We would like to thank the viral core NRDDC at the Jan and Dan Duncan Neurological Institute for producing the AAVs and lentiviruses used in these experiments. Additionally, we would like to thank the Baylor College of Medicine Center for Comparative Medicine for mouse husbandry and general maintenance of the mice used. We would like to thank the American Heart Association for their support under award number 20PRE35040011, and BRASS: Baylor Research Advocates for Student Scientists for their support (PJH). Finally, we would like to thank Logos for providing our lab with the Logos X-Clarity electrophoretic tissue clearing system.

The following protocol follows all animal care guidelines for Baylor College of Medicine.
1. Dissection and tissue preparation
<ol>
	<li>Euthanize the mouse with an overdose of isoflurane by placing the mouse in a closed container with a towel soaked in isoflurane (or by other IUCAC approved means).</li>
	<li>Perfuse the animal transcardially using a 25 G needle with 10 mL of ice cold PBS, followed by 10 mL of 4% PFA.</li>
	<li>Dissect the brain region (or tissue) of interest.</li>
	<li>Place the dissected tissue into 4% PFA overnight at 4 &#176;C. Proper fixation is key for this protocol. Do not skip or shorten this step.</li>
	<li>After fixation in 4% PFA for at least 12 h, transfer the tissue to a 4% acrylamide hydrogel mixture for 24 h at 4 &#176;C. 
	&#8203;NOTE: When thawing the hydrogel, ensure that it has not polymerized, this can happen if the hydrogel becomes too warm. Thaw on ice to prevent premature polymerization.</li>
</ol>
2. Tissue clearing
<ol>
	<li>Place the brain tissue (still submerged in hydrogel) in a vacuum incubator for 3 h at 37 &#176;C&#160;with a -90 kPa vacuum. Leave the tube top unscrewed to allow the vacuum to form properly.</li>
	<li>Wash the tissue with PBS for 10 min at room temperature (25 &#176;C) with gentle shaking.</li>
	<li>Place the polymerized tissue sample in the electrophoresis chamber, keeping note of the orientation of the tissue within the chamber.</li>
	<li>Fill the chamber and reservoir with the supplied electrophoresis SDS buffer.</li>
	<li>Run the sample at 70 V, 1 A, and 35 C, with constant current for about 2 h/mm of tissue.
	<ol>
		<li>Check the sample periodically; it may require more time in the chamber to properly clear. A good starting point for clearing is 1-2 h per mm of brain tissue. A whole mouse brain requires 8-10 h for sufficient clearing. Remember the orientation of the sample before removing it from the chamber and be sure to replace it back into the chamber in the same orientation. 
		&#8203;NOTE: Figure 3A shows what a fully cleared brain will look like. The brain will look opaque and not clear at this stage due to the presence of dissolved SDS, but there should be little to no yellow tissue colored hue after lipid extraction.</li>
	</ol>
	</li>
</ol>
3. Preparing and mounting the cleared tissue
<ol>
	<li>After the sample has finished clearing and looks sufficiently clear, wash in PBS overnight at room temperature. Replace the PBS with fresh PBS as often as possible. This step is critical to remove residual SDS that can form precipitates in later steps.</li>
	<li>Following the final wash in PBS, wash the tissue for 5 min in deionized water at room temperature three times. The tissue will become opaque at this step and may expand.</li>
	<li>Incubate the tissue in the refractive index matching solution (see Table 1) for at least 4 h at room temperature. Figure 3B shows a piece of cleared tissue after incubation in refractive index matching solution.</li>
	<li>During the incubation of tissue in refractive index matching solution, construct a suitable housing chamber to image the sample if necessary.</li>
