This work was funded by NIH grants: NS100624, DA013137, HD043680, MH106392 &#38; by T32 Training Grant 5T32GM081740

All animal protocols were reviewed and approved by the Animal Care and Use Committee at the University of South Carolina.
1. Stock solution preparation
<ol>
	<li>Solution 1 (1 L): To 900 mL of deionized H2O add 100 mL of 10x phosphate buffered saline (PBS) and 2 mL of Triton X-100.</li>
	<li>Solution 2 (1 L): To 900 mL of deionized H2O, add 100 mL of 10x PBS, 2mL of Tween-20 and 1 mL of 10 mg/mL Heparin stock solution.</li>
	<li>Solution 3 (500 mL): To 400 mL of Solution 1, add 11.5 g of glycine and 100 mL of dimethylsulfoxide (DMSO).</li>
	<li>Solution 4 (50 mL): To 42 mL of Solution 1, add 3 mL of corresponding serum and 5 mL of DMSO.</li>
	<li>Primary antibody solution (50 mL): To 46 mL of Solution 2, add 2.5 mL of DMSO and 1.5 mL of corresponding serum.</li>
	<li>Secondary antibody solution (50 mL): To 48.5 mL of Solution 2, add 1.5 mL of corresponding serum. 
	NOTE: For Solution 4, primary solution, and secondary solution, use a serum that corresponds with the secondary antibody (e.g., goat, horse, bovine).</li>
</ol>
2. Sample preparation
NOTE: Perform steps 2.1 through 2.7 in a fume hood due to limit the exposure to PFA.
<ol>
	<li>Deeply anesthetize young (3-6 week) rat using sevoflurane; proceed to 2.2 when rat is not responsive and fails to respond to toe and tail pinch.</li>
	<li>Lay the rat in a supine position and make an incision through the abdominal wall using a pair of Iris scissors.</li>
	<li>Cut up through the bottom of the rib cage to the collarbone on both sides of the rib cage using Mayo scissors.</li>
	<li>Affix the sternum and ribs away from the heart and lungs with a hemostat.</li>
	<li>Pierce the left ventricle with a 23 G needle attached to a perfusion pump and cut the right atrium with Iris scissors.</li>
	<li>Perform transcardial perfusion with approximately 75 mL of 100 mM (1x) PBS.</li>
	<li>Continue transcardial perfusion with approximately 100 mL of 4% paraformaldehyde (PFA) buffered in 1x PBS.</li>
	<li>Remove the brain from the skull using forceps.</li>
	<li>Place the brain in sagittal position and slice into 4 equal sections (approximately 3 mm in width) using a razor blade and a rat brain matrix.</li>
	<li>Fix each section in 4% PFA in 1x PBS in a sealed, 5mL plastic tube overnight at 4 &#176;C on a shaker.</li>
	<li>Continue to fix in 4% PFA in 1x PBS at room temperature (RT) on a shaker for 1 h.</li>
	<li>Wash each section with 1x PBS at RT for 30 min, 3 times.</li>
</ol>
3. Dehydration and depigmentation
<ol>
	<li>Incubate the sample in a 20% methanol/80% deionized water solution for 1 h at RT.</li>
	<li>Incubate the sample in a 40% methanol/60% deionized water solution for 1 h at RT.</li>
	<li>Incubate the sample in a 60% methanol/40% deionized water solution for 1 h at RT.</li>
	<li>Incubate the sample in an 80% methanol/20% deionized water solution for 1 h at RT.</li>
	<li>Incubate the sample in 100% methanol for 1 h at RT. Repeat with more 100% methanol, incubating for 1 h at RT.</li>
	<li>Chill the sample in 100% methanol at 4&#176; C for approximately 10 min.</li>
	<li>Prepare a 66% dichloromethane (DCM)/33% methanol solution.</li>
	<li>Incubate the sample overnight in the DCM/methanol solution at RT, with shaking.</li>
	<li>Wash with methanol at RT for 30 min, 2 times.</li>
	<li>Chill the sample in 100% methanol at 4&#176; C for approximately 10 min.</li>
	<li>Bleach in chilled, freshly prepared 5% hydrogen peroxide in methanol overnight at 4&#176; C.</li>
	<li>Incubate the sample in an 80% methanol/20% deionized water solution for 1 h at RT.</li>
	<li>Incubate the sample in a 60% methanol/40% deionized water solution for 1 h at RT.</li>
	<li>Incubate the sample in a 40% methanol/60% deionized water solution for 1 h at RT.</li>
	<li>Incubate the sample in a 20% methanol/80% deionized water solution for 1 h at RT.</li>
	<li>Incubate the sample in 1x PBS for 1 h at RT.</li>
