The authors thank Thomas C. Mettenleiter and Verena te Kamp for critically reading the manuscript. This work was supported by the Federal Excellence Initiative of Mecklenburg Western Pomerania and the European Social Fund (ESF) Grant KoInfekt (ESF/14-BM-A55-0002/16) and an intramural collaborative research grant on Lyssaviruses at the Friedrich-Loeffler-Institute (Ri-0372).

RABV-infected, PFA-fixed archived brain material was used. The respective animal experimental studies were evaluated by the responsible animal care, use, and ethics committee of the State Office for Agriculture, Food Safety, and Fishery in Mecklenburg-Western Pomerania (LALFF M-V) and gained approval with permissions 7221.3-2.1-002/11 (mice) and 7221.3-1-068/16 (ferrets). General care and methods used in the animal experiments were carried out according to the approved guidelines.
CAUTION: This protocol uses various toxic and/or harmful substances, including PFA, methanol (MeOH), hydrogen peroxide (H2O2), sodium azide (NaN3), TBA, BA, BB, and DPE. MeOH and TBA are highly flammable. Avoid exposure by wearing appropriate personal protective equipment (a lab coat, gloves, and eye protection) and conducting experiments in a fume hood. Collect waste separately in appropriate containers and dispose of it according to local regulations. Rabies virus is classified as a biosafety level (BSL)-2 pathogen and can, therefore, generally be handled under BSL-2 conditions. Some activities, including procedures that may generate aerosols, work with high virus concentrations, or work with novel lyssaviruses, may require BSL-3 classification. Pre-exposure prophylaxis is recommended for high-risk personnel, including animal caretakers and laboratory workers18,19. Refer to local authority regulations.
1. Fixation of brain tissue and sectioning
<ol>
	<li>Fix brain samples in an appropriate volume of 4% PFA in phosphate-buffered saline (PBS [pH 7.4]) for at least 48 h at 4 &#176;C (with an approximate tissue-to-fixative ratio of 1:10 [v/v]).</li>
	<li>Wash the tissue samples 3x in PBS for at least 30 min each wash and store them in 0.02% NaN3/PBS at 4 &#176;C until use.</li>
	<li>Section the tissue into 1 mm-thick sections using a vibratome (blade feed rate: 0.3&#8211;0.5 mm/s, amplitude: 1 mm, slice thickness: 1,000 &#181;m).</li>
	<li>To retain the correct slice sequence, store each tissue section separately in a well of a multiwell cell culture plate. Add 0.02% NaN3/PBS and store the tissue sections at 4 &#176;C until use.</li>
</ol>
2. Sample pretreatment with methanol
NOTE: Perform all incubation steps with gentle oscillation and, if not indicated otherwise, at room temperature. Protect the samples from light. The sample pretreatment serves the overall purpose of improving antibody diffusion and reducing tissue autofluorescence by exposure to MeOH and H2O2, respectively14.
<ol>
	<li>Prepare 20% (v/v), 40%, 60%, and 80% MeOH solutions in distilled water. For instance, for 20% MeOH, add 10 mL of 100% MeOH to 40 mL of distilled water in an appropriate sealable vessel and mix by inverting it.</li>
	<li>Transfer the samples to reasonably sized vessels (e.g., 5 mL reaction tubes). Take care to use materials chemically resistant to the reagents used in this protocol. For instance, note that while polypropylene is suitable, polystyrene is not. 
	NOTE: The volume specifications in this protocol refer to 5 mL reaction tubes. If a different vessel is used, adjust the volumes accordingly.</li>
	<li>Incubate the samples in 4 mL of each concentration of the prepared series of MeOH solutions in ascending order for 1 h each.</li>
	<li>Incubate the samples 2x for 1 h each in pure (100%) MeOH.</li>
	<li>Cool the samples to 4 &#176;C (e.g., in a laboratory-safe refrigerator).</li>
	<li>Prepare bleaching solution (5% H2O2 in MeOH) by, for instance, diluting a 30% H2O2 stock solution at 1:6 in pure (100%) MeOH, and chill it at 4 &#176;C.</li>
	<li>Remove the 100% MeOH from the refrigerated samples and add 4 mL of prechilled bleaching solution (5% H2O2 in MeOH). Incubate overnight at 4 &#176;C.</li>
	<li>Exchange the bleaching solution for 4 mL of 80% MeOH and incubate for 1 h. Continue using the prepared series of MeOH solutions in descending order for 1 h each until the samples have been incubated for 1 h in 4 mL of 20% MeOH.</li>
	<li>Wash the samples 1x for 1 h with 4 mL of PBS.</li>
</ol>
3. Immunostaining
NOTE: Perform all incubation steps with gentle oscillation and, if not indicated otherwise, at room temperature. Protect the samples from light. To prevent microbial growth, add NaN3 to a final concentration of 0.02% to the solutions in this section. Tissue samples are further permeabilized by treatment with nonionic detergents Triton X-100 and Tween 20. Normal serum is used to block unspecific antibody binding. Glycine and heparin are added to reduce the immunolabeling background14.
<ol>
	<li>Wash the samples 2x for 1 h each in 4 mL of 0.2% Triton X-100/PBS.</li>
	<li>Permeabilize the samples for 2 days at 37 &#176;C with 4 mL of 0.2% Triton X-100/20% DMSO/0.3 M glycine/PBS.</li>
	<li>Block the unspecific binding of antibodies by incubating the samples for 2 days at 37 &#176;C in 4 mL of 0.2% Triton X-100/10% DMSO/6% normal serum/PBS. 
	NOTE: Use normal serum from the same species the secondary antibody was raised in to achieve ideal blocking results.</li>
	<li>Incubate the samples in 2 mL of primary antibody solution (3% normal serum/5% DMSO/PTwH [PBS-Tween 20 with heparin] + primary antibody/antibodies) for 5 days at 37 &#176;C. Refresh the primary antibody solution&#160;after 2.5 days.
	<ol>
		<li>For PTwH, reconstitute the heparin sodium salt in distilled water to make a 10 mg/mL stock solution (store this solution, aliquoted, at 4 &#176;C). Add the stock solution to 0.2% Tween 20/PBS to a final concentration of 10 &#181;g/mL. 
		NOTE: Choosing the correct antibody dilution may require optimization. Generally, standard immunohistochemistry concentrations are a good starting point.</li>
	</ol>
	</li>
	<li>Wash the samples for 1 day in 4 mL of PTwH, exchanging the wash buffer at least 4x&#8211;5x during the course of the day and leaving the final wash on overnight.</li>
	<li>Incubate the samples in 2 mL of secondary antibody solution (3% normal serum/PTwH + secondary antibody/antibodies) for 5 days at 37 &#176;C. Refresh the secondary antibody solution after 2.5 days.
	<ol>
		<li>Dilute the secondary antibody/antibodies at 1:500 in secondary antibody solution (i.e., 4 &#181;L in 2 mL).</li>
	</ol>
	</li>
	<li>Wash the samples for 1 day as described in step 3.5, leaving the final wash on overnight.</li>
</ol>
4. Nuclear staining
NOTE: Perform all incubation steps with gentle oscillation and, if not indicated otherwise, at room temperature. Protect the samples from light. If no nuclear staining is required or the excitation wavelength/emission spectrum of TO-PRO-3 is required for the excitation or detection of another fluorophore, skip this step.