	<li>Constructing an imaging chamber for small tissue samples/slices
	<ol>
		<li>Using a glass slide as a base for mounting, lay down either rubber or plastic spacers and secure with super glue. If premade spacers are not available, use plastic rings made from conical tube cross sections.</li>
		<li>Ensure to secure these pieces to the glass slide without any holes (Figure 3C).</li>
		<li>Place the cleared tissue into the mounting chamber prefilled with refractive index matching solution.</li>
		<li>Securely mount the tissue by placing a glass coverslip on top and sealing it with nail polish.</li>
		<li>Image this tissue by adding a drop of refractive index matching mounting solution directly on top of the glass.</li>
	</ol>
	</li>
	<li>Large tissue imaging chamber
	<ol>
		<li>Construct this chamber if the tissue is larger than 5 mm thick (suitable for whole brains or hemispheres).</li>
		<li>Using a 10 cm glass dish with a high wall, place a 50 mL conical tube in the center, making sure that the diameter of conical is large enough to accept the barrel of the objective lens used.</li>
		<li>Make 3% agarose in water and pour it in the space between the glass dish and conical tube, allow to cool for 1 h (Figure 3D). This will form a ring of solid agarose (Figure 3E).</li>
		<li>Securely adhere the tissue to the bottom of the chamber using super glue and fill the chamber with refractive index matching solution. Apply glue to adhere the tissue on a region that will not be imaged to allow reclamation of the tissue from the dish without damaging the regions of interest. 
		NOTE: This preparation is time sensitive, as the refractive index media may start to polymerize unless preserved from air and stored at 4 &#176;C.</li>
	</ol>
	</li>
</ol>
4. Imaging cleared tissue samples
<ol>
	<li>Acquire the image using a confocal microscope fit with a 25x/0.95 NA objective with a 4 mm working distance.</li>
	<li>Turn on all the relevant imaging equipment. Place the sample on the stage and place a drop of refractive index matching solution onto the top of the mounting chamber.</li>
	<li>Carefully approach the immersion media with the objective and form a continuous column of media.</li>
	<li>Using epifluorescence, find an appropriate imaging field.</li>
	<li>Begin the image acquisition procedure by testing the appropriate settings.
	<ol>
		<li>Start by setting the resolution and scan speed settings using Figure 4A as a guide. If imaging using a confocal microscope, fully close the pinhole to obtain the smallest optical section and thus best z-resolution.</li>
		<li>Gradually increase the laser power/sensor gain until a suitable image is obtained with a high signal-to-noise ratio.</li>
		<li>If utilizing standard EGFP/tdTomato two-color imaging, set the light collection settings using Figure 4B as a guide.</li>
		<li>Set the z-stack parameters based on the observed start and end points of the tissue. Set the step size based on the desired z-resolution using Figure 4C as a guide. 
		NOTE: Smaller step sizes will yield a greater z-resolution but will also introduce more laser dwell time, potentially leading to sample bleaching.</li>
		<li>When satisfied with image acquisition settings, acquire the image.</li>
		<li>Ensure that the image has a high signal-to-noise ratio and shows distinct boundaries of structures (Figure 4D).</li>
	</ol>
	</li>
</ol>
5. Image processing and 3D quantification using microscopy analysis software
NOTE: Microscopic image analysis software packages are powerful tools for three-dimensional image visualization and processing. Many of these programs are perfectly suited for the handling of large datasets that are generated from imaging cleared tissue samples. The following steps and associated figures correspond to the Imaris software workflow.
<ol>
