The authors thank Yoko Ishida (Juntendo University) for AAV vector production and Kisara Hoshino (Juntendo University) for technical assistance. This study was supported by JSPS KAKENHI (JP20K07231 to K.Y.; JP21H03529 to T.F.; JP20K07743 to M.K.; JP21H02592 to H.H.) and Scientific Research on Innovative Area &#8220;Resonance Bio&#8221; (JP18H04743 to H.H.). This study was also supported by the Japan Agency for Medical Research and Development (AMED) (JP21dm0207112 to T.F. and H.H.), Moonshot R&#38;D from the Japan Science and Technology Agency (JST) (JPMJMS2024 to H.H.), Fusion Oriented Research for disruptive Science and Technology (FOREST) from JST (JPMJFR204D to H.H.), Grants-in-Aid from the Research Institute for Diseases of Old Age at the Juntendo University School of Medicine (X2016 to K.Y.; X2001 to H.H.), and the Private School Branding Project.

All the experiments were approved by the Institutional Animal Care and Use Committees of Juntendo University (Approval No. 2021245, 2021246) and performed in accordance with Fundamental Guidelines for Proper Conduct of Animal Experiments by the Science Council of Japan (2006). Here, male C57BL/6J mice injected with AAV vector carrying enhanced green fluorescent protein (EGFP) gene and parvalbumin (PV)/myristoylation-EGFP-low-density lipoprotein receptor C-terminal bacterial artificial chromosome (BAC) transgenic mice (PV-FGL mice)9 were used. PV-FGL mice were maintained in C57BL/6J background. No sex-based differences were found with regard to this study.
1. Tissue preparation
<ol>
	<li>Perfusion fixation 
	NOTE: Perform steps 1.1.1 through 1.1.3 in a fume hood to limit the exposure to paraformaldehyde (PFA).
	<ol>
		<li>Anesthetize adult male mice (8&#8211;16 weeks old) by an intraperitoneal injection of overdose of sodium pentobarbital (200 mg/kg). Confirm adequacy of anesthesia by the absence of toe-pinch withdrawal and eye-blink reflexes.</li>
		<li>Open the thoracic cavity and cut the right atrial appendage with surgical scissors. Perfuse the mice with 20 mL of ice-cold phosphate buffer saline (PBS) using a 23 G needle attached to a 20 mL syringe, followed by perfusion of 20 mL of ice-cold 4% PFA in 0.1 M phosphate buffer (PB) using another 20 mL syringe. 
		CAUTION: PFA is toxic and teratogenic. Avoid inhalation or contact with skin, eyes, and mucous membrane.</li>
		<li>Remove brain tissues from the skull with tweezers. Transfer the brain tissues to a 15 mL tube containing 4% PFA in 0.1 M PB, protect the samples from light, and gently rock overnight at 4 &#176;C on a shaker at 50&#8211;100 rpm.</li>
		<li>NOTE: The harvested brain tissues can be stored for several weeks in 0.02% sodium azide (NaN3) in PBS at 4 &#176;C. 
		CAUTION: NaN3 is toxic. Avoid inhalation or contact with skin, eyes, and mucous membrane. Handle it inside a fume hood.</li>
	</ol>
	</li>
	<li>Brain slice preparation
	<ol>
		<li>Prepare 4% agar in PBS by adding 2 g of agar to 50 mL of PBS. Microwave the mixture until the agar is fully dissolved. Let the solution cool to 40-45 &#176;C. 
		NOTE: The viscous property provided by agaropectin, a major component of agar, improves ease of cutting of tissue slices.</li>
		<li>Add 10 mL of the agar solution to a 6-well culture plate. Submerge the brain tissue in the agar solution using forceps. Let the agar solidify on ice.</li>
		<li>Remove the embedded brain tissue from the well and trim the agar with a razor blade. Secure the agar block onto the bottom of the vibratome bath with superglue and pour 0.1 M PB in the buffer tray.</li>
		<li>Clean another razor blade using a lint-free tissue paper soaked in ethanol and attach the blade to the blade holder of the vibrating tissue slicer.</li>
		<li>Set the sectioning speed to 0.14 mm/s with 1.4 mm amplitude and the frequency to 75&#8211;77 Hz. Cut the brain tissue into 1-mm-thick slices and collect the slices in a 6-well cell culture plate containing PBS. 
		NOTE: The brain slices can be stored for several weeks in 0.02% NaN3 in PBS at 4 &#176;C.</li>
	</ol>
	</li>
</ol>
2. Tissue clarification
NOTE: The compositions of ScaleS solutions used are listed in Table 1. Samples should be protected from light by covering with a foil. The clearing steps is shown in Figure 1A.
<ol>
	<li>Add 8 mL of ScaleS0 solution to one well of a 6-well cell culture plate and add 8 mL of ScaleS4 solution to another well of the plate and pre-warm to 37 &#176;C in an incubator.</li>
	<li>Transfer the brain slices to the pre-warmed ScaleS0 solution with a spatula and incubate for 2 h at 37 &#176;C in a shaking incubator at 90 rpm.</li>
	<li>Transfer the permeabilized brain slices in 8 mL of PBS(&#8211;) in a 6-well cell culture plate with a spatula and wash for 15 min by keeping in an orbital shaker at 40&#8211;60 rpm. Repeat this twice.</li>
	<li>Transfer the brain slices in the pre-warmed 8 mL of ScaleS4 solution with a spatula and clear them by incubating in a shaking incubator at 90 rpm for 8&#8211;12 h at 37 &#176;C. A cleared brain slices can be seen in Figure 1.</li>