<ol>
	<li>Dilute the nucleic acid stain TO-PRO-3 at 1:1,000 in PTwH and incubate the samples in 4 mL of nuclear staining solution for 5 h.</li>
	<li>Wash the samples for 1 day as described in step 3.5, leaving the final wash on overnight. 
	NOTE: Following the washes, the samples can be stored in PBS at 4 &#176;C until optical clearing.</li>
</ol>
5. Tissue clearing
NOTE: Perform all incubation steps with gentle oscillation and, if not indicated otherwise, at room temperature. Protect the samples from light. The tissue samples are dehydrated in a graded series of TBA solutions. As immunostaining requires aqueous solutions, all staining procedures have to be finished prior to tissue clearing. Optical clearance and refractive index matching are achieved by treatment with a mixture of BA, BB, and DPE. The clearing solution is supplemented with DL-&#945;-tocopherol as an antioxidant13.
<ol>
	<li>Prepare 30% (v/v), 50%, 70%, 80%, 90%, and 96% TBA solutions in distilled water. For instance, for 30% TBA, add 15 mL of 100% TBA to 35 mL of distilled water in an appropriate sealable vessel and mix by inverting. 
	NOTE: TBA has a melting point of 25&#8211;26 &#176;C; thus, it tends to be solid at room temperature. In order to prepare TBA solutions, heat the well-sealed bottle at 37 &#176;C in an incubator or water bath.</li>
	<li>Dehydrate the samples with 4 mL of each concentration of the prepared series of TBA solutions in ascending order for 2 h each. Leave the 96% TBA on overnight.</li>
	<li>Dehydrate the samples further in pure (100%) TBA for 2 h.</li>
	<li>Prepare clearing solution BABB-D15. 
	NOTE: BABB-D15 is a combination of BA and BB (BABB) which is mixed with DPE at a ratio of x:1, where x is specified in the solution&#8217;s name, in this case 15.
	<ol>
		<li>For BABB, mix one-part BA with two parts BB.</li>
		<li>Mix BABB and DPE at a ratio of 15:1.</li>
		<li>Add 0.4 vol% DL-&#945;-tocopherol (vitamin E). 
		NOTE: For example, for 20 mL of BABB-D15, mix 6.25 mL of BA with 12.5 mL of BB. Add 1.25 mL of DPE and supplement it with 0.08 mL of DL-&#945;-tocopherol.</li>
	</ol>
	</li>
	<li>Clear the samples in clearing solution until they are optically transparent (2&#8211;6 h).</li>
	<li>The samples can be stored at 4 &#176;C in BABB-D15, protected from light, until mounting and imaging.</li>
</ol>
6. Sample mounting
<ol>
	<li>Using a 3D printer, print the&#160;imaging chamber and the lid (material: copolyester [CPE], nozzle: 0.25 mm, layer height: 0.06 mm, wall thickness: 0.88 mm, wall count: 4, infill: 100%, no support structure; the corresponding .STL file can be found in the <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/59402/3D_ImagingChamber.stl" target="_blank">Supplementary Materials</a> of this protocol).</li>
	<li>Assemble the imaging chamber (Figure 2).
	<ol>
		<li>Mount a round coverslip (diameter: 30 mm) on the imaging chamber with RTV-1 (one-component room-temperature-vulcanizing) silicone rubber. Remove the excess silicone rubber with a water-wetted cotton swab and cure overnight.</li>
		<li>Mount a round coverslip (diameter: 22 mm) on the lid with RTV-1 silicone rubber. Remove the&#160;excess silicone rubber&#160;with a water-wetted cotton swab and cure overnight.</li>
	</ol>
	</li>
	<li>Place the sample in an imaging chamber, add a small volume of BABB-D15, and insert the lid. Fill the chamber up with BABB-D15 through the inlet, using a hypodermic needle (27 G x 3/4 inch [0.40 mm x 20 mm]).</li>
	<li>Plug the inlet and seal the imaging chamber with RTV-1 silicone rubber. Cure overnight in the dark.</li>
</ol>
7. Imaging and image processing
<ol>
	<li>Set up the image acquisition by selecting the respective laser lines to match the fluorophores used. Adjust the detection ranges of each detector to prevent signal overlap between channels. 
	NOTE: Exemplary detection ranges for Alexa Fluor 488, Alexa Fluor 568, and TO-PRO-3 are 500&#8211;550 nm, 590&#8211;620 nm, and 645&#8211;700 nm, respectively.</li>
	<li>Choose the acquisition parameters, define the upper and lower border of the z-stack, and acquire the image stack. 
	NOTE: Exemplary acquisition parameters are a sequential scan with a pixel size of 60-90 nm, a z-step size of 0.5 &#181;m, a line average of 1, a scan speed of 400 Hz, and a pinhole size of 1 Airy unit.</li>
	<li>Process the image stack using appropriate image analysis software (e.g., Fiji) to generate 3D projections or perform in-depth analyses. 
	NOTE: Because of the large size of the acquired image files, the use of a workstation is usually necessary.
	<ol>
		<li>Open the acquisition or image file(s) in Fiji (File | Open | Select files).
		<ol>
			<li>If using, for instance, .LIF files, select or deselect the desired options in the Bio-Formats dialog window. View stack with Hyperstack. Other than that, no specific selections or ticks are necessary. Press OK.</li>
			<li>If the file contains multiple image stacks, select the ones to be analyzed and confirm by pressing OK.</li>
		</ol>
		</li>
		<li>Perform bleach correction by splitting the merged image into individual channels (Image | Color | Channels Tool, then select More | Split Channels). For each channel, select bleach correction (Image | Adjust | Bleach Correction) and choose Simple Ratio (background intensity: 0.0). 
		NOTE: In some cases, for instance, when there is no linear decay of the signal or the signal is too weak overall, Simple Ratio may fail. Alternatively, try Exponential Fit or skip the bleach correction.</li>
		<li>Adjust the brightness and contrast for each channel using the sliders (Image | Adjust | Brightness | Contrast).</li>
		<li>Merge the channels (Image | Color | Merge Channels), make a composite (Image | Color | Channels Tool, then select More | Make Composite), and convert it to RGB format (Image | Color | Channels Tool, then select More | Convert to RGB).</li>
		<li>If necessary, resize the image stack to reduce the computation time and file size (Image | Adjust | Size, both options ticked plus bilinear interpolation).</li>
		<li>Generate a 3D projection (Image | Stacks | 3D Project). Choose Brightest Point as projection method and set the slice spacing to match the z-step size of the acquired image stack. For maximum quality, set the rotation angle increment to 1 and enable interpolation. Modify the total rotation, transparency thresholds, and opacity as needed.</li>
		<li>If necessary, readjust the contrast and brightness by converting the 3D projection back to 8-bit format (Image | Type | 8-bit). Use the respective sliders (Image | Adjust | Brightness | Contrast) and reconvert the image stack to RGB format as described in step 7.3.4.</li>
		<li>Save the 3D projection as .TIF file (image file format) and .AVI file (video file format).</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