	<li>Open the image stack and import it into the selected analysis software.</li>
	<li>View the image in three-dimensional space and make any desired changes to the lookup tables using the display adjuster to better visualize the image. 
	NOTE: Figure 5A demonstrates the possible extreme imaging depth accessible through 2-photon microscopy paired with CLARITY tissue clearing. Figure 5B shows distinct dendrite processes as well as clearly visible spine morphologies from the z-stack presented in Figure 5A.</li>
	<li>To filter out any consistent background, click on the Image Processing button and select the background subtraction filter. 
	NOTE: Figure 5C shows the image with a consistent hazy background signal before processing. Figure 5D shows the image after the background subtraction filter has been applied.</li>
	<li>Observe the 3D image and become familiar with it by looking at it from multiple angles and zoom levels.</li>
	<li>Start the dendrite tracing by first selecting the Filament tracer tool.</li>
	<li>Click on Edit the Filament Manually, Skip Automatic Creation.</li>
	<li>Set the mode to auto path and check Auto-center and Auto-diameter corrections.</li>
	<li>Shift + right click on the cell body to set a starting point.</li>
	<li>Trace the neuron along the entire length of the dendrite; left click on the end of the dendrite to set the termination point to allow the software to automatically calculate the path in between the start and end points.</li>
	<li>Repeat this step for all the dendrites, and fully trace out the cell structure.</li>
	<li>Visualize the traced cell and confirm its accuracy. Make manual adjustments as needed.</li>
	<li>Select the Creation tab.</li>
	<li>Select the Recompute Dendrite Diameter option.</li>
	<li>Follow the wizard to completion for a more accurately traced dendrite.</li>
	<li>Select the Draw tab.</li>
	<li>Click on the Spine radio button to start drawing spines.</li>
	<li>Set the approximate spine diameter as needed, and use the measurement tool to get an accurate representation of spine diameters.</li>
	<li>Click on the center of the spine heads to add a new spine.</li>
	<li>Repeat this for all spines on the dendrite. 
	NOTE: It is important to observe the dendrite from all possible angles when manually adding spines.</li>
	<li>Check the newly added spines for accuracy and make changes as needed.</li>
	<li>Select the Creation tab.</li>
	<li>Select the Recompute Spine Diameter option to allow the software to determine the proper head and neck diameters, which are crucial for downstream data analysis.</li>
	<li>Follow the spine diameter creation wizard to completion.</li>
	<li>Observe the result of the computation and make any manual adjustments as needed. Figure 5E shows a fully traced neuron complete with spines.</li>
	<li>To create a classified list of spines, select the Tools tab. 
	NOTE: The MATLAB extension must be installed for this to work.
	<ol>
		<li>To install the MATLAB extension, open the preferences window.</li>
		<li>Select the Custom Tools option.</li>
		<li>Add the appropriate MATLAB runtime MCR.</li>
	</ol>
	</li>
	<li>Click on Classify Spines.</li>
	<li>Edit the desired parameters for spine classification.</li>
	<li>Click on Classify Spines on the MATLAB extension. Figure 5F shows the dendrite overlaid with color-coded classified spines.</li>
	<li>Select the Statistics tab.</li>
	<li>Configure the desired statistics to quantify the data by clicking on the Configure button in the bottom-left corner of this tab.</li>
	<li>Once the method of statistics representation has been chosen, export the data using the Export Statistics on Tab Display to File button located on the bottom right of this window.</li>
	<li>Graph the statistics using a preferred graphing method.</li>
</ol>