	<li>Wash in Solution 1 for 1 h at RT, 2 times.</li>
</ol>
4. Cholera Toxin Subunit B labeling
<ol>
	<li>Insert needle tip of 1 mL syringe into nucleus accumbens site.</li>
	<li>Slowly inject 2.5 &#956;L of 1% biotin-CTB over 20 s.</li>
	<li>Leave the needle tip in place for 1 min and remove slowly over 10 s to prevent leakage.</li>
</ol>
5. Antibody application
<ol>
	<li>Incubate the sample in Solution 3 for 2 days at 37 &#176;C in a water bath.</li>
	<li>Incubate the sample in Solution 4 for 2 days at 37 &#176;C in a water bath.</li>
	<li>Incubate with primary antibody for 7 days at 37 &#176;C in a water bath.</li>
	<li>Wash in Solution 2 for 5 times, incubating 1 h per wash. Store in Solution 2 at RT overnight.</li>
	<li>Incubate with secondary antibody for 7 days at 37 &#176;C in water bath.</li>
	<li>Wash in Solution 2 for 5 times, incubating 1 h per wash; store in Solution 2 at RT overnight. 
	NOTE: All steps from 5.6 onwards, including final storage, should occur in low light. The sample tubes can be shielded with aluminum foil. The concentrations used for the antibodies in this protocol were: TH at 1:100, Iba1 at 1:200, and the secondary antibody at 1:100. Add additional Solution 2 to steps 5.1-5.6 to ensure tubes are completely filled.</li>
</ol>
6. Tissue clearing
<ol>
	<li>Incubate the sample in a 20% methanol/80% deionized water solution for 1 h at RT.</li>
	<li>Incubate the sample in a 40% methanol/60% deionized water solution for 1 h at RT.</li>
	<li>Incubate the sample in a 60% methanol/40% deionized water solution for 1 h at RT.</li>
	<li>Incubate the sample in an 80% methanol/20% deionized water solution for 1 h at RT.</li>
	<li>Incubate the sample in 100% methanol for 1 h at RT.</li>
	<li>Incubate in fresh 100% methanol overnight at RT.</li>
	<li>Incubate in 66% DCM/33% methanol for 3 h, with shaking, at room temperature.</li>
	<li>Incubate in dibenzyl ether (DBE) without shaking. Leave the sample in DBE until clear and store in DBE until imaging.</li>
</ol>
7. Mounting
<ol>
	<li>Obtain a 3D printed chamber made of Visijet M3 Crystal resin (templates available on the idisco.info website).</li>
	<li>Secure the chamber to a microscope slide using an epoxy.</li>
	<li>Place the sample in the square space in the middle of the chamber.</li>
	<li>Fill the chamber completely with DBE.</li>
	<li>Place a 0.17 mm thick coverslip over the chamber and apply pressure until the coverslip has full contact with the chamber walls.</li>
	<li>Seal the edges of the coverslip to the chamber using the epoxy.</li>
	<li>Rotate the chamber to allow air bubbles to escape from the filling inlet.</li>
	<li>Fill the chamber with additional DBE.</li>
	<li>Seal the filling inlet with epoxy and allow to cure. 
	NOTE: Excess DBE will spill out during mounting.</li>
</ol>
8. Imaging
<ol>
	<li>Use a confocal microscope system to perform z-scan at 4x magnification (see the attached 3D video).</li>
	<li>Achieve fluorescent excitation of the sample using a laser set to emit at a wavelength similar to the wavelength specified by the secondary antibody. TH excitation was achieved utilizing a HeNe laser set to emit at 632 nm.</li>
	<li>Set the detector to a wavelength close to the laser emission. For the present protocol, the detector for TH was set to 650 nm.</li>
	<li>Raise the gain of the detector until there is a visible signal. The 650LP gain for images was set to 7.50 B.</li>
	<li>Using the Scan option in the confocal imaging software, move the microscope stage until the sample is brought into focus.</li>
	<li>Navigate around the tissue in all plains and obtain either single images or z-stack images using the confocal imaging software.</li>
	<li>Once imaging is completed, remove the sample from the chamber. Place it back into a tube completely filled with DBE and store at RT in a dark place, preferably covered in aluminum foil. 
	NOTE: Samples can be remounted and imaged as long as the florescence signal is still strong.</li>
</ol>

