain hemorrhage and traumatic brain injury, CNS tumors, and Alzheimers disease1-10. Rodent models are critical for investigating CNS disease mechanisms and therapeutics, however, both collecting CSF and injecting materials into CSF in small animals are technically challenging and often result in high rates of postoperative mortality. Here, we present an easy-to-practice and cost-effective protocol with minimum instrument requirements to access the CSF in live rodents for collection and infusion purposes. By introducing a metal needle tool bent at a unique angle and length, we could steadily reach the CSF via the foramen magnum. Compared with prior methods, this protocol requires neither the operator to discern the changes in resistance from solid tissues while puncturing the needle, nor surgical opening of the skin and muscle covering the rodent neck. Using this method, we frequently obtain 5-15 L of CSF from mice and 70-120 L from rats to enable diverse downstream analyses including mass spectrometry. Due to the minimal invasiveness, this procedure allows iterative CSF collection from the same animal every few days - a major improvement over prior protocols that require extensive surgical operations. Moreover, we demonstrate that this method could be used for injecting desired solutions including dyes into mouse CSF with high success rates. Our method shortens the time required for CSF collection or injection to 3-5 minutes. Notably, we could reach near 100% postoperative recovery rates in both mice and rats even with repetitive collections. Together, we establish an efficient and minimally-invasive protocol for collecting CSF and inoculating reagents into the CSF in live rodents to enable various longitudinal studies at the forefronts of CNS investigation.

Spatially resolved transcriptomics was named Method of the Year 20201 and has continued to evolve rapidly since then, providing novel insights in development, physiology, and disease processes. Many approaches now offer excellent performance in formalin-fixed paraffin embedded tissue, opening up a wealth of archived clinical samples to these high-throughput spatial biology modalities. In this Chapter, we introduce a methodology to apply 10x Xenium In Situ to FFPE tissue microarrays. Using this method we have achieved a cost-effective approach to profile hundreds of genes with subcellular resolution in over a hundred human liver samples. Our method can easily be adapted to other tumour types, diseases, or model systems.

The Default Mode Network (DMN) is a central large-scale brain network implicated in a range of cognitive functions and neuropsychiatric disorders, such as major depressive disorder (MDD). Studying the DMNs complex dynamics in animal models provides invaluable insights into its function in both healthy and pathological states. However, performing a stable, long-term, and large-scale electrophysiological recordings from the multiple, deep, and distributed nodes of the DMN in awake, behaving mice has been a significant challenge. Here, we present a novel, two-phase surgical protocol developed to create a large (4x7.6 mm), durable, and resealable cranial window in mice. The procedure is designed to preserve the integrity of the dura mater, which is paramount for long-term brain health and recording stability. This window facilitates repeated, longitudinal recordings from over 1,000 electrodes simultaneously by combining surface-level micro-electrocorticography ({micro}ECoG) with two high-density intracranial electrode probes, allowing unprecedented access to the DMN. This technique provides a robust platform for multi-modal, multi-scale interrogation of network-wide electrophysiological dynamics over several weeks, opening new avenues for investigating the neuroplastic changes underlying the pathophysiology of brain disorders and for evaluating the chronic effects of novel therapeutics. SUMMARYThis protocol describes a two-phase surgical method to create a large, resealable, dura-sparing cranial window in mice. This technique enables chronic, multi-modal electrophysiological recordings from distributed, deep brain networks, such as the Default Mode Network, over several weeks.

Single-nucleus RNA sequencing enables high-resolution transcriptomic profiling of brain tissue, facilitating detailed analysis of cell identity in models of neurodegeneration and repair. Here, we describe a protocol for isolating nuclei from long-term human stem cell-derived grafts in the rat brain, incorporating vibratome sectioning, graft dissection, nuclear extraction, and fluorescence-activated sorting. This workflow supports analysis of human neurons embedded within host tissue or sensitive to dissociation, offering a powerful approach to assess graft composition, integration, and neuronal identity in living brain. For complete details on the use and execution of this protocol, please refer to Fiorenzano et al.1 Subject areasSingle nucleus RNA sequencing, stem cell biology, neuroscience, transplantation, cell replacement therapy Graphical abstract O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=61 SRC="FIGDIR/small/672369v1_ufig1.gif" ALT="Figure 1"> View larger version (18K): org.highwire.dtl.DTLVardef@1ac3b90org.highwire.dtl.DTLVardef@7aab65org.highwire.dtl.DTLVardef@18a8fd4org.highwire.dtl.DTLVardef@1e8bfae_HPS_FORMAT_FIGEXP M_FIG C_FIG HighlightsO_LIStep-by-step vibratome sectioning of xenografted rat brain tissue. C_LIO_LIPrecise dissection of human stem cell-derived grafts from host brain sections. C_LIO_LIExtraction of intact nuclei from fragile, grafted neurons. C_LIO_LIIsolation of single nuclei via fluorescence-activated nuclei sorting (FANS) for snRNA-seq sample preparation. C_LI

Two-bottle choice home cage drinking is one of the most widely used paradigms to study ethanol consumption in rodents. In its simplest form, animals are provided with access to two drinking bottles, one of which contains regular tap water and the other ethanol, for 24 hr/day with daily intake measured via change in bottle weight over the 24 hr period. Consequently, this approach requires no specialized laboratory equipment. While such ease of implementation is likely the greatest contributor to its widespread adoption by preclinical alcohol researchers, the resolution of drinking data acquired using this approach is limited by the number of times the researcher measures bottle weight (e.g., once daily). However, the desire to examine drinking patterns in the context of overall intake, pharmacological interventions, and neuronal manipulations has prompted the development of home cage lickometer systems that can acquire data at the level of individual licks. Although a number of these systems have been developed recently, the open-source system, LIQ HD, has garnered significant attention in the field for its affordability and user friendliness. Although exciting, this system was designed for use in mice. Here, we review appropriate procedures for standard and lickometer-equipped two-bottle choice home cage drinking. We also introduce methods for adapting the LIQ HD system to rats including hardware modifications to accommodate larger cage size and a redesigned 3D printed bottle holder compatible with standard off-the-shelf drinking bottles. Using this approach, researchers can examine daily drinking patterns in addition to levels of intake in many rats in parallel thereby increasing the resolution of acquired data with minimal investment in additional resources. These methods provide researchers with the flexibility to use either standard bottles or a lickometer-equipped apparatus to interrogate the neurobiological mechanisms underlying alcohol drinking depending on their precise experimental needs. SUMMARYThis protocol describes a standard intermittent-access two-bottle choice home cage drinking paradigm to model alcohol consumption in rats. In addition, it provides step-by-step instructions to augment the standard protocol with a DIY lickometer system that enables microstructural analysis of drinking behavior.