</ol>
3. Brain slice mounting
NOTE: A customizable imaging chamber is used for reliable mounting of cleared brain slices (Figure 2)6. The chamber consists of the chamber frame and bottom coverslip. The microscope stage adaptors are also designed to mount the imaging chamber on microscope stages directly (Figure 2A,B). The chamber frame and microscope stage adaptors can be 3D-printed using in-house or outsourced 3D-printing services. 3D computer-aided design (CAD) data of the imaging chamber are provided in Furuta et al. 20226.
<ol>
	<li>For chamber preparation, attach the chamber frame to a coverslip using a pressure-sensitive adhesive.</li>
	<li>Prepare 1.5% agarose in ScaleS4D25(0) solution (ScaleS4 gel) by adding 1.5 g of agarose to 100 mL of the solution in a bottle. Mix the solution well by stirring and microwave the solution until the agarose is fully dissolved. Once done, allow the solution to cool to 37 &#176;C.</li>
	<li>Mount the cleared brain slice onto the bottom coverslip of the imaging chamber with a spatula. Wipe away the excess solution from the cleared slice using a clean lint-free tissue paper.</li>
	<li>Add the ScaleS4 gel on the brain slice using a micropipette to fill the imaging chamber. Place another coverslip on top with forceps and place a piece of lint-free tissue paper and a glass slide on the coverslip in this order.</li>
	<li>Transfer the imaging chamber to a refrigerator at 4 &#176;C. Place metal weights on the glass slide and leave them for 30 min.</li>
	<li>Remove the metal weights, glass slide, lint-free tissue paper, and coverslip from the imaging chamber, and wipe the excess gel away (Figure 2A,B).</li>
	<li>Place the imaging chamber in a 60 mm glass Petri dish and attach the rim of the imaging chamber to the dish with a putty-like pressure sensitive adhesive. Attach the chamber at multiple points to the Petri dish.</li>
	<li>Pour ScaleS4 solution in the dish and shake gently for 1 h at 20-25 &#176;C on an orbital shaker at 40-60 rpm. Substitute with fresh solution and remove air bubbles on the gel surface by gently scraping the surface using a 200 &#181;L pipette tip. Mount the immersed imaging chamber on a microscope stage (Figure 2C).</li>
</ol>
4. CLSM imaging
<ol>
	<li>Acquire images using a CLSM equipped with a multi-immersion objective lens of a long working distance (WD) (16x/0.60 numerical aperture [NA], WD = 2.5 mm). 
	NOTE: High NA objective lenses can provide high diffraction-limited resolution.</li>
	<li>Turn on all the relevant imaging equipment (workstation, microscope, scanner, lasers, and mercury lamp) and launch a CLSM imaging software.</li>
	<li>Set the correction collar of the multi-immersion objective lens to 1.47. ScaleS4 solution has a refractive index (RI) of around 1.475,7. RI mismatch-induced aberrations can disturb the image formation (Figure 3).</li>
	<li>Immerse the objective lens in the solution, and let it approach the slice slowly. Remove any air bubbles trapped on the tip of the objective lens. Find regions of interest (ROIs) in the cleared tissues using epifluorescence.</li>
	<li>Set image acquisition parameters by testing appropriate settings.
	<ol>
		<li>Determine the bit depth for image acquisition. The data size of the image increases with the bit depth.</li>
		<li>Set the&#160;detection wavelength. Adjust the appropriate gate for the detector according to the emission spectrum. Ensure that the detection wavelength does not cover any laser lines.</li>
		<li>Set the xy resolution. Larger formats provide better xy resolutions, but it takes longer time to collect the images.</li>
		<li>Set the scan speed. A slower scan speed provides a high signal-to-noise ratio. However, it also increases the pixel dwell time and the risk of photobleaching. Choose accordingly.</li>
		<li>Adjust the pinhole size. The pinhole size controls optical section thickness. A smaller pinhole size creates a thinner optical section, and thus better z resolution, but reduces fluorescence signal. Making the pinhole size larger provides a thicker optical section with stronger fluorescence signal.</li>
		<li>Set the laser power, the detector/amplifier gain, and offset. Gradually increase the laser power and detector/amplifier gain until a suitable image is obtained. A high laser power carries the risk of photobleaching. Adjust the offset (contrast) appropriately to obtain a high signal-to-noise ratio.</li>
		<li>Determine the tilling area needed based on the size of ROI. Ensure that the entire length and width of the ROI is captured.</li>
		<li>Navigate the cleared tissues in all planes, and set the start and end points of the stack. Set the z-step size according to the desired z-resolution.</li>
	</ol>
	</li>
	<li>Collect images when satisfied with the image acquisition settings, and record captured images. Process the images using an image analysis software.</li>
</ol>