The resurgence and further development of tissue clearing techniques in recent years2,3,4,5,6,7,8,9,10,11,12,13,...

Conventional histology techniques mostly rely on thin sections of organ tissues, which can inherently provide only 2D insights into a complex 3D environment. Although feasible in principle, 3D reconstruction from serial thin sections requires demanding technical pipelines for both slicing and subsequent in silico alignment of the acquired images1. Moreover, seamless reconstruction of z-volumes after microtome slicing is critical as both mechanical and computational artifacts can remain because of suboptimal image registration caused by nonoverlapping image planes, staining variations, and physical destruction of tissue by, for instance, the microtome blade. In contrast, pure optical slicing of intact thick tissue samples allows the acquisition of overlapping image planes (oversampling) and, thereby, facilitates 3D reconstruction. This, in turn, is highly beneficial for the analysis of infection processes in complex cell populations (e.g., neuronal networks in the context of the surrounding glial and immune cells). However, inherent obstacles of thick tissue sections include light scattering and limited antibody penetration into the tissue. In recent years, a variety of techniques has been developed and optimized to overcome these issues2,3,4,5,6,7,8,9,10,11,12,13. Essentially, target tissues are turned optically transparent by treatment with either aqueous2,3,4,5,6,7,8,9 or organic solvent-based10,11,12,13 solutions. The introduction of 3DISCO (3D imaging of solvent-cleared organs)11,12 and its successor uDISCO (ultimate 3D imaging of solvent-cleared organs)13 provided a relatively fast, simple, and inexpensive tool with excellent clearing capabilities. The main constituents of the clearing protocol are the organic solvents tert-butanol (TBA), benzyl alcohol (BA), benzyl benzoate (BB), and diphenyl ether (DPE). The development and addition of iDISCO (immunolabeling-enabled 3D imaging of solvent-cleared organs)14, a compatible immunostaining protocol, constituted another advantage over existing methods and enabled the deep-tissue labeling of antigens of interest, as well as the long-term storage of immunostained samples. Thus, the combination of iDISCO14 and uDISCO13 allows for the high-resolution imaging of antibody-labeled proteins in large tissue sections (up to 1 mm) using conventional CLSM.
The preservation of an organ&#8217;s complex structure in all three dimensions is particularly important for brain tissue. Neurons comprise a very heterogeneous cellular subpopulation with highly diverse 3D morphologies based on their neurite projections (reviewed by Masland15). Furthermore, the brain consists of a number of compartments and subcompartments, each composed of different cellular subpopulations and ratios thereof, including glial cells and neurons (reviewed by von Bartheld et al.16). As a neurotropic virus, the rabies virus (RABV, reviewed by Fooks et al.17) primarily infects neurons, using their transport machinery to travel in retrograde direction along axons from the primary site of infection to the central nervous system (CNS). The protocol described here (Figure 1A) allows for the immunostaining-assisted detection and visualization of RABV and RABV-infected cells in large, coherent image stacks obtained from infected brain tissue. This enables an unbiased, 3D high-resolution assessment of the infection environment. It is applicable to brain tissue from a variety of species, can be performed immediately after fixation or after the long-term storage of samples in paraformaldehyde (PFA), and allows the storage and reimaging of stained and cleared samples for months.