<ol>
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The authors have nothing to disclose at this time.

Before the advent of contemporary tissue clearing techniques, studying neuronal morphology consisted of time-intensive sectioning, imaging, and reconstruction of adjacent very thin sections. Using electrophoretic tissue clearing in combination with confocal imaging provides an unobstructed view of complete neuronal morphology. From intact dendritic trees, down to the smallest synaptic bouton, imaging and quantifying neuronal morphology has never been more feasible.
The preparation of cleared b...

Understanding the spatial organization, patterns of connectivity, and morphology of complex three-dimensional biological structures is essential for delineating the functions of specific cells and tissues. This is especially true in neuroscience, in which tremendous effort has been dedicated to building high-resolution neuroanatomical maps of the central nervous system1,2. Close examination of the neurons that comprise these maps yields varied morphologies, with connections and locations that reflect the function of these diverse sets of neurons3,4. Moreover, investigation of subcellular structures, especially dendritic spines, can inform the maturity of synapses, thereby reflecting developmental processes and neurological disease states5,6,7. Thus, approaches that improve imaging resolution and throughput are essential toward better understanding brain function at all scales.
Recent advances have expanded the molecular and genetic toolkit for marking and manipulating populations of neurons. The development of new fluorescent markers, combined with new methods of introducing these markers into neurons, allows for differential labeling of populations of interacting neurons within the same animal or brain sample8,9,10,11. Because light is scattered by opaque lipids, and given the high lipid content of brain tissue, imaging neuronal populations has primarily been limited to thin sections or has relied on advanced microscropy techniques (e.g., confocal, multi-photon, and light-sheet microscopy) to image deep structures. However, these efforts have been greatly bolstered by advances in tissue clearing techniques. Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging/immunostaining/in situ hybridization-compatible Tissue-hYdrogel (CLARITY) is one such technique, in which tissues of interest are infused with hydrogel monomers (acrylamide and bis-acrylamide) and then washed with detergents12. The hydrogel monomers hybridize to create a stable 3D hydrogel scaffold that is optically transparent and permeable to macromolecule labels. Nucleic acids and proteins are maintained within the hybridized matrix, whereas lipids are removed by the detergent washes (Figure 1). This results in a stable tissue that is rigid enough to maintain the original shape and orientation of cells and non-lipid molecules, while optically transparent enough to easily image deep structures at high resolution. This maintenance of tissue structure and orientation allows for the imaging of thick slices, thereby preserving cell-to-cell connections and spatial relationships. Moreover, because the location and availability of proteins and nucleic acids is maintained during the clearing process, cleared tissues are able to hold expression-based markers, as well as exogenous labels. Thus, CLARITY lends itself as a potent method for imaging large amounts of deep brain structures and the connections between these structures at high resolution.
The use of CLARITY greatly improves approaches to imaging neuronal populations. This technique is especially adept at generating large amounts of imaging data. CLARITY works well with multiple forms of protein-based fluorescence. This protocol utilizes a lentiviral-based approach to sparsely label cells with EGFP and tdTomato; however, transgenic reporter alleles expressing tdTomato or EGFP to label cells for reconstruction have been routinely used. It is important to choose a fluorophore that is both photo-stable and bright (e.g., EGFP or tdTomato). Additionally, using a strong promoter to express the fluorophore yields superior contrast and image quality. The drawbacks of this technique arise as properly analyzing this large amount of data can be both labor- and time-intensive. Specialized microscopes can help improve throughput and decrease workload. However, building, owning, and/or operating advanced microscopes are often cost-prohibitive for many laboratories. This work presents a high throughput, relatively rapid, and simple method of visualizing large amounts of neural tissue at high-resolution using CLARITY tissue clearing of large sections, combined with standard confocal microscopy. This protocol describes this approach through the following steps: 1) dissecting and preparing the neural tissue, 2) clearing the tissue, 3) mounting the tissue, 4) imaging the prepared slices, and 5) processing full slice images using microscopy visualization software reconstruction and analysis (Figure 2). These efforts result in high-resolution images that can be used to analyze populations of neurons, neuronal connection patterns, 3D dendritic morphology, dendritic spine abundance and morphology, and molecular expression patterns within intact brain tissue.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>15 mL Conical Tube</td>
			<td>Thermo Scientific</td>
			<td>339650</td>
			<td></td>
		</tr>
		<tr>
			<td>25 G x 1&#34; Needle</td>
			<td>BD</td>
			<td>305127</td>
			<td></td>
		</tr>
		<tr>
			<td>30% Acrylamide (No-Bis)</td>
			<td>National Diagnostics</td>
			<td>EC-810</td>
			<td></td>
		</tr>
		<tr>
			<td>50 mL Conical Tube</td>
			<td>Thermo Scientific</td>
			<td>339653</td>
			<td></td>
		</tr>
		<tr>
			<td>Electrophoretic Tissue Clearing Solution</td>
			<td>Logos</td>
			<td>C13001</td>
			<td></td>
		</tr>
		<tr>
			<td>Histodenz</td>
			<td>Sigma</td>
			<td>D2158-100G</td>
			<td></td>
		</tr>
		<tr>
			<td>Hydrogel Solution Kit</td>
			<td>Logos</td>
			<td>C1310X</td>
			<td></td>
		</tr>
		<tr>
			<td>Imaris</td>
			<td>Oxford Instruments</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde 16%</td>
			<td>EMS</td>
			<td>15710</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS, 1x, 500 mL, 6 bottles/case</td>
			<td>fisher</td>
			<td>MT21040CV</td>
			<td></td>
		</tr>
		<tr>
			<td>VA-044</td>
			<td>Wako</td>
			<td>925-41020</td>
			<td></td>
		</tr>
		<tr>
			<td>X-CLARITY Polymerization System</td>
			<td>Logos</td>
			<td>C20001</td>
			<td></td>
		</tr>
		<tr>
			<td>X-CLARITY Tissue Clearing System II</td>
			<td>Logos</td>
			<td>C30001</td>
			<td></td>
		</tr>
	</tbody>
</table>

imaging and quantification of intact neuronal dendrites via clarity tissue clearing