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The authors have nothing to disclose.

Critical steps within the protocol 
There are a few critical steps in the protocol that should be conducted with utmost caution to obtain meaningful results. Uniform fixation of samples is imperative for 3D imaging within large-scale tissues. The objective lens, sample, and immersion fluid should have matching RI. RI-mismatch among them will lead to highly disturbed imaging of EGFP-expressing cells within the cleared brain slices (Figure 3). The correction collar adjustment of the o...

Tissue clearing methods have improved depth-independent imaging of biological and clinical samples with light microscopy, allowing for extraction of structural information on intact tissues1,2. Optical clearing techniques could also potentially speed up, and reduce the cost for histological analysis. Currently, three major clearing approaches are available: hydrophilic, hydrophobic, and hydrogel-based methods1,2. Hydrophilic approaches surpass in preserving fluorescence signals and tissue integrity and are less toxic compared to the other two approaches3,4.
A hydrophilic clearing method, ScaleS, holds a distinctive position with its preservation of structural and molecular integrity as well as potent clearing capability (clearing-preservation spectrum)5. In a previous study, we developed a rapid and isometric clearing protocol, ScaleSF, for tissue slices (~1-mm thickness) by modifying the clearing procedure of ScaleS6. This clearing protocol requires sequential incubations of brain slices in three solutions for 10.5-14.5 h. The method is featured with a high clearing-preservation spectrum, which is compatible even with electron microscopy (EM) analysis (Supplementary Figure 1), allowing for multi-scale high-resolution three dimensional (3D) imaging with accurate signal reconstruction6. Thus, ScaleSF should be effective especially in the brain, where neuronal cells elaborate exuberant processes of tremendous length, and arrange specialized fine subcellular structures for transmitting and receiving information. Extracting structural information with scales from circuit to subcellular levels on neuronal cells is quite useful toward better understanding of brain functions.
Here, we provide a detailed protocol to visualize neuronal structures with scales from the mesoscopic/circuit to microscopic/subcellular level using ScaleSF. The protocol includes tissue preparation, tissue clarification, handling of cleared tissues, and confocal laser scanning microscopy (CLSM) imaging of cleared tissues. Our protocol focuses on interrogating neuronal structures from circuit to subcellular component scales. For a detailed procedure for preparation of the solutions and stereotaxic injection of adeno-associated virus (AAV) vectors into mouse brains, refer to Miyawaki et al. 20167 and Okamoto et al. 20218, respectively.

