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. S. A. 98, 9271-9276 (2001).</li><li>Vigorito, M., Connaghan, K. P., Chang, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+HIV-1+transgenic+rat+model+of+neuroHIV.">The HIV-1 transgenic rat model of neuroHIV.</a> Brain, Behavior, and Immunity. 48, 336-349 (2015).</li><li>McLaurin, K. A., 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=Synaptic+connectivity+in+medium+spiny+neurons+of+the+nucleus+accumbens:+A+sex-dependent+mechanism+underlying+apathy+in+the+HIV-1+transgenic+rat.">Synaptic connectivity in medium spiny neurons of the nucleus accumbens: A sex-dependent mechanism underlying apathy in the HIV-1 transgenic rat.</a> Frontiers in Behavior Neurosciences. 12, 285 (2018).</li><li>Fitting, S., Booze, R. M., Mactutus, C. 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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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3.8K 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

Preparation of Mouse Brain Tissue for Immunoelectron Microscopy

Transmission electron microscopy (TEM) is extremely useful for visualizing microglial, oligodendrocytic, astrocytic, and neuronal subcellular compartments (dendrite, dendritic spine, axon, axon terminal, perikaryon), as well as their intracellular organelles and cytoskeleton, in the central nervous system at high spatial resolution. Combined with TEM, pre-embedding immunocytochemistry allows the discrimination of cellular elements with few distinctive features and identification criteria (e.g., microglial perikarya and processes, when using an antibody against the microglia-specific marker Iba1 (ionized calcium binding adaptor molecule 1; as presented here)), identifying the neurotransmitter contents of cellular elements (e.g., serotonergic) and their ultrastructural localization of soluble or membrane-bound proteins (e.g., 5 HT1A and EphA4 receptors). Here, we describe a protocol for transcardiac perfusion of mice with acrolein fixative, removal and sectioning of the brain, as well as immunoperoxidase-diaminobenzidine (DAB) staining, resin embedding, and ultrathin sectioning of the brain sections. Upon completion of these procedures, the immunostained material is ready for examination with TEM. When rigorously performed, this technique provides an excellent compromise between optimal ultrastructural preservation and immunocytochemical detection.

We describe a protocol for transcardiac perfusion of mice, removal and sectioning of the brain, as well as immunoperoxidase staining, resin embedding, and ultrathin sectioning of the brain sections. Upon completion of these procedures, the immunostained material is ready for examination with transmission electron microscopy.

Transmission electron microscopy (TEM) is extremely useful for visualizing microglial, oligodendrocytic, astrocytic, and neuronal subcellular compartments ...

Chromatin Immunoprecipitation from Dorsal Root Ganglia Tissue following Axonal Injury

Axons in the central nervous system (CNS) do not regenerate while those in the peripheral nervous system (PNS) do regenerate 
to a limited extent after injury (Teng et al., 2006). It is recognized that transcriptional programs essential for neurite and axonal outgrowth are 
reactivated upon injury in the PNS (Makwana et al., 2005). However the tools available to analyze neuronal gene regulation in vivo are limited and 
often challenging.
The dorsal root ganglia (DRG) offer an excellent injury model system because both the CNS and PNS are innervated by a 
bifurcated axon originating from the same soma. The ganglia represent a discrete collection of cell bodies where all transcriptional events occur, 
and thus provide a clearly defined region of transcriptional activity that can be easily and reproducibly removed from the animal. Injury of nerve 
fibers in the PNS (e.g. sciatic nerve), where axonal regeneration does occur, should reveal a set of transcriptional programs that are distinct from 
those responding to a similar injury in the CNS, where regeneration does not take place (e.g. spinal cord). Sites for transcription factor binding, 
histone and DNA modification resulting from injury to either PNS or CNS can be characterized using chromatin immunoprecipitation (ChIP). 
Here, we describe a ChIP protocol using fixed mouse DRG tissue following axonal injury. This powerful combination provides a means for characterizing the pro-regeneration chromatin environment necessary for promoting axonal regeneration.

We present a method for chromatin immunoprecipitation from dorsal root ganglia tissue following axonal injury. The approach can be used to identify specific transcription factor binding sites and epigenetic modification of histone and DNA important for the regeneration of injured axons in both the peripheral and central nervous system.

Axons in the central nervous system (CNS) do not regenerate while those in the peripheral nervous system (PNS) do regenerate 
to a limited extent after ...