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R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tissue+clearing+and+its+applications+in+neuroscience.">Tissue clearing and its applications in neuroscience.</a> Nature Reviews Neuroscienc. 21, 298 (2020).</li><li>Renier, N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=iDISCO:+a+simple,+rapid+method+to+immunolabel+large+tissue+samples+for+volume+imaging.">iDISCO: a simple, rapid method to immunolabel large tissue samples for volume imaging.</a> Cell. 159 (4), 896-910 (2014).</li><li>Renier, N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mapping+of+brain+activity+by+automated+volume+analysis+of+immediate+early+genes.">Mapping of brain activity by automated volume analysis of immediate early genes.</a> Cell. 165 (7), 1789-1802 (2016).</li><li>Reid, W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+HIV-1+transgenic+rat+that+develops+HIV-related+pathology+and+immunologic+dysfunction.">An HIV-1 transgenic rat that develops HIV-related pathology and immunologic dysfunction.</a> Proceedings of the National Academy of Science U. 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M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=HIV-1+proteins+dysregulate+motivational+processes+and+dopamine+circuitry.">HIV-1 proteins dysregulate motivational processes and dopamine circuitry.</a> Science Report. 8, 6152 (2018).</li><li>Denton, A. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Selective+monoaminergic+and+histaminergic+circuit+dysregulation+following+long-term+HIV-1+protein+exposure.">Selective monoaminergic and histaminergic circuit dysregulation following long-term HIV-1 protein exposure.</a> Journal of neurovirology. 25 (4), 540-550 (2019).</li><li>Berendse, H. W., Groenewegen, H. J., Lohman, A. 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None of the authors have conflicts of interest to declare.

Tissue clearing offers a solution to the limitations of traditional IHC protocol. A sample that is transparent minimizes the scattering and absorption of light, which provides cellular level optical access to intact tissues18,19. Tissue clearing techniques turn a tissue transparent through delipidation, decolorization, and decalcification with the administration of solvents. Once the refractive index of the tissue sample matches the refractive index of the chosen...

