This work was supported by the Brain Korea 21 PLUS Project for Medical Science, Yonsei University. In addition, this work was supported by a grant from the National Research Foundation of Korea (NRF-2017R1D1A1B03030315).

All procedures have been approved by the appropriate research ethics committee at Yonsei University College of Medicine. All experimental animals are sacrificed in accordance with the guidelines of the laboratory animal care committee at Yonsei University College of Medicine.
1. Preparation of Reagents
Caution: Paraformaldehyde (PFA), acrylamide and sodium dodecyl sulfate (SDS) are toxic irritants and thus should be handled in a fume hood with appropriate personal protective equipment (PPE; lab coat, gloves, protective eyewear).
<ol>
	<li>4% acrylamide (A.A) solution (A4P0): Add 20 mL of 40% acrylamide solution to 180 mL of 0.1 M phosphate-buffered saline (PBS).</li>
	<li>0.25% VA-044 solution: Add 0.5 g of 2,2&#180;-Azobis[2-(2-imidazolin-2-yl)propane] dihydrochloride (VA-044) powder to 200 mL of 0.1 M phosphate-buffered saline (PBS). While the VA-044 powder can be stored at room temperature, it must be stored at 4 &#176;C upon solubilization in PBS. For best results, prepare only the amount of solution needed for the experiment.</li>
	<li>PACT cocktail solution: Add 10 g of acrylamide to 225 mL of 4% paraformaldehyde (PFA) solution and adjust the volume to 200 mL with 4% PFA. Prior to use, add 100 mg of VA-044 powder to 40 mL of mixture in a 50 mL conical tube.</li>
	<li>PACT clearing solution buffer: Add 40 g of sodium dodecyl sulfate (SDS) to 350 mL of 0.1 M PBS and adjust the volume to 500 mL with 0.1 M PBS. For mPACT clearing solution, add 0.5% &#945;-thioglycerol to the PACT clearing solution buffer.</li>
	<li>The mice used in this study were 2-week-old BALB male mice; the rats used in this study were 2 week old male SD-rats.</li>
</ol>
2. Anesthesia and Perfusion