<table><tbody><tr><td>&lt;strong&gt;Reagents&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Benzyl alcohol</td><td>Alfa Aesar</td><td>41218</td><td>Clearing reagent</td></tr><tr><td>Benzyl benzoate</td><td>Sigma-Aldrich</td><td>BB6630-500ML</td><td>Clearing reagent</td></tr><tr><td>Dimethyl sulfoxide</td><td>Carl Roth</td><td>4720.2</td><td>Various buffers</td></tr><tr><td>Diphenyl ether</td><td>Sigma-Aldrich</td><td>240834-100G</td><td>Clearing reagent</td></tr><tr><td>DL-&amp;alpha;-Tocopherol</td><td>Alfa Aesar</td><td>A17039</td><td>Antioxidant</td></tr><tr><td>Donkey serum</td><td>Bio-Rad</td><td>C06SBZ</td><td>Blocking reagent</td></tr><tr><td>Glycine</td><td>Carl Roth</td><td>3908.2</td><td>Background reduction</td></tr><tr><td>Goat serum</td><td>Merck</td><td>S26-100ML</td><td>Blocking reagent</td></tr><tr><td>Heparin sodium salt</td><td>Carl Roth</td><td>7692.1</td><td>Background reduction</td></tr><tr><td>Hydrogen peroxide solution (30 %)</td><td>Carl Roth</td><td>8070.2</td><td>Sample bleaching</td></tr><tr><td>Methanol</td><td>Carl Roth</td><td>4627.4</td><td>Sample pretreatment</td></tr><tr><td>Paraformaldehyde</td><td>Carl Roth</td><td>0335.3</td><td>Crystalline powder to make fixative solution</td></tr><tr><td>Sodium azide</td><td>Carl Roth</td><td>K305.1</td><td>Prevention of microbial growth in stock solutions</td></tr><tr><td>tert-Butanol</td><td>Alfa Aesar</td><td>33278</td><td>Sample dehydration for tissue clearing</td></tr><tr><td>TO-PRO-3</td><td>Thermo Fisher</td><td>T3605</td><td>Nucleic acid stain</td></tr><tr><td>Triton X-100</td><td>Carl Roth</td><td>3051.2</td><td>Detergent</td></tr><tr><td>Tween 20</td><td>AppliChem</td><td>A4974,0500</td><td>Detergent</td></tr><tr><td>&lt;strong&gt;Miscellaneous&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>5 mL reaction tubes</td><td>Eppendorf</td><td>0030119401</td><td>Sample tubes</td></tr><tr><td>Coverslip, circular (diameter: 22 mm)</td><td>Marienfeld</td><td>0111620</td><td>Part of imaging chamber</td></tr><tr><td>Coverslip, circular (diameter: 30 mm)</td><td>Marienfeld</td><td>0111700</td><td>Part of imaging chamber</td></tr><tr><td>Hypodermic needle (27 G x &amp;frac34;&amp;rdquo; [0.40 mm x 20 mm])</td><td>B. Braun</td><td>4657705</td><td>Filling of the imaging chamber with clearing solution</td></tr><tr><td>RTV-1 silicone rubber</td><td>Wacker</td><td>Elastosil E43</td><td>Adhesive for the assembly of the imaging chamber</td></tr><tr><td>Ultimaker CPE 2.85 mm transparent</td><td>Ultimaker</td><td>8718836374869</td><td>Copolyester filament for 3D printer to print parts of the imaging chamber</td></tr><tr><td>&lt;strong&gt;Technical equipment and software&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>3D printer</td><td>Ultimaker</td><td>Ultimaker 2+</td><td>Printing of imaging chamber</td></tr><tr><td>Automated water immersion system</td><td>Leica</td><td>15640019</td><td>Software-controlled water pump</td></tr><tr><td>Benchtop orbital shaker</td><td>Elmi</td><td>DOS-20M</td><td>Sample incubation at room temperature (~ 150 rpm)</td></tr><tr><td>Benchtop orbital shaker, heated</td><td>New Brunswick Scientific</td><td>G24 Environmental Shaker</td><td>Sample incubation at 37 &amp;deg;C (~ 150 rpm)</td></tr><tr><td>Confocal laser scanning microscope</td><td>Leica</td><td>DMI 6000 TCS SP5</td><td>Inverted confocal microscope for sample imaging</td></tr><tr><td>Fiji</td><td>NIH (ImageJ)</td><td>open source software (v1.52h)</td><td>Image processing package based on ImageJ</td></tr><tr><td>Long working distance water immersion objective</td><td>Leica</td><td>15506360</td><td>HC PL APO 40x/1.10 W motCORR CS2</td></tr><tr><td>Vibratome</td><td>Leica</td><td>VT1200S</td><td>Sample slicing</td></tr><tr><td>Workstation</td><td>Dell</td><td>Precision 7920</td><td>CPU: Intel Xeon Gold 5118&lt;br/&gt; GPU: Nvidia Quadro P5000&lt;br/&gt; RAM: 128 GB 2666 MHz DDR4&lt;br/&gt; SSD: 2 TB</td></tr><tr><td>&lt;strong&gt;Primary antibodies&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Goat anti-RABV N</td><td>Friedrich-Loeffler-Institut</td><td></td><td>Monospecific polyclonal goat anti-RABV N serum, generated by goat immunization with baculovirus-expressed and His-tag-purified RABV nucleoprotein N&lt;br/&gt; Dilution: 1:400</td></tr><tr><td>Rabbit anti-GFAP</td><td>Dako</td><td>Z0334</td><td>Polyclonal antibody (RRID:AB_10013382)&lt;br/&gt; Dilution: 1:100</td></tr><tr><td>Rabbit anti-MAP2</td><td>Abcam</td><td>ab32454</td><td>Polyclonal antibody (RRID:AB_776174)&lt;br/&gt; Dilution: 1:250</td></tr><tr><td>Rabbit anti-RABV P 160-5</td><td>Friedrich-Loeffler-Institut</td><td></td><td>Monospecific polyclonal rabbit anti-RABV P serum, generated by rabbit immunization with baculovirus-expressed and His-tag-purified RABV phosphoprotein P (see reference 23: Orbanz et al., 2010)&lt;br/&gt; Dilution: 1:1,000</td></tr><tr><td>&lt;strong&gt;Secondary antibodies&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Donkey anti-goat IgG</td><td>Thermo Fisher Scientific</td><td>depending on conjugated fluorophore</td><td>Highly cross-absorbed&lt;br/&gt; Dilution: 1:500</td></tr><tr><td>Donkey anti-mouse IgG</td><td>Thermo Fisher Scientific</td><td>depending on conjugated fluorophore</td><td>Highly cross-absorbed&lt;br/&gt; Dilution: 1:500</td></tr><tr><td>Donkey anti-rabbit IgG</td><td>Thermo Fisher Scientific</td><td>depending on conjugated fluorophore</td><td>Highly cross-absorbed&lt;br/&gt; Dilution: 1:500</td></tr><tr><td>Goat anti-mouse IgG</td><td>Thermo Fisher Scientific</td><td>depending on conjugated fluorophore</td><td>Highly cross-absorbed&lt;br/&gt; Dilution: 1:500</td></tr><tr><td>Goat anti-rabbit IgG</td><td>Thermo Fisher Scientific</td><td>depending on conjugated fluorophore</td><td>Highly cross-absorbed&lt;br/&gt; Dilution: 1:500</td></tr></tbody></table>