Several tissue clearing techniques have been validated for the use in the brain: hydrogel, hydrophilic, and hydrophobic. These techniques aim to turn a tissue transparent through delipidation, decolorization, and decalcification via the administration of solvents. Once the refractive index of the tissue sample matches the refractive index of the chosen imaging medium, a clear image of the sample can be obtained. Hydrogel based techniques, such as CLARITY, secure biomolecules in the tissue by linking them to acryl-based hydrogels, which prevents structural damage and loss of proteins1. However, hydrogel techniques utilize harsh chemicals than have the potential to damage more fragile tissues, and denser tissue samples may not be compatible with hydrogel protocol. Certain hydrogel techniques may require expensive equipment or lead to tissue expansion. Hydrophilic techniques, like CUBIC, preserve 3D structure through the formation of hydrogen bonds within the tissue2. Tissue expansion can also occur in certain hydrophilic protocol. The clearing ability of hydrophilic techniques often does not match the ability that hydrophobic techniques have, which is important for denser and thicker tissues3.
Hydrophobic techniques are usually fast, do not require special equipment, and produce a sample that is easy to handle and store. iDISCO is a hydrophobic technique that eliminates the shrinkage that can occur in other hydrophobic protocol. Originally, the iDISCO tissue clearing technique was described by Renier et al.4 for the embryos and dense, adult organs of mice. This technique removes water from the tissue in the initial dehydration step, thereby reducing the light scatter. Tissue is permeabilized during pretreatment to allow deep antibody penetration. Alexa Fluor dyes in the far-red spectrum are used for immunolabeling to avoid autofluorescence of the tissue at lower wavelengths5. Tissue that has undergone hydrophobic clearing protocols are able to be handled easily and can be imaged several times due to the longevity of the Alexa Fluor dyes. If properly stored, tissues can provide images for months to a year after initial processing.
In the present protocol, tyrosine hydroxylase (TH) antibody, Iba1 antibody, and Cholera Toxin Subunit B (CTB) were used on both male and female control and HIV-1 transgenic (Tg) rat brains. The HIV-1 Tg rat possesses seven of the nine genes that comprise the HIV-1 viral genome, which leads to a non-infectious model of long-term HIV-1 protein exposure6,7,8. Dopaminergic alterations have previously been demonstrated in the HIV-1 Tg rat, and HIV-1 itself is an inflammatory disease, so the antibody choice was relevant to the experimental design9,10,11,12. CTB is a tracer that attaches to neurons via ganglioside binding and can be used to trace afferent projections in the brain. CTB has been previously used to study projections from the nucleus accumbens to the substantia nigra area, two areas of the brain heavily involved with dopaminergic pathways13,14,15.
In this protocol, TH will serve as a marker for dopamine production, and Iba1 will mark activated microglia. The TH antibody used selectively labels a single band at approximately 62 kDa that corresponds to tyrosine hydroxylase. The Iba1 antibody corresponds to the Iba1 carboxy-terminal sequence. Both antibodies were raised in rabbit and were polyclonal. CTB will be used for retrograde tracing of neurons from the nucleus accumbens area to the substantia nigra area. The current protocol also offers a guideline for adjustment of the original iDISCO protocol in two distinct ways: 1) overall reduction of the background staining by changing the serum reagents in the blocking steps, and 2) scaling of the incubation time to be suitable for larger tissue samples. Overall, the present protocol provides continued evidence that hydrophobic clearing techniques are feasible in brain tissues of the rat, have no discernible detrimental interactions with HIV-1 viral proteins, and are compatible with TH antibody, Iba1 antibody, and CTB.

<table><tbody><tr><td>Cholera Toxin Subunit B (Recombinant), Alexa Fluor 488</td><td>Invitrogen</td><td>C34775</td><td></td></tr><tr><td>DBE</td><td>Sigma-Aldrich</td><td>108014-1KG</td><td></td></tr><tr><td>DCM</td><td>Sigma-Aldrich</td><td>270997-100mL</td><td></td></tr><tr><td>DMSO</td><td>Sigma-Aldrich</td><td>472301-1L</td><td></td></tr><tr><td>Glycine</td><td>Fisher Chemical</td><td>G46-500</td><td></td></tr><tr><td>Goat anti-rabbit Alexa Fluor Plus 647</td><td>Invitrogen</td><td>A32733</td><td></td></tr><tr><td>Goat serum</td><td>Sigma Life Science</td><td>G9023-10mL</td><td></td></tr><tr><td>Heparin</td><td>Acros Organics</td><td>41121-0010</td><td></td></tr><tr><td>Iba1 primary antibody</td><td>FUJIFILM Wako</td><td>019-19741</td><td></td></tr><tr><td>Kwik-sil epoxy</td><td>VWR</td><td>70730-062</td><td></td></tr><tr><td>Methanol</td><td>Sigma-Aldrich</td><td>34860-1l-R</td><td></td></tr><tr><td>PBS</td><td>Fisher Bioreagents</td><td>BP2944-100</td><td></td></tr><tr><td>Perfusion machine</td><td>VWR</td><td>70730-062</td><td>mini pump variable flow</td></tr><tr><td>PFA</td><td>Sigma-Aldrich</td><td>158127-3KG</td><td></td></tr><tr><td>TH primary antibody</td><td>Millipore Sigma</td><td>AB152</td><td></td></tr><tr><td>TritonX-100</td><td>Fisher Bioreagents</td><td>BP151-500</td><td></td></tr><tr><td>Tween-20</td><td>Fisher Bioreagents</td><td>BP337-500</td><td></td></tr></tbody></table>

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.