<table><tbody><tr><td>16x multi-immersion objective lens</td><td>Leica Microsystems</td><td>HC FLUOTAR 16x/0.60 IMM CORR VISIR</td><td></td></tr><tr><td>Agar</td><td>Nacalai Tesque</td><td>01028-85</td><td></td></tr><tr><td>Agarose</td><td>TaKaRa Bio</td><td>L03</td><td></td></tr><tr><td>Dimethyl sulfoxide</td><td>Nacalai Tesque</td><td>13407-45</td><td></td></tr><tr><td>D-Sorbitol</td><td>Nacalai Tesque</td><td>06286-55</td><td></td></tr><tr><td>&amp;gamma;-cyclodextrin</td><td>Wako Pure Chemical Industries</td><td>037-10643</td><td></td></tr><tr><td>Glycerol</td><td>Sigma-Aldrich</td><td>G9012</td><td></td></tr><tr><td>Huygens Essential</td><td>Scientific Volume Imaging</td><td>ver. 18.10.0p8/21.10.1p0 64b</td><td></td></tr><tr><td>Imaris</td><td>Bitplane</td><td>ver. 9.0.0</td><td></td></tr><tr><td>Leica Application Suite X</td><td>Leica Microsystems</td><td>LAS X, ver. 3.5.5.19976</td><td></td></tr><tr><td>Methyl-&amp;beta;-cyclodextrin</td><td>Tokyo Chemical Industry</td><td>M1356</td><td></td></tr><tr><td>Paraformaldehyde</td><td>Merck Millipore</td><td>1.04005.1000</td><td></td></tr><tr><td>Phosphate Buffered Saline (10x; pH 7.4)</td><td>Nacalai Tesque</td><td>27575-31</td><td>10x PBS(&amp;ndash;)</td></tr><tr><td>Sodium azide</td><td>Nacalai Tesque</td><td>31233-55</td><td></td></tr><tr><td>Sodium pentobarbital</td><td>Kyoritsu Seiyaku</td><td>N/A</td><td></td></tr><tr><td>TCS SP8</td><td>Leica Microsystems</td><td>N/A</td><td></td></tr><tr><td>Triton X-100</td><td>Nacalai Tesque</td><td>35501-15</td><td></td></tr><tr><td>Urea</td><td>Nacalai Tesque</td><td>35940-65</td><td></td></tr><tr><td>Vibrating tissue slicer</td><td>Dosaka EM</td><td>PRO7N</td><td></td></tr></tbody></table>

a tissue clearing method for neuronal imaging from mesoscopic to microscopic scales

A detailed protocol is provided here to visualize neuronal structures from mesoscopic to microscopic levels in brain tissues. Neuronal structures ranging from neural circuits to subcellular neuronal structures are visualized in mouse brain slices optically cleared with ScaleSF. This clearing method is a modified version of ScaleS and is a hydrophilic tissue clearing method for tissue slices that achieves potent clearing capability as well as a high-level of preservation of fluorescence signals and structural integrity. A customizable three dimensional (3D)-printed imaging chamber is designed for reliable mounting of cleared brain tissues. Mouse brains injected with an adeno-associated virus vector carrying enhanced green fluorescent protein gene were fixed with 4% paraformaldehyde and cut into slices of 1-mm thickness with a vibrating tissue slicer. The brain slices were cleared by following the clearing protocol, which include sequential incubations in three solutions, namely, ScaleS0 solution, phosphate buffer saline (&#8211;), and ScaleS4 solution, for a total of 10.5&#8211;14.5 h. The cleared brain slices were mounted on the imaging chamber and embedded in 1.5% agarose gel dissolved in ScaleS4D25(0) solution. The 3D image acquisition of the slices was carried out using a confocal laser scanning microscope equipped with a multi-immersion objective lens of a long working distance. Beginning with mesoscopic neuronal imaging, we succeeded in visualizing fine subcellular neuronal structures, such as dendritic spines and axonal boutons, in the optically cleared brain slices. This protocol would facilitate understanding of neuronal structures from circuit to subcellular component scales.

Optical clearing of a mouse brain slice of 1-mm thickness was achieved using this protocol. Figure 1B represents transmission images of a mouse brain slice before and after the clearing treatment. The tissue clearing method rendered a 1-mm-thick mouse brain slice transparent. A slight expansion in final sizes of brain slices was found after the incubation in the clearing solution for 12 h (linear expansion: 102.5% &#177; 1.3%).&#160;The preservation of fluorescence and structural integrity of the tissues...

Watch this Scientific Journal Video about A Tissue Clearing Method for Neuronal Imaging from Mesoscopic to Microscopic Scales at JoVE.com

A Tissue Clearing Method for Neuronal Imaging from Mesoscopic to Microscopic Scales

The protocol provides a detailed method of neuronal imaging in brain slice using a tissue clearing method, ScaleSF. The protocol includes brain tissue preparation, tissue clarification, handling of cleared slices and confocal laser scanning microscopy imaging of neuronal structures from mesoscopic to microscopic levels.

A detailed protocol is provided here to visualize neuronal structures from mesoscopic to microscopic levels in brain tissues. Neuronal structures ranging ...