high-resolution 3d imaging of rabies virus infection in solvent-cleared brain tissue

The visualization of infection processes in tissues and organs by immunolabeling is a key method in modern infection biology. The ability to observe and study the distribution, tropism, and abundance of pathogens inside of organ tissues provides pivotal data on disease development and progression. Using conventional microscopy methods, immunolabeling is mostly restricted to thin sections obtained from paraffin-embedded or frozen samples. However, the limited 2D image plane of these thin sections may lead to the loss of crucial information on the complex structure of an infected organ and the cellular context of the infection. Modern multicolor, immunostaining-compatible tissue clearing techniques now provide a relatively fast and inexpensive way to study high-volume 3D image stacks of virus-infected organ tissue. By exposing the tissue to organic solvents, it becomes optically transparent. This matches the sample&#8217;s refractive indices and eventually leads to a significant reduction of light scattering. Thus, in combination with long free working distance objectives, large tissue sections up to 1 mm in size can be imaged by conventional confocal laser scanning microscopy (CLSM) at high resolution. Here, we describe a protocol to apply deep-tissue imaging after tissue clearing to visualize rabies virus distribution in infected brains in order to study topics like virus pathogenesis, spread, tropism, and neuroinvasion.

The combination of iDISCO14 and uDISCO13 coupled with high-resolution CLSM provides deep insights into the spatiotemporal resolution and plasticity of RABV infection of brain tissue and the surrounding cellular context.
Using immunostaining of RABV phosphoprotein (P), complex layers of infected neuronal cells can be visualized in thick sections of mouse brains (Figure 3). Subsequently, seamless 3D projections of the ...

Watch this Scientific Journal Video about High-Resolution 3D Imaging of Rabies Virus Infection in Solvent-Cleared Brain Tissue at JoVE.com

High-Resolution 3D Imaging of Rabies Virus Infection in Solvent-Cleared Brain Tissue

Novel, immunostaining-compatible tissue clearing techniques like the ultimate 3D imaging of solvent-cleared organs allow the 3D visualization of rabies virus brain infection and its complex cellular environment. Thick, antibody-labeled brain tissue slices are made optically transparent to increase imaging depth and to enable 3D analysis by confocal laser scanning microscopy.