Full clearing of large sections of both F344/N and HIV-1 rat brain tissue was achieved using this modified hydrophobic tissue clearing protocol. Figure 1 displays a typical confocal image for TH in the substantia nigra region. Figure 1A represents dense, positive staining. Dense areas such as these can be parsed out by focusing through the &#8220;Z&#8221; plane to confirm positive staining and proper cell morphology. Figure 1B repre...

Watch this Scientific Journal Video about A Hydrophobic Tissue Clearing Method for Rat Brain Tissue at JoVE.com

A Hydrophobic Tissue Clearing Method for Rat Brain Tissue

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 ...

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3.9K Views.  University of South Carolina. 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.

Video: A Hydrophobic Tissue Clearing Method for Rat Brain Tissue

2-Vessel Occlusion/Hypotension: A Rat Model of Global Brain Ischemia

Cardiac arrest followed by resuscitation often results in dramatic brain damage caused by ischemia and subsequent reperfusion of the brain. Global brain ischemia produces damage to specific brain regions shown to be highly sensitive to ischemia 1. Hippocampal neurons have higher sensitivity to ischemic insults compared to other cell populations, and specifically, the CA1 region of the hippocampus is particularly vulnerable to ischemia/reperfusion 2.
The design of therapeutic interventions, or study of mechanisms involved in cerebral damage, requires a model that produces damage similar to the clinical condition and in a reproducible manner. Bilateral carotid vessel occlusion with hypotension (2VOH) is a model that produces reversible forebrain ischemia, emulating the cerebral events that can occur during cardiac arrest and resuscitation. We describe a model modified from Smith et al. (1984) 2, as first presented in its current form in Sanderson, et al. (2008) 3, which produces reproducible injury to selectively vulnerable brain regions 3-6. The reliability of this model is dictated by precise control of systemic blood pressure during applied hypotension, the duration of ischemia, close temperature control, a specific anesthesia regimen, and diligent post-operative care. An 8-minute ischemic insult produces cell death of CA1 hippocampal neurons that progresses over the course of 6 to 24 hr of reperfusion, while less vulnerable brain regions are spared. This progressive cell death is easily quantified after 7-14 days of reperfusion, as a near complete loss of CA1 neurons is evident at this time.
In addition to this brain injury model, we present a method for CA1 damage quantification using a simple, yet thorough, methodology. Importantly, quantification can be accomplished using a simple camera-mounted microscope, and a free ImageJ (NIH) software plugin, obviating the need for cost-prohibitive stereology software programs and a motorized microscopic stage for damage assessment.

Bilateral carotid occlusion coupled with systemic hypotension produces global brain ischemia in the rat, resulting in damage to the hippocampus with reproducible severity. Animal subjects are impaired with predictable patterns of brain damage, they recover expediently, and mortality rates are comparatively low.

Cardiac arrest followed by resuscitation often results in dramatic brain damage caused by ischemia and subsequent reperfusion of the brain. Global brain ...

Medicine

Metabolic Support of Excised, Living Brain Tissues During Magnetic Resonance Microscopy Acquisition