Intact Histological Characterization of Brain-implanted Microdevices and Surrounding Tissue

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

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

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

Immunohistochemical Analysis in the Rat Central Nervous System and Peripheral Lymph Node Tissue Sections

Immunohistochemistry (IHC) provides highly specific, reliable and attractive protein visualization. Correct performance and interpretation of an IHC-based multicolor labeling is challenging, especially when utilized for assessing interrelations between target proteins in the tissue with a high fat content such as the central nervous system (CNS).
Our protocol represents a refinement of the standard immunolabeling technique particularly adjusted for detection of both structural and soluble proteins in the rat CNS and peripheral lymph nodes (LN) affected by neuroinflammation. Nonetheless, with or without further modifications, our protocol could likely be used for detection of other related protein targets, even in other organs and species than here presented.

We here present an optimized, detailed protocol for double immunostaining in formalin-fixed, paraffin-embedded rat central nervous system (CNS) and peripheral lymph node (LN) tissue sections.

Immunohistochemistry (IHC) provides highly specific, reliable and attractive protein visualization. Correct performance and interpretation of an IHC-based ...

Primer for Immunohistochemistry on Cryosectioned Rat Brain Tissue: Example Staining for Microglia and Neurons

Immunohistochemistry is a widely used technique for detecting the presence, location, and relative abundance of antigens in situ. This introductory level protocol describes the reagents, equipment, and techniques required to complete immunohistochemical staining of rodent brain tissue, using markers for microglia and neuronal elements as an example. Specifically, this paper is a step-by-step protocol for fluorescent visualization of microglia and neurons via immunohistochemistry for Iba1 and Pan-neuronal, respectively. Fluorescence double-labeling is particularly useful for the localization of multiple proteins within the same sample, providing the opportunity to accurately observe interactions between cell types, receptors, ligands, and/or the extracellular matrix in relation to one another as well as protein co-localization within a single cell. Unlike other visualization techniques, fluorescence immunohistochemistry staining intensity may decrease in the weeks to months following staining, unless appropriate precautions are taken. Despite this limitation, in many applications fluorescence double-labeling is preferred over alternatives such as 3,3&#39;-diaminobenzidine tetrahydrochloride (DAB) or alkaline phosphatase (AP), as fluorescence is more time efficient and allows for more precise differentiation between two or more markers. The discussion includes troubleshooting tips and advice to promote success.

This introductory level protocol describes the reagents, equipment, and techniques required to complete immunohistochemical staining of rodent brains, using markers for microglia and neuronal elements as an example.

Immunohistochemistry is a widely used technique for detecting the presence, location, and relative abundance of antigens in situ. This introductory level ...

Using Fluorescence Activated Cell Sorting to Examine Cell-Type-Specific Gene Expression in Rat Brain Tissue

The brain is comprised of four primary cell types including neurons, astrocytes, microglia and oligodendrocytes. Though they are not the most abundant cell type in the brain, neurons are the most widely studied of these cell types given their direct role in impacting behaviors. Other cell types in the brain also impact neuronal function and behavior via the signaling molecules they produce. Neuroscientists must understand the interactions between the cell types in the brain to better understand how these interactions impact neural function and disease. To date, the most common method of analyzing protein or gene expression utilizes the homogenization of whole tissue samples, usually with blood, and without regard for cell type. This approach is an informative approach for examining general changes in gene or protein expression that may influence neural function and behavior; however, this method of analysis does not lend itself to a greater understanding of cell-type-specific gene expression and the effect of cell-to-cell communication on neural function. Analysis of behavioral epigenetics has been an area of growing focus which examines how modifications of the deoxyribonucleic acid (DNA) structure impact long-term gene expression and behavior; however, this information may only be relevant if analyzed in a cell-type-specific manner given the differential lineage and thus epigenetic markers that may be present on certain genes of individual neural cell types. The Fluorescence Activated Cell Sorting (FACS) technique described below provides a simple and effective way to isolate individual neural cells for the subsequent analysis of gene expression, protein expression, or epigenetic modifications of DNA. This technique can also be modified to isolate more specific neural cell types in the brain for subsequent cell-type-specific analysis.

The goal of this protocol is to use the fluorescence activated cell sorting (FACS) technique to sort specific types of neural cells for subsequent analysis of cell-type-specific gene expression, epigenetic markers, and or protein expression.

The brain is comprised of four primary cell types including neurons, astrocytes, microglia and oligodendrocytes. Though they are not the most abundant ...

A Rat Model of Mild Intrauterine Hypoperfusion with Microcoil Stenosis

Intrauterine hypoperfusion/ischemia is one of the major causes of intrauterine/fetal growth restriction, preterm birth, and low birth weight. Most studies of this phenomenon have been performed in either models with severe intrauterine ischemia or models with gradient degree of intrauterine hypoperfusion. No study has been performed in a model on uniform mild intrauterine hypoperfusion (MIUH). Two models have been used for studies of MIUH: a model based on suture ligation of either side of the arterial arcade formed with the uterine and ovarian arteries, and a transient model based on clipping the bilateral ovarian arteries and aorta having patency. Those two rodent models of MIUH have some limitations, e.g., not all fetuses are subjected to MIUH, depending on their position in the uterine horn. In our MIUH model, all fetuses are subjected to a comparable level of intrauterine hypoperfusion. MIUH was achieved by mild stenosis of all four arteries feeding the uterus, i.e., the bilateral uterine and ovarian arteries.
Arterial stenosis was induced by metal microcoils wrapped around the feeding arteries. Producing arterial stenosis with microcoils allowed us to control, optimize, and reproduce decreased blood flow with very little inter-animal variability and a low mortality rate, thus enabling accurate evaluation. When microcoils with an inner diameter of 0.24 mm were used, the blood flow in both the placenta and fetus was mildly decreased (approximately 30% from the pre-stenosis level in the placenta). The offspring of our MIUH model clearly demonstrates long-lasting alterations in neurological, neuroanatomical and behavioral test results.

Mild intrauterine hypoperfusion was produced by artery stenosis with metal microcoils wrapped around the uterine and ovarian arteries in rats at embryonic day 17. This procedure produced prenatal hypoperfusion and intrauterine growth restriction.

Intrauterine hypoperfusion/ischemia is one of the major causes of intrauterine/fetal growth restriction, preterm birth, and low birth weight. Most studies ...

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

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

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

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

Isolation of Cerebral Capillaries from Fresh Human Brain Tissue

Understanding blood-brain barrier function under physiological and pathophysiological conditions is critical for the development of new therapeutic strategies that hold the promise to enhance brain drug delivery, improve brain protection, and treat brain disorders. However, studying the human blood-brain barrier function is challenging. Thus, there is a critical need for appropriate models. In this regard, brain capillaries isolated from human brain tissue represent a unique tool to study barrier function as close to the human in vivo situation as possible. Here, we describe an optimized protocol to isolate capillaries from human brain tissue at a high yield and with consistent quality and purity. Capillaries are isolated from fresh human brain tissue using mechanical homogenization, density-gradient centrifugation, and filtration. After the isolation, the human brain capillaries can be used for various applications including leakage assays, live cell imaging, and immune-based assays to study protein expression and function, enzyme activity, or intracellular signaling. Isolated human brain capillaries are a unique model to elucidate the regulation of the human blood-brain barrier function. This model can provide insights into central nervous system (CNS) pathogenesis, which will help the development of therapeutic strategies for treating CNS disorders.

Isolated brain capillaries from human brain tissue can be used as a preclinical model to study barrier function under physiological and pathophysiological conditions. Here, we present an optimized protocol to isolate brain capillaries from fresh human brain tissue.

Understanding blood-brain barrier function under physiological and pathophysiological conditions is critical for the development of new therapeutic ...

Preparation of Newborn Rat Brain Tissue for Ultrastructural Morphometric Analysis of Synaptic Vesicle Distribution at Nerve Terminals

Our laboratory and many others have exploited the high resolving power of transmission electron microscopy to study the morphology and spatial organization of synaptic vesicles. In order to obtain high-quality electron micrographs that can yield the degree of morphological detail necessary for quantitative analysis of pre-synaptic vesicle distribution, optimal specimen preparation is critical. Chemical fixation is the first step in the process of specimen preparation, and of utmost importance to preserve fine ultrastructure. Vascular fixation with a glutaraldehyde-formaldehyde solution, followed by treatment of vibratome-sectioned specimens with osmium tetroxide, stabilizes the maximum number of molecules, especially proteins and lipids, and results in superior conservation of ultrastructure. Tissue is then processed with counterstaining, sequential dehydration and resin-embedding. En bloc staining with uranyl acetate (i.e., staining of vibratome-sectioned tissue before resin embedding) enhances endogenous contrast and stabilizes cell components against extraction during specimen processing. Contrast can be further increased by applying uranyl acetate as a post-stain on ultrathin sections. Double-staining of ultrathin sections with lead citrate after uranyl acetate treatment also improves image resolution, by intensifying electron-opacity of nucleic acid-containing structures through selective binding of lead to uranyl acetate. Transmission electron microscopy is a powerful tool for characterization of the morphological details of synaptic vesicles and quantification of their size and spatial organization in the terminal bouton. However, because it uses fixed tissue, transmission electron microscopy can only provide indirect information regarding living or evolving processes. Therefore, other techniques should be considered when the main objective is to study dynamic or functional aspects of synaptic vesicle trafficking and exocytosis.

We describe procedures for processing newborn rat brain tissue to obtain high-resolution electron micrographs for morphometric analysis of synaptic vesicle distribution at nerve terminals. The micrographs obtained with these methods can also be used to study the morphology of a number of other cellular components and their dimensional structural relationships.