Brain activity, the electrochemical signals passed between neurons, is determined by the connectivity patterns of neuronal networks, and from the morphology of processes and substructures within these neurons. As such, much of what is known about brain function has arisen alongside developments in imaging technologies that allow further insight into how neurons are organized and connected in the brain. Improvements in tissue clearing have allowed for high-resolution imaging of thick brain slices, facilitating morphological reconstruction and analyses of neuronal substructures, such as dendritic arbors and spines. In tandem, advances in image processing software provide methods of quickly analyzing large imaging datasets. This work presents a relatively rapid method of processing, visualizing, and analyzing thick slices of labeled neural tissue at high-resolution using CLARITY tissue clearing, confocal microscopy, and image analysis. This protocol will facilitate efforts toward understanding the connectivity patterns and neuronal morphologies that characterize healthy brains, and the changes in these characteristics that arise in diseased brain states.

After image acquisition, the representative cell morphology was analyzed using embedded statistics and classifying scripts within the analysis software. The collected data (Figure 6A) reflects that neuron 2 has a larger dendritic structure with a higher density of spines. As a whole, the data suggests that neuron 2 has a more complex dendritic structure compared to neuron 1. To substantiate this result, standard Sholl analysis was performed, which affirms that neuron 2 is more dendritically ...

Watch this Scientific Journal Video about Imaging and Quantification of Intact Neuronal Dendrites via CLARITY Tissue Clearing at JoVE.com

Imaging and Quantification of Intact Neuronal Dendrites via CLARITY Tissue Clearing

Neuronal dendritic morphology often underlies function. Indeed, many disease processes that affect the development of neurons manifest with a morphological phenotype. This protocol describes a simple and powerful method for analyzing intact dendritic arbors and their associated spines.

Brain activity, the electrochemical signals passed between neurons, is determined by the connectivity patterns of neuronal networks, and from the ...

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3.4K Views.  Baylor College of Medicine. Neuronal dendritic morphology often underlies function. Indeed, many disease processes that affect the development of neurons manifest with a morphological phenotype. This protocol describes a simple and powerful method for analyzing intact dendritic arbors and their associated spines.

Video: Imaging and Quantification of Intact Neuronal Dendrites via CLARITY Tissue Clearing

Intact Histological Characterization of Brain-implanted Microdevices and Surrounding Tissue

Research into the design and utilization of brain-implanted microdevices, such as microelectrode arrays, aims to produce clinically relevant devices that interface chronically with surrounding brain tissue. Tissue surrounding these implants is thought to react to the presence of the devices over time, which includes the formation of an insulating "glial scar" around the devices. However, histological analysis of these tissue changes is typically performed after explanting the device, in a process that can disrupt the morphology of the tissue of interest.
Here we demonstrate a protocol in which cortical-implanted devices are collected intact in surrounding rodent brain tissue. We describe how, once perfused with fixative, brains are removed and sliced in such a way as to avoid explanting devices. We outline fluorescent antibody labeling and optical clearing methods useful for producing an informative, yet thick tissue section. Finally, we demonstrate the mounting and imaging of these tissue sections in order to investigate the biological interface around brain-implanted devices.

Here we present a histological method for capturing, labeling, optically clearing, and imaging the intact brain tissue interface around chronically implanted microdevices in rodent brain tissue. Results from the techniques comprising this method are useful for understanding the impact of various penetrating brain-implants on their surrounding tissue.

Research  into the design and utilization of brain-implanted microdevices, such as  microelectrode arrays, aims to produce clinically relevant devices ...

Imaging Serotonergic Fibers in the Mouse Spinal Cord Using the CLARITY/CUBIC Technique

Long descending fibers to the spinal cord are essential for locomotion, pain perception, and other behaviors. The fiber termination pattern in the spinal cord of the majority of these fiber systems have not been thoroughly investigated in any species. Serotonergic fibers, which project to the spinal cord, have been studied in rats and opossums on histological sections and their functional significance has been deduced based on their fiber termination pattern in the spinal cord. With the development of CLARITY and CUBIC techniques, it is possible to investigate this fiber system and its distribution in the spinal cord, which is likely to reveal previously unknown features of serotonergic supraspinal pathways. Here, we provide a detailed protocol for imaging the serotonergic fibers in the mouse spinal cord using the combined CLARITY and CUBIC techniques. The method involves perfusion of a mouse with a hydrogel solution and clarification of the tissue with a combination of clearing reagents. Spinal cord tissue was cleared in just under two weeks, and the subsequent immunofluorescent staining against serotonin was completed in less than ten days. With a multi-photon fluorescent microscope, the tissue was scanned and a 3D image was reconstructed using Osirix software.

Supraspinal projections are important for pain perception and other behaviors, and serotonergic fibers are one of these fiber systems. The present study focused on the application of the combined CLARITY/CUBIC protocol to the mouse spinal cord in order to investigate the termination of these serotonergic fibers.

Long descending fibers to the spinal cord are essential for locomotion, pain perception, and other behaviors. The fiber termination pattern in the spinal ...

Optical Clearing of the Mouse Central Nervous System Using Passive CLARITY

Traditionally, tissue visualization has required that the tissue of interest be serially sectioned and imaged, subjecting each tissue section to unique non-linear deformations, dramatically hampering one's ability to evaluate cellular morphology, distribution and connectivity in the central nervous system (CNS). However, optical clearing techniques are changing the way tissues are visualized. These approaches permit one to probe deeply into intact organ preparations, providing tremendous insight into the structural organization of tissues in health and disease. Techniques such as Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tissue-hYdrogel (CLARITY) achieve this goal by providing a matrix that binds important biomolecules while permitting light-scattering lipids to freely diffuse out. Lipid removal, followed by refractive index matching, renders the tissue transparent and readily imaged in 3 dimensions (3D). Nevertheless, the electrophoretic tissue clearing (ETC) used in the original CLARITY protocol can be challenging to implement successfully and the use of a proprietary refraction index matching solution makes it expensive to use the technique routinely. This report demonstrates the implementation of a simple and inexpensive optical clearing protocol that combines passive CLARITY for improved tissue integrity and 2,2&#8242;-thiodiethanol (TDE), a previously described refractive index matching solution.

Optical clearing techniques are revolutionizing the way tissues are visualized. In this report we describe modifications of the original Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tissue-hYdrogel (CLARITY) protocol that yields more consistent and less expensive results.

Traditionally, tissue visualization has required that the tissue of interest be serially sectioned and imaged, subjecting each tissue section to unique ...

Prolonged Incubation of Acute Neuronal Tissue for Electrophysiology and Calcium-imaging

Acute neuronal tissue preparations, brain slices and retinal wholemount, can usually only be maintained for 6 - 8 h following dissection. This limits the experimental time, and increases the number of animals that are utilized per study. This limitation specifically impacts protocols such as calcium imaging that require prolonged pre-incubation with bath-applied dyes. Exponential bacterial growth within 3 - 4 h after slicing is tightly correlated with a decrease in tissue health. This study describes a method for&#160;limiting the proliferation of bacteria in acute preparations to maintain viable neuronal tissue for prolonged periods of time (&#62;24 h) without the need for antibiotics, sterile procedures, or tissue culture media containing growth factors. By cycling the extracellular fluid through UV irradiation and keeping the tissue in a custom holding chamber at 15 - 16 &#176;C, the tissue shows no difference in electrophysiological properties, or calcium signaling through intracellular calcium dyes at &#62;24 h postdissection. These methods will not only extend experimental time for those using acute neuronal tissue, but will reduce the number of animals required to complete experimental goals, and will set a gold standard for acute neuronal tissue incubation.

Once removed from the body, neuronal tissue is greatly affected by environmental conditions, leading to eventual degradation of the tissue after 6 - 8 h. Using a unique incubation method, which closely monitors and regulates the extracellular environment of the tissue, tissue viability can be significantly extended for &#62;24 h.

Acute neuronal tissue preparations, brain slices and retinal wholemount, can usually only be maintained for 6 - 8 h following dissection. This limits the ...

Two-photon Calcium Imaging in Neuronal Dendrites in Brain Slices

Calcium (Ca2+) imaging is a powerful tool to investigate the spatiotemporal dynamics of intracellular Ca2+ signals in neuronal dendrites. Ca2+ fluctuations can occur through a variety of membrane and intracellular mechanisms and play a crucial role in the induction of synaptic plasticity and regulation of dendritic excitability. Hence, the ability to record different types of Ca2+ signals in dendritic branches is valuable for groups studying how dendrites integrate information. The advent of two-photon microscopy has made such studies significantly easier by solving the problems inherent to imaging in live tissue, such as light scattering and photodamage. Moreover, through combination of conventional electrophysiological techniques with two-photon Ca2+ imaging, it is possible to investigate local Ca2+ fluctuations in neuronal dendrites in parallel with recordings of synaptic activity in soma. Here, we describe how to use this method to study the dynamics of local Ca2+ transients (CaTs) in dendrites of GABAergic inhibitory interneurons. The method can be also applied to studying dendritic Ca2+ signaling in different neuronal types in acute brain slices.

We present a method combining whole-cell patch-clamp recordings and two-photon imaging to record Ca2+ transients in neuronal dendrites in acute brain slices.

Calcium (Ca2+) imaging is a powerful tool to investigate the spatiotemporal dynamics of intracellular Ca2+ signals in neuronal dendrites. Ca2+ ...

Novel Passive Clearing Methods for the Rapid Production of Optical Transparency in Whole CNS Tissue

Since the development of CLARITY, a bioelectrochemical clearing technique that allows for three-dimensional phenotype mapping within transparent tissues, a multitude of novel clearing methodologies including CUBIC (clear, unobstructed brain imaging cocktails and computational analysis), SWITCH (system-wide control of interaction time and kinetics of chemicals), MAP (magnified analysis of the proteome), and PACT (passive clarity technique), have been established to further expand the existing toolkit for the microscopic analysis of biological tissues. The present study aims to improve upon and optimize the original PACT procedure for an array of intact rodent tissues, including the whole central nervous system (CNS), kidneys, spleen, and whole mouse embryos. Termed psPACT (process-separate PACT) and mPACT (modified PACT), these novel techniques provide highly efficacious means of mapping cell circuitry and visualizing subcellular structures in intact normal and pathological tissues. In the following protocol, we provide a detailed, step-by-step outline on how to achieve maximal tissue clearance with minimal invasion of their structural integrity via psPACT and mPACT.

Here, we present two novel methodologies, psPACT and mPACT, for achieving maximal optical transparency and subsequent microscopic analysis of tissue vasculature in the intact rodent whole CNS.

Since the development of CLARITY, a bioelectrochemical clearing technique that allows for three-dimensional phenotype mapping within transparent tissues, ...

Monitoring Neuronal Survival via Longitudinal Fluorescence Microscopy

Standard cytotoxicity assays, which require the collection of lysates or fixed cells at multiple time points, have limited sensitivity and capacity to assess factors that influence neuronal fate. These assays require the observation of separate populations of cells at discrete time points. As a result, individual cells cannot be followed prospectively over time, severely limiting the ability to discriminate whether subcellular events, such as puncta formation or protein mislocalization, are pathogenic drivers of disease, homeostatic responses, or merely coincidental phenomena. Single-cell longitudinal microscopy overcomes these limitations, allowing the researcher to determine differences in survival between populations and draw causal relationships with enhanced sensitivity. This video guide will outline a representative workflow for experiments measuring single-cell survival of rat primary cortical neurons expressing a fluorescent protein marker. The viewer will learn how to achieve high-efficiency transfections, collect and process images enabling the prospective tracking of individual cells, and compare the relative survival of neuronal populations using Cox proportional hazards analysis.

Here, we present a protocol to monitor survival on a single-cell basis and identify variables that significantly predict cell death.

Standard cytotoxicity assays, which require the collection of lysates or fixed cells at multiple time points, have limited sensitivity and capacity to ...

A Hydrophobic Tissue Clearing Method for Rat Brain Tissue

Hydrophobic tissue clearing methods are easily adjustable, fast, and low-cost procedures that allows for the study of a molecule of interest in unaltered tissue samples. Traditional immunolabeling procedures require cutting the sample into thin sections, which restricts the ability to label and examine intact structures. However, if brain tissue can remain intact during processing, structures and circuits can remain intact for the analysis. Previously established clearing methods take significant time to completely clear the tissue, and the harsh chemicals can often damage sensitive antibodies. The iDISCO method quickly and completely clears tissue, is compatible with many antibodies, and requires no special lab equipment. This technique was initially validated for the use in mice tissue, but the current protocol adapts this method to image hemispheres of control and transgenic rat brains. In addition to this, the present protocol also makes several adjustments to preexisting protocol to provide clearer images with less background staining. Antibodies for Iba-1 and tyrosine hydroxylase were validated in the HIV-1 transgenic rat and in F344/N control rats using the present hydrophobic tissue clearing method. The brain is an interwoven network, where structures work together more often than separately of one another. Analyzing the brain as a whole system as opposed to a combination of individual pieces is the greatest benefit of this whole brain clearing method.

Here we present a hydrophobic tissue clearing method that allows for the viewing of target molecules as part of intact brain structures. This technique has now been validated for F344/N control and HIV-1 transgenic rats of both sexes.

Hydrophobic tissue clearing methods are easily adjustable, fast, and low-cost procedures that allows for the study of a molecule of interest in unaltered ...

Preparing Adult Drosophila melanogaster for Whole Brain Imaging during Behavior and Stimuli Responses

We present a method developed specifically to image the whole Drosophila brain during ongoing behavior such as walking. Head fixation and dissection are optimized to minimize their impact on behavior. This is first achieved by using a holder that minimizes movement hindrances. The back of the fly&#8217;s head is glued to this holder at an angle that allows optical access to the whole brain while retaining the fly&#8217;s ability to walk, groom, smell, taste and see. The back of the head is dissected to remove tissues in the optical path and muscles responsible for head movement artefacts. The fly brain can subsequently be imaged to record brain activity, for instance using calcium or voltage indicators, during specific behaviors such as walking or grooming, and in response to different stimuli. Once the challenging dissection, which requires considerable practice, has been mastered, this technique allows to record rich data sets relating whole brain activity to behavior and stimulus responses.

Preparing Adult Drosophila melanogaster for Whole Brain Imaging during Behavior and Stimuli Responses

We present a method specifically tailored to image the whole brain of adult Drosophila during behavior and in response to stimuli. The head is positioned to allow optical access to the whole brain, while the fly can move its legs and the antennae, the tip of the proboscis, and the eyes can receive sensory stimuli.

We present a method developed specifically to image the whole Drosophila brain during ongoing behavior such as walking. Head fixation and dissection are ...

Immunostaining of Whole-Mount Retinas with the CLARITY Tissue Clearing Method

The tissue hydrogel delipidation method (CLARITY), originally developed by the Deisseroth laboratory, has been modified and widely used for immunostaining and imaging of thick brain samples. However, this advanced technology has not yet been used for whole-mount retinas. Although the retina is partially transparent, its thickness of approximately 200 &#181;m (in mice) still limits the penetration of antibodies into the deep tissue as well as reducing light penetration for high-resolution imaging. Here, we adapted the CLARITY method for whole-mount mouse retinas by polymerizing them with an acrylamide monomer to form a nanoporous hydrogel and then clearing them in sodium dodecyl sulfate to minimize protein loss and avoid tissue damage. CLARITY-processed retinas were immunostained with antibodies for retinal neurons, glial cells, and synaptic proteins, mounted in a refractive index matching solution, and imaged. Our data demonstrate that CLARITY&#160;can improve the quality of standard immunohistochemical staining and imaging for retinal neurons and glial cells in whole-mount preparation. For instance, 3D resolution of fine axon-like and dendritic structures of dopaminergic amacrine cells were much improved by CLARITY. Compared to non-processed whole-mount retinas, CLARITY can reveal immunostaining for synaptic proteins such as postsynaptic density protein 95. Our results show that CLARITY renders the retina more optically transparent after the removal of lipids and preserves fine structures of retinal neurons and their proteins, which can be routinely used for obtaining high-resolution imaging of retinal neurons and their subcellular structures in whole-mount preparation.

Here we present a protocol to adapt the CLARITY method of the brain tissues for whole-mount retinas to improve the quality of standard immunohistochemical staining and high-resolution imaging of retinal neurons and their subcellular structures.

The tissue hydrogel delipidation method (CLARITY), originally developed by the Deisseroth laboratory, has been modified and widely used for immunostaining ...