This protocol describes the procedures necessary to support normal metabolic functions of acute brain slice preparations during the collection of magnetic resonance (MR) microscopy data. While it is possible to perform MR collections on living, excised mammalian tissue, such experiments have traditionally been constrained by resolution limits and are thus incapable of visualizing tissue microstructure. Conversely, MR protocols that did achieve microscopic image resolution required the use of fixed samples to accommodate the need for static, unchanging conditions over lengthy scan times. The current protocol describes the first available MR technique that enables imaging of living, mammalian tissue samples at microscopic resolutions. Such data is of great importance to the understanding of how pathology-based contrast changes occurring at the microscopic level influence the content of macroscopic MR scans such as those used in the clinic. Once such an understanding is realized, diagnostic methods with greater sensitivity and accuracy can be developed, which will translate directly to earlier disease treatment, more accurate therapy monitoring and improved patient outcomes.
While the described methodology focuses on brain slice preparations, the protocol is adaptable to any excised tissue slice given that changes are made to the gas and perfusate preparations to accommodate the tissue&#39;s specific metabolic needs. Successful execution of the protocol should result in living, acute slice preparations that exhibit MR diffusion signal stability for periods up to 15.5 h. The primary advantages of the current system over other MR compatible perfusion apparatuses are its compatibility with the MR microscopy hardware required to attain higher resolution images and ability to provide constant, uninterrupted flow with carefully regulated perfusate conditions. Reduced sample throughput is a consideration with this design as only one tissue slice may be imaged at a time.

The current protocol describes a method by which users can maintain viability of acute hippocampal and cortical slice preparations during the collection of magnetic resonance microscopy data.

This protocol describes the procedures necessary to support normal metabolic functions of acute brain slice preparations during the collection of magnetic ...

Bioengineering

Sublimation of DAN Matrix for the Detection and Visualization of Gangliosides in Rat Brain Tissue for MALDI Imaging Mass Spectrometry

Sample preparation is key for optimal detection and visualization of analytes in Matrix-assisted Laser Desorption/Ionization (MALDI) Imaging Mass Spectrometry (IMS) experiments. Determining the appropriate protocol to follow throughout the sample preparation process can be difficult as each step must be optimized to comply with the unique characteristics of the analytes of interest. This process involves not only finding a compatible matrix that can desorb and ionize the molecules of interest efficiently, but also selecting the appropriate matrix deposition technique. For example, a wet matrix deposition technique, which entails dissolving a matrix in solvent, is superior for desorption of most proteins and peptides, whereas dry matrix deposition techniques are particularly effective for ionization of lipids. Sublimation has been reported as a highly efficient method of dry matrix deposition for the detection of lipids in tissue by MALDI IMS due to the homogeneity of matrix crystal deposition and minimal analyte delocalization as compared to many wet deposition methods 1,2. Broadly, it involves placing a sample and powdered matrix in a vacuum-sealed chamber with the samples pressed against a cold surface. The apparatus is then lowered into a heated bath (sand or oil), resulting in sublimation of the powdered matrix onto the cooled tissue sample surface. Here we describe a sublimation protocol using 1,5-diaminonaphthalene (DAN) matrix for the detection and visualization of gangliosides in the rat brain using MALDI IMS.

A protocol for the sublimation of DAN matrix onto rat brain tissue for the detection of gangliosides using MALDI Imaging Mass Spectrometry is presented.

Sample preparation is key for optimal detection and visualization of analytes in Matrix-assisted Laser Desorption/Ionization (MALDI) Imaging Mass ...

A Micro-CT-based Method for Characterizing Lesions and Locating Electrodes in Small Animal Brains

Lesion and electrode location verification are traditionally done via histological examination of stained brain slices, a time-consuming procedure that requires manual estimation. Here, we describe a simple, straightforward method for quantifying lesions and locating electrodes in the brain that is less laborious and yields more detailed results. Whole brains are stained with osmium tetroxide, embedded in resin, and imaged with a micro-CT scanner. The scans result in 3D digital volumes of the brains with resolutions and virtual section thicknesses dependent on the sample size (12&#8211;15 and 5&#8211;6 &#181;m&#160;per voxel for rat and zebra finch brains, respectively). Surface and deep lesions can be characterized, and single tetrodes, tetrode arrays, electrolytic lesions, and silicon probes can also be localized. Free and proprietary software allows experimenters to examine the sample volume from any plane and segment the volume manually or automatically. Because this method generates whole brain volume, lesions and electrodes can be quantified to a much higher degree than in current methods, which will help standardize comparisons within and across studies.

This article describes a straightforward method to prepare small animal brains for micro-CT imaging, in which lesions can be quantified and electrodes located with high precision in the context of the whole brain.