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Neuroscience

2.8K Views.  Juntendo University Graduate School of Medicine. Ons protocol zou het onderzoek van neuronale structuren van circuit- tot componentschalen vergemakkelijken, wat essentieel is voor een beter begrip van hersenfuncties. Onze clearingtechniek, ScaleSF, oefent krachtige clearingcapaciteit uit, evenals een hoog niveau van weefselbehoud dat nodig is voor gelijktijdige visualisatie van zowel grootschalige als kleinschalige structuren. Ons protocol is vooral effectief in de hersenen, waar neuronale cellen uitgebreide processen vertonen, enorme verbi...

Video: A Tissue Clearing Method for Neuronal Imaging from Mesoscopic to Microscopic Scales

A Rapid Approach to High-Resolution Fluorescence Imaging in Semi-Thick Brain Slices

A fundamental goal to both basic and clinical neuroscience is to better understand the identities, molecular makeup, and patterns of connectivity that are characteristic to neurons in both normal and diseased brain. Towards this, a great deal of effort has been placed on building high-resolution neuroanatomical maps1-3. With the expansion of molecular genetics and advances in light microscopy has come the ability to query not only neuronal morphologies, but also the molecular and cellular makeup of individual neurons and their associated networks4. Major advances in the ability to mark and manipulate neurons through transgenic and gene targeting technologies in the rodent now allow investigators to 'program' neuronal subsets at will5-6. Arguably, one of the most influential contributions to contemporary neuroscience has been the discovery and cloning of genes encoding fluorescent proteins (FPs) in marine invertebrates7-8, alongside their subsequent engineering to yield an ever-expanding toolbox of vital reporters9. Exploiting cell type-specific promoter activity to drive targeted FP expression in discrete neuronal populations now affords neuroanatomical investigation with genetic precision.
Engineering FP expression in neurons has vastly improved our understanding of brain structure and function. However, imaging individual neurons and their associated networks in deep brain tissues, or in three dimensions, has remained a challenge. Due to high lipid content, nervous tissue is rather opaque and exhibits auto fluorescence. These inherent biophysical properties make it difficult to visualize and image fluorescently labelled neurons at high resolution using standard epifluorescent or confocal microscopy beyond depths of tens of microns. To circumvent this challenge investigators often employ serial thin-section imaging and reconstruction methods10, or 2-photon laser scanning microscopy11. Current drawbacks to these approaches are the associated labor-intensive tissue preparation, or cost-prohibitive instrumentation respectively.
Here, we present a relatively rapid and simple method to visualize fluorescently labelled cells in fixed semi-thick mouse brain slices by optical clearing and imaging. In the attached protocol we describe the methods of: 1) fixing brain tissue in situ via intracardial perfusion, 2) dissection and removal of whole brain, 3) stationary brain embedding in agarose, 4) precision semi-thick slice preparation using new vibratome instrumentation, 5) clearing brain tissue through a glycerol gradient, and 6) mounting on glass slides for light microscopy and z-stack reconstruction (Figure 1).
For preparing brain slices we implemented a relatively new piece of instrumentation called the 'Compresstome' VF-200 (http://www.precisionary.com/products_vf200.html). This instrument is a semi-automated microtome equipped with a motorized advance and blade vibration system with features similar in function to other vibratomes. Unlike other vibratomes, the tissue to be sliced is mounted in an agarose plug within a stainless steel cylinder. The tissue is extruded at desired thicknesses from the cylinder, and cut by the forward advancing vibrating blade. The agarose plug/cylinder system allows for reproducible tissue mounting, alignment, and precision cutting. In our hands, the 'Compresstome' yields high quality tissue slices for electrophysiology, immunohistochemistry, and direct fixed-tissue mounting and imaging. Combined with optical clearing, here we demonstrate the preparation of semi-thick fixed brain slices for high-resolution fluorescent imaging.

Here we describe a rapid and simple method to image fluorescently labeled cells in semi-thick brain slices. By fixing, slicing, and optically clearing brain tissue we describe how standard epifluorescent or confocal imaging can be used to visualize individual cells and neuronal networks within intact nervous tissue.

A fundamental goal to both basic and clinical neuroscience is to better understand the identities, molecular makeup, and patterns of connectivity that are ...

Imaging Cleared Intact Biological Systems at a Cellular Level by 3DISCO

Tissue clearing and subsequent imaging of transparent organs is a powerful method to analyze fluorescently labeled cells and molecules in 3D, in intact organs. Unlike traditional histological methods, where the tissue of interest is sectioned for fluorescent imaging, 3D imaging of cleared tissue allows examination of labeled cells and molecules in the entire specimen. To this end, optically opaque tissues should be rendered transparent by matching the refractory indices throughout the tissue. Subsequently, the tissue can be imaged at once using laser-scanning microscopes to obtain a complete high-resolution 3D image of the specimen. A growing list of tissue clearing protocols including 3DISCO, CLARITY, Sca/e, ClearT2, and SeeDB provide new ways for researchers to image their tissue of interest as a whole. Among them, 3DISCO is a highly reproducible and straightforward method, which can clear different types of tissues and can be utilized with various microscopy techniques. This protocol describes this straightforward procedure and presents its various applications. It also discusses the limitations and possible difficulties and how to overcome them.

To obtain high-resolution images of fluorescently labeled cells within large tissues, ideally, the biological samples should be imaged without sectioning. 3DISCO is a straightforward tissue clearing procedure based on sequential incubation with organic solvents. Upon clearing, the organs become transparent allowing an end-to-end laser scan of the specimen.

Tissue clearing and subsequent imaging of transparent organs is a powerful method to analyze fluorescently labeled cells and molecules in 3D, in intact ...

Large-scale Reconstructions and Independent, Unbiased Clustering Based on Morphological Metrics to Classify Neurons in Selective Populations

This protocol outlines large-scale reconstructions of neurons combined with the use of independent and unbiased clustering analyses to create a comprehensive survey of the morphological characteristics observed among a selective neuronal population. Combination of these techniques constitutes a novel approach for the collection and analysis of neuroanatomical data. Together, these techniques enable large-scale, and therefore more comprehensive, sampling of selective neuronal populations and establish unbiased quantitative methods for describing morphologically unique neuronal classes within a population.
The protocol outlines the use of modified rabies virus to selectively label neurons. G-deleted rabies virus acts like a retrograde tracer following stereotaxic injection into a target brain structure of interest and serves as a vehicle for the delivery and expression of EGFP in neurons. Large numbers of neurons are infected using this technique and express GFP throughout their dendrites, producing &#34;Golgi-like&#34; complete fills of individual neurons. Accordingly, the virus-mediated retrograde tracing method improves upon traditional dye-based retrograde tracing techniques by producing complete intracellular fills.
Individual well-isolated neurons spanning all regions of the brain area under study are selected for reconstruction in order to obtain a representative sample of neurons. The protocol outlines procedures to reconstruct cell bodies and complete dendritic arborization patterns of labeled neurons spanning multiple tissue sections. Morphological data, including positions of each neuron within the brain structure, are extracted for further analysis. Standard programming functions were utilized to perform independent cluster analyses and cluster evaluations based on morphological metrics. To verify the utility of these analyses, statistical evaluation of a cluster analysis performed on 160 neurons reconstructed in the thalamic reticular nucleus of the thalamus (TRN) of the macaque monkey was made. Both the original cluster analysis and the statistical evaluations performed here indicate that TRN neurons are separated into three subpopulations, each with unique morphological characteristics.

This protocol describes large-scale reconstructions of selective neuronal populations, labeled following retrograde infection with a modified rabies virus expressing fluorescent markers, and independent, unbiased cluster analyses that enable comprehensive characterization of morphological metrics among distinct neuronal subclasses.

This protocol outlines large-scale reconstructions of neurons combined with the use of independent and unbiased clustering analyses to create a ...

Large-scale Three-dimensional Imaging of Cellular Organization in the Mouse Neocortex

The mammalian neocortex is composed of many types of excitatory and inhibitory neurons, each with specific electrophysiological and biochemical properties, synaptic connections, and in vivo functions, but their basic functional and anatomical organization from cellular to network scale is poorly understood. Here we describe a method for the three-dimensional imaging of fluorescently-labeled neurons across large areas of the brain for the investigation of the cortical cellular organization. Specific types of neurons are labeled by the injection of fluorescent retrograde neuronal tracers or expression of fluorescent proteins in transgenic mice. Block brain samples, e.g., a hemisphere, are prepared after fixation, made transparent with tissue clearing methods, and subjected to fluorescent immunolabeling of the specific cell types. Large areas are scanned using confocal or two-photon microscopes equipped with large working distance objectives and motorized stages. This method can resolve the periodic organization of the cell type-specific microcolumn functional modules in the mouse neocortex. The procedure can be useful for the study of three-dimensional cellular architecture in the diverse brain areas and other complex tissues.

Here we describe a procedure for tissue clearing, fluorescent labeling, and large-scale imaging of mouse brain tissue which, thereby, enables visualization of the three-dimensional organization of cell types in the neocortex.

The mammalian neocortex is composed of many types of excitatory and inhibitory neurons, each with specific electrophysiological and biochemical ...

Microsurgical Dissection and Tissue Clearing for High Resolution Intact Whole Retina and Vitreous Imaging

Neuronal and vascular structures of the retina in physiologic and pathologic conditions can be better visualized and characterized by using intact whole retina imaging techniques compared to conventional retinal flat mount preparations and sections. However, immunofluorescent imaging of intact whole retina is hindered by the opaque coatings of the eyeball, i.e., sclera, choroid, and retinal pigment epithelium (RPE) and the light scattering properties of retinal layers that prevent full thickness high resolution optical imaging. Chemical bleaching of the pigmented layers and tissue clearing protocols have been described to address these obstacles; however, currently described methods are not suitable for imaging endogenous fluorescent molecules such as green fluorescent protein (GFP) in intact whole retina. Other approaches bypassed this limitation by surgical removal of pigmented layers and the anterior segment of the eyeball allowing intact eye imaging, though the peripheral retina and hyaloid structures were disrupted. Presented here is an intact whole retina and vitreous immunofluorescent imaging protocol that combines surgical dissection of the sclera/choroid/retina pigment epithelium (RPE) layers with a modified tissue clearing method and light sheet fluorescent microscopy (LSFM). The new approach offers an unprecedented view of&#160;unperturbed vascular and neuronal elements of the retina as well as the vitreous and hyaloid vascular system in pathologic conditions.

Presented here is a protocol for intact whole retina imaging in which the outer opaque/pigmented layers of the eyeball are surgically removed, and optical clearing is applied to render retina transparent enabling the visualization of the peripheral retina and hyaloid vasculature in intact retina using light sheet fluorescent microscopy.

Neuronal and vascular structures of the retina in physiologic and pathologic conditions can be better visualized and characterized by using intact whole ...

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.

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 ...

Whole-Brain Single-Cell Imaging and Analysis of Intact Neonatal Mouse Brains Using MRI, Tissue Clearing, and Light-Sheet Microscopy

Tissue clearing followed by light-sheet microscopy (LSFM) enables cellular-resolution imaging of intact brain structure, allowing quantitative analysis of structural changes caused by genetic or environmental perturbations. Whole-brain imaging results in more accurate quantification of cells and the study of region-specific differences that may be missed with commonly used microscopy of physically sectioned tissue. Using light-sheet microscopy to image cleared brains greatly increases acquisition speed as compared to confocal microscopy. Although these images produce very large amounts of brain structural data, most computational tools that perform feature quantification in images of cleared tissue are limited to counting sparse cell populations, rather than all nuclei.
Here, we demonstrate NuMorph (Nuclear-Based Morphometry), a group of analysis tools, to quantify all nuclei and nuclear markers within annotated regions of a postnatal day 4 (P4) mouse brain after clearing and imaging on a light-sheet microscope. We describe magnetic resonance imaging (MRI) to measure brain volume prior to shrinkage caused by tissue clearing dehydration steps, tissue clearing using the iDISCO+ method, including immunolabeling, followed by light-sheet microscopy using a commercially available platform to image mouse brains at cellular resolution. We then demonstrate this image analysis pipeline using NuMorph, which is used to correct intensity differences, stitch image tiles, align multiple channels, count nuclei, and annotate brain regions through registration to publicly available atlases.
We designed this approach using publicly available protocols and software, allowing any researcher with the necessary microscope and computational resources to perform these techniques. These tissue clearing, imaging, and computational tools allow measurement and quantification of the three-dimensional (3D) organization of cell-types in the cortex and should be widely applicable to any wild-type/knockout mouse study design.

This protocol describes methods for conducting magnetic resonance imaging, clearing, and immunolabeling of intact mouse brains using iDISCO+, followed by a detailed description of imaging using light-sheet microscopy, and downstream analyses using NuMorph.

Tissue clearing followed by light-sheet microscopy (LSFM) enables cellular-resolution imaging of intact brain structure, allowing quantitative analysis of ...

ACT-PRESTO: Biological Tissue Clearing and Immunolabeling Methods for Volume Imaging

The identification and exploration of the detailed organization of organs or of the whole body at the cellular level are fundamental challenges in biology. Transitional methods require a substantial amount of time and effort to obtain a 3D image and including sectioning the intact tissue, immunolabeling, and imaging serially-sectioned tissue, which produces a loss of information at each step of the process. In recently developed approaches for high-resolution imaging within intact tissue, molecular characterization has been restricted to the labeling of proteins. However, currently available protocols for organ clearing require a considerably long process time, making it difficult to implement tissue clearing techniques in the lab. We recently established a rapid and highly-reproducible protocol termed ACT-PRESTO (active clarity technique&#8211;pressure related efficient and stable transfer of macromolecules into organs), which allows for tissue clearance within several hours. Moreover, ACT-PRESTO enables rapid immunolabeling with conventional methods and accelerates antibody penetration into the deep layer of densely-formed, thick specimens by applying pressure or convection flow. We describe how to prepare tissues, how to clear by lipid removal using electrophoresis, and how to immuno-stain by a pressure-assisted delivery. The rapidity and consistency of the protocol will expedite the performance of 3D histological research and volume-based diagnoses.

ACT-PRESTO (active clarity technique&#8211;pressure related efficient and stable transfer of macromolecules into organs) enables rapid, efficient, but inexpensive tissue clearing and rapid antibody penetration into cleared tissues by passive diffusion or pressure-assisted delivery. Using this method, a wide range of tissues can be cleared, immunolabeled by multiple antibodies, and volume-imaged.

The identification and exploration of the detailed organization of organs or of the whole body at the cellular level are fundamental challenges in ...

Biology

High-Resolution 3D Reconstruction of Human Brain with SHORT and LSFM

Optical Clearing and Labeling for Light-sheet Fluorescence Microscopy in Large-scale Human Brain Imaging

Despite the numerous clearing techniques that emerged in the last decade, processing postmortem human brains remains a challenging task due to its dimensions and complexity, which make imaging with micrometer resolution particularly difficult. This paper presents a protocol to perform the reconstruction of volumetric portions of the human brain by simultaneously processing tens of sections with the SHORT (SWITCH - H2O2 - Antigen Retrieval - 2,2'-thiodiethanol [TDE]) tissue transformation protocol, which enables clearing, labeling, and sequential imaging of the samples with light-sheet fluorescence microscopy (LSFM). SHORT provides rapid tissue clearing and homogeneous multi-labeling of thick slices with several neuronal markers, enabling the identification of different neuronal subpopulations in both white and grey matter. After clearing, the slices are imaged via LSFM with micrometer resolution and in multiple channels simultaneously for a rapid 3D reconstruction. By combining SHORT with LSFM analysis within a routinely high-throughput protocol, it is possible to obtain the 3D cytoarchitecture reconstruction of large volumetric areas at high resolution in a short time, thus enabling comprehensive structural characterization of the human brain.

Author Spotlight: Advancing 3D Cytoarchitecture Analysis - Rapid Volumetric Reconstruction of the Human Brain 

The present protocol provides a step-by-step procedure for rapid and simultaneous optical clearing, muti-round labeling, and 3D volumetric reconstruction of tens of postmortem human brain sections by combining the (SWITCH - H2O2 - Antigen Retrieval - 2,2'-thiodiethanol [TDE]) SHORT tissue transformation technique with light-sheet fluorescence microscopy imaging in a routinely high-throughput protocol.

Despite the numerous clearing techniques that emerged in the last decade, processing postmortem human brains remains a challenging task due to its ...

Extraction and Tissue Clearing Preparation of Mouse Brain-Spinal Cord Samples

Neuroscience research focused on the central nervous system necessitates a thorough understanding of various factors, including cell distribution, neuronal connectivity, and molecular dynamics. Traditional methodologies for investigating the entire brain and spinal cord often involve isolating, sectioning, and scanning tissue slices, followed by the labor-intensive process of three-dimensional image reconstruction. This conventional approach can be both time-consuming and cumbersome. Advancements in tissue clearing and whole-organ imaging techniques have revolutionized the analysis of the entire brain and spinal cord. To maximize the potential of these innovative methods, it is essential to extract and clear brain samples while maintaining their connection to the spinal cord. This protocol provides a detailed and systematic guide for preparing samples of the brain connected to the spinal cord, outlining the extraction and clearing procedures. By streamlining these processes, this approach significantly enhances experimental efficiency and data integrity, thereby fostering advancements in neuroscience research and enabling more comprehensive investigations into the complexities of the central nervous system.

This study presents a protocol for extracting and preparing cleared whole-brain and spinal cord samples with preserved fluorescent signals, enhancing experimental efficiency and data integrity to advance neuroscience research.

A Tissue Clearing Method for Neuronal Imaging from Mesoscopic to Microscopic Scales

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