The visualization of infection processes in tissues and organs by immunolabeling is a key method in modern infection biology. The ability to observe and ...

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Neuroscience

10.8K Views.  Federal Research Institute for Animal Health. Novel, immunostaining-compatible tissue clearing techniques like the ultimate 3D imaging of solvent-cleared organs allow the 3D visualization of rabies virus brain infection and its complex cellular environment. Thick, antibody-labeled brain tissue slices are made optically transparent to increase imaging depth and to enable 3D analysis by confocal laser scanning microscopy.

Video: High-Resolution 3D Imaging of Rabies Virus Infection in Solvent-Cleared Brain Tissue

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 ...

3-D Imaging and Analysis of Neurons Infected In Vivo with Toxoplasma gondii

Toxoplasma gondii is an obligate, intracellular parasite with a broad host range, including humans and rodents. In both humans and rodents, Toxoplasma establishes a lifelong persistent infection in the brain. While this brain infection is asymptomatic in most immunocompetent people, in the developing fetus or immunocompromised individuals such as acquired immune deficiency syndrome (AIDS) patients, this predilection for and persistence in the brain can lead to devastating neurologic disease. Thus, it is clear that the brain-Toxoplasma interaction is critical to the symptomatic disease produced by Toxoplasma, yet we have little understanding of the cellular or molecular interaction between cells of the central nervous system (CNS) and the parasite. In the mouse model of CNS toxoplasmosis it has been known for over 30 years that neurons are the cells in which the parasite persists, but little information is available about which part of the neuron is generally infected (soma, dendrite, axon) and if this cellular relationship changes between strains. In part, this lack is secondary to the difficulty of imaging and visualizing whole infected neurons from an animal. Such images would typically require serial sectioning and stitching of tissue imaged by electron microscopy or confocal microscopy after immunostaining. By combining several techniques, the method described here enables the use of thick sections (160 &#181;m) to identify and image whole cells that contain cysts, allowing three-dimensional visualization and analysis of individual, chronically infected neurons without the need for immunostaining, electron microscopy, or serial sectioning and stitching. Using this technique, we can begin to understand the cellular relationship between the parasite and the infected neuron.

3-D Imaging and Analysis of Neurons Infected In Vivo with Toxoplasma gondii

Using this protocol, we were able to image 160 &#181;m thick brain sections from mice infected with the parasite Toxoplasma gondii, which enables visualization and analysis of the spatial relationship between the encysting parasite and the infected neuron.

Toxoplasma gondii is an obligate, intracellular parasite with a broad host range, including humans and rodents. In both humans and rodents, Toxoplasma ...

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

This video shows the craniotomy procedure that allows chronic imaging of neurons in the mouse retrosplenial cortex (RSC) using in vivo two-photon microscopy in Thy1-GFP transgenic mouse line. This approach creates a possibility to investigate the correlation of behavioural manipulations with changes in neuronal morphology in vivo.
The cranial window implantation procedure was considered to be limited only to the easily accessible cortex regions such as the barrel field. Our approach allows visualization of neurons in the highly vascularized RSC. RSC is an important element of the brain circuit responsible for spatial memory, previously deemed to be problematic for in vivo two-photon imaging.
The cranial window implantation over the RSC is combined with an injection of mCherry-expressing recombinant adeno-associated virus (rAAVmCherry) into the dorsal hippocampus. The expressed mCherry spreads out to axonal projections from the hippocampus to RSC, enabling the visualization of changes in both presynaptic axonal boutons and postsynaptic dendritic spines in the cortex. 
This technique allows long-term monitoring of experience-dependent structural plasticity in RSC.

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

This video shows the craniotomy procedure that allows chronic imaging of neurons in mouse retrosplenial cortex using in vivo two photon microscopy in Thy1-GFP transgenic line. This approach is combined with injection of mCherry-expressing adeno-associated virus into dorsal hippocampus. These techniques allow long-term monitoring of experience-dependent structural plasticity in RSC.

This video shows the craniotomy procedure that allows chronic imaging of neurons in the mouse retrosplenial cortex (RSC) using in vivo two-photon ...

Enhanced Rabies Surveillance Using a Direct Rapid Immunohistochemical Test

Laboratory-based surveillance is integral for rabies prevention, control and management efforts. While the DFA is the gold standard for rabies diagnosis, there is a need to validate additional diagnostic techniques to improve rabies surveillance, particularly in developing countries. Here, we present a standard protocol for the DRIT as an alternative, laboratory or field-based testing option that uses light microscopy as compared to the DFA. Touch impressions of brain tissue collected from suspect animals are fixed in 10% buffered formalin. The DRIT uses rabies virus-specific monoclonal or polyclonal antibodies (conjugated to biotin), a streptavidin-peroxidase enzyme, and a chromogen reporter (such as acetyl 3-amino-9-ethylcarbazole) to detect viral inclusions within infected tissue. In approximately 1 h, a brain tissue sample can be tested and interpreted by the DRIT. Evaluation of suspect animal brains tested from a variety of species in North America, Asia, Africa, and Europe have illustrated high sensitivity and specificity by the DRIT approaching 100% with results compared to DFA. Since 2005, the United States Department of Agriculture&#8217;s Wildlife Services (USDA WS) program has conducted large-scale enhanced rabies surveillance efforts using the DRIT to test &#62;94,000 samples collected from wildlife in strategic rabies management areas. The DRIT provides a powerful, economical tool for rabies diagnosis that can be used by laboratorians and field biologists to improve current rabies surveillance, prevention and control programs globally.

The direct rapid immunohistochemical test (DRIT) offers a World Organization for Animal Health and World Health Organization (OIE/WHO) recognized alternative to the direct fluorescent antibody (DFA) test for rabies diagnosis. This test allows for field-based applications that can be performed in approximately 1 h upon brain impressions using light microscopy.

Laboratory-based surveillance is integral for rabies prevention, control and management efforts. While the DFA is the gold standard for rabies diagnosis, ...

Immunology and Infection

Field Postmortem Rabies Rapid Immunochromatographic Diagnostic Test for Resource-Limited Settings with Further Molecular Applications

Functional rabies surveillance systems are crucial to provide reliable data and increase the political commitment necessary for disease control. To date, animals suspected as rabies-positive must be submitted to a postmortem confirmation using classical or molecular laboratory methods. However, most endemic areas are in low- and middle-income countries where animal rabies diagnosis is restricted to central veterinary laboratories. Poor availability of surveillance infrastructure leads to serious disease underreporting from remote areas. Several diagnostic protocols requiring low technical expertise have been recently developed, providing opportunity to establish rabies diagnosis in decentralized laboratories. We present here a complete protocol for field postmortem diagnosis of animal rabies using a rapid immunochromatographic diagnostic test (RIDT), from brain biopsy sampling to the final interpretation. We complete the protocol by describing a further use of the device for molecular analysis and viral genotyping. RIDT easily detects rabies virus and other lyssaviruses in brain samples. The principle of such tests is simple: brain material is applied on a test strip where gold conjugated antibodies bind specifically to rabies antigens. The antigen-antibody complexes bind further to fixed antibodies on the test line, resulting in a clearly visible purple line. The virus is inactivated in the test strip, but viral RNA can be subsequently extracted. This allows the test strip, rather than the infectious brain sample, to be safely and easily sent to an equipped laboratory for confirmation and molecular typing. Based on a modification of the manufacturer&#39;s protocol, we found increased test sensitivity, reaching 98% compared to the gold standard reference method, the direct immunofluorescence antibody test. The advantages of the test are numerous: rapid, easy-to-use, low cost and no requirement for laboratory infrastructure, such as microscopy or cold-chain compliance. RIDTs represent a useful alternative for areas where reference diagnostic methods are not available.

We present a complete protocol for postmortem diagnosis of animal rabies under field conditions using a rapid immunochromatographic diagnostic test (RIDT), from brain biopsy sampling to final interpretation. We also describe further applications using the device for molecular analysis and viral genotyping.

Functional rabies surveillance systems are crucial to provide reliable data and increase the political commitment necessary for disease control. To date, ...

Immunohistochemistry Test for the Lyssavirus Antigen Detection from Formalin-Fixed Tissues

One of the primary diagnostic modalities for rabies is the detection of viral ribonucleoprotein (RNP) complex (antigen) in the infected tissue samples. While the direct fluorescent antibody (DFA) test or the direct rapid immunohistochemical test (DRIT) are most commonly utilized for the antigen detection, both tests require fresh and/or frozen tissues for impressions on slides prior to the antigen detection using antibodies. If samples are collected and fixed in formalin, neither test is optimal for the antigen detection, however, testing can be performed by conventional immunohistochemistry (IHC) after embedding in paraffin blocks and sectioning. With this IHC method, tissues are stained with anti-rabies antibodies, sections are deparaffinized, antigen retrieved by partial proteolysis or other methods, and incubated with primary and secondary antibodies. Antigens are stained using horseradish peroxidase / amino ethyl carbazole and counterstained with hematoxylin for the visualization using a light microscope. In addition to the specific antigen detection, formalin fixation offers other advantages like the determination of histological changes, relaxed conditions for specimen storage and transport (under ambient temperatures), ability to test retrospective cases and improved biological safety through the inactivation of infectious agents.

Here, we present an immunohistochemistry test protocol for the detection of rabies virus antigen as an alternative diagnostic test for formalin-fixed tissues.

One of the primary diagnostic modalities for rabies is the detection of viral ribonucleoprotein (RNP) complex (antigen) in the infected tissue samples. ...

Quantitation of Rabies Virus in Various Bovine Brain Structures

Bovine paralytic rabies (BPR) is a form of viral encephalitis that is of substantial economic importance throughout Latin America, where it poses a major zoonotic risk. Here, our objective was to utilize a laboratory protocol to determine the relative copy number of the rabies virus (RABV) genome in different bovine brain anatomical structures using quantitative real-time reverse transcription polymerase chain reaction (qRT-PCR). qRT-PCR quantifies the specific number of gene copies present in a sample based on fluorescence emitted after amplification that is directly proportional to the amount of target nucleic acid present in the sample. This method is advantageous owing to its short duration, reduced risk of contamination, and potential to detect viral nucleic acids in different samples more easily compared to other techniques. The brains of six rabid animals were divided into six anatomical structures, namely the Ammon&#8217;s horn, cerebellum, cortex, medulla, pons, and thalamus. All brains were identified as positive for RABV antigens based on a direct immunofluorescence test. The same anatomical structures from the brains of four RABV-negative bovines were also assessed. RNA was extracted from each structure and used for qRT-PCR. An assay was performed to determine the copy numbers of RABV genes using an in vitro transcribed nucleoprotein gene. The standard curve used to quantify viral RNA exhibited an efficiency of 100% and linearity of 0.99. Analysis revealed that the cortex, medulla, and thalamus were the ideal CNS portions for use in RABV detection, based on the observation that these structures possessed the highest levels of RABV. The test specificity was 100%. All samples were positive, no false positives were detected. This method can be used to detect RABV in samples that contain low levels of RABV during diagnosis of BPR.

This protocol presents a qRT-PCR-based approach for determining the rabies virus nucleoprotein (N) gene copy number within various bovine brain anatomical structures using in vitro transcription.

Bovine paralytic rabies (BPR) is a form of viral encephalitis that is of substantial economic importance throughout Latin America, where it poses a major ...

A Rat Model of EcoHIV Brain Infection

It has been well studied that the EcoHIV infected mouse model is of significant utility in investigating HIV associated neurological complications. Establishment of the EcoHIV infected rat model for studies of drug abuse and neurocognitive disorders, would be beneficial in the study of neuroHIV and HIV-1 associated neurocognitive disorders (HAND). In the present study, we demonstrate the successful creation of a rat model of active HIV infection using chimeric HIV (EcoHIV). First, the lentiviral construct of EcoHIV was packaged in cultured 293 FT cells for 48 hours. Then, the conditional medium was concentrated and titered. Next, we performed bilateral stereotaxic injections of the EcoHIV-EGFP into F344/N rat brain tissue. One week after infection, EGFP fluorescence signals were detected in the infected brain tissue, indicating that EcoHIV successfully induces an active HIV infection in rats. In addition, immunostaining for the microglial cell marker, Iba1, was performed. The results indicated that microglia were the predominant cell type harboring EcoHIV. Furthermore, EcoHIV rats exhibited alterations in temporal processing, a potential underlying neurobehavioral mechanism of HAND as well as synaptic dysfunction eight weeks after infection. Collectively, the present study extends the EcoHIV model of HIV-1 infection to the rat offering a valuable biological system to study HIV-1 viral reservoirs in the brain as well as HAND and associated comorbidities such as drug abuse.

Here, we present a protocol to establish a new rat model of active HIV infection using chimeric HIV (EcoHIV), which is critical for enhancing our understanding of HIV-1 viral reservoirs in the brain and offering a system to study HIV-associated neurocognitive disorders and associated comorbidities (i.e., drug abuse).

It has been well studied that the EcoHIV infected mouse model is of significant utility in investigating HIV associated neurological complications. ...

Detection of Viral Proteins in Rabies Virus-Infected Mouse Brain Tissue

Collect fixed brain sections from intracerebral rabies virus-infected mice at defined postmortem intervals.&#160;
The virus infects pyramidal neurons in the cerebral cortex, and expressed viral proteins localize within intracytoplasmic granular structures known as inclusion bodies.
Treat the sections with ammonium chloride to neutralize fixation-induced reactive groups and minimize background staining.
Immerse the sections in hydrogen peroxide to inactivate endogenous peroxidase and reduce nonspecific signals.
Apply a blocking solution containing a detergent to permeabilize cellular membranes and mask nonspecific binding sites.
Incubate with primary antibodies specific to the viral proteins.
Wash and incubate with biotin-conjugated secondary antibodies that bind to the primary antibodies.
Wash again and introduce peroxidase enzyme-conjugated avidin, which binds to biotin, followed by a chromogenic substrate that the enzyme converts into a colored product.
Visualize the immunoreactive inclusion bodies containing viral proteins using a microscope.
With an increase in postmortem time, neuronal degradation and the disappearance of inclusion bodies become evident.

JoVE Encyclopedia of Experiments

Neuropathology

Infectious Diseases

An Immunohistochemistry Staining for the Detection of Rabies Virus Antigens

Source: Niezgoda, M. et al., <a href="https://appjove.unicartagenaproxy.elogim.com/v/60138">Immunohistochemistry Test for the Lyssavirus Antigen Detection from Formalin-Fixed Tissues.</a> J. Vis. Exp. (2021)
This video demonstrates immunohistochemistry staining for rabies virus antigen detection in infected mouse brain tissue. The process includes deparaffinization and alcohol treatment, followed by proteolytic treatment to enhance antigen accessibility. Immunohistochemistry staining reveals red precipitates, confirming the presence of rabies antigens in the tissue section.

All procedures involving sample collection have been performed in accordance with the institute&#39;s IRB guidelines.
1. Formalin fixation of tissues

	...

High-Resolution 3D Imaging of Rabies Virus Infection in Solvent-Cleared Brain Tissue

Summary

Explore More Videos

Chapters in this video

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

3-D Imaging and Analysis of Neurons Infected In Vivo with Toxoplasma gondii

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

Enhanced Rabies Surveillance Using a Direct Rapid Immunohistochemical Test

Field Postmortem Rabies Rapid Immunochromatographic Diagnostic Test for Resource-Limited Settings with Further Molecular Applications

Immunohistochemistry Test for the Lyssavirus Antigen Detection from Formalin-Fixed Tissues

Quantitation of Rabies Virus in Various Bovine Brain Structures

A Rat Model of EcoHIV Brain Infection

Detection of Viral Proteins in Rabies Virus-Infected Mouse Brain Tissue

An Immunohistochemistry Staining for the Detection of Rabies Virus Antigens