Lesion and electrode location verification are traditionally done via histological examination of stained brain slices, a time-consuming procedure that ...

Massive Pontine Hemorrhage by Dual Injection of Autologous Blood

We provide a protocol to establish a massive pontine hemorrhage model in a rat. Rats weighing about 250 grams were used in this study. One hundred microliters of autologous blood was taken from the tail vein and stereotaxically injected into the pons. The injection process was divided into 2 steps: First, 10 &#181;L of blood was injected into a specific location, anteroposterior position (AP) -9.0 mm; lateral (Lat) 0 mm; vertical (Vert) -9.2 mm, followed by a second injection of the residual blood located at AP -9.0 mm; Lat 0 mm; Vert -9.0 mm with a 20-minute interval. The balance beam test, limb placement test, and the modified Voestch neuroscore were used to evaluate neurological function. Magnetic Resonance Imaging (MRI) was used to assess the volume of hemorrhage in vivo. The symptoms of this model were in line with patients with massive pontine hemorrhage.

We present a protocol to establish a massive pontine hemorrhage model in a rat via dual injection of autologous blood.

We provide a protocol to establish a massive pontine hemorrhage model in a rat. Rats weighing about 250 grams were used in this study. One hundred ...

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 ...

Investigation of Spatial Interaction Between Astrocytes and Neurons in Cleared Brains

Combining viral vector transduction and tissue clearing using the CLARITY method makes it possible to simultaneously investigate several types of brain cells and their interactions. Viral vector transduction enables the marking of diverse cell types in different fluorescence colors within the same tissue. Cells can be identified genetically by activity or projection. Using a modified CLARITY protocol, the potential sample size of astrocytes and neurons has grown by 2-3 orders of magnitude. The use of CLARITY allows the imaging of complete astrocytes, which are too large to fit in their entirety in slices, and the examination of the somata with all their processes. In addition, it provides the opportunity to investigate the spatial interaction between astrocytes and different neuronal cell types, namely, the number of pyramidal neurons in each astrocytic domain or the proximity between astrocytes and specific inhibitory neuron populations. This paper describes, in detail, how these methods are to be applied.

Combining viral vector transduction and brain clearing using the CLARITY method allows the investigation of a large number of neurons and astrocytes simultaneously.

Combining viral vector transduction and tissue clearing using the CLARITY method makes it possible to simultaneously investigate several types of brain ...

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.

Neuroscience research focused on the central nervous system necessitates a thorough understanding of various factors, including cell distribution, ...

A Hydrophobic Tissue Clearing Method for a Rat Brain Tissue Sample

<a href='https://app.jove.com/v/61821/a-hydrophobic-tissue-clearing-method-for-rat-brain-tissue'>Source: Kirchner, K.N., et al.,&#160;A Hydrophobic Tissue Clearing Method for Rat Brain Tissue. J. Vis. Exp.&#160;(2020).</a>This video demonstrates the method of hydrophobic tissue clearing of a rat brain tissue sample. The sample is sequentially dehydrated in methanol, treated with a dichloromethane and methanol mixture to dissolve lipids and pigments, and transferred to dibenzyl ether for complete tissue clearing, enabling detailed imaging without thin sectioning.

Source: Kirchner, K.N., et al.,&#160;A Hydrophobic Tissue Clearing Method for Rat Brain Tissue. J. Vis. Exp.&#160;(2020).This video demonstrates the ...

JoVE Encyclopedia of Experiments

Neuropathology

Neurodegenerative Diseases

A Method for the Isolation of Neurovascular Units from Rat Brain

Source: Brzica, H., et al,. <a href="https://app.jove.com/v/57698">A simple and reproducible method to prepare membrane samples from freshly isolated rat brain microvessels.</a> J. Vis. Exp. (2018).
This video demonstrates the process of isolating brain microvessels from euthanized rats. It includes the dissection of the brain, cleaning of the tissue, and mechanical digestion, followed by repeated centrifugation to isolate the pure fraction of microvessels.

All procedures involving sample collection have been performed in accordance with the institute&#39;s IRB guidelines. The procedural flow for the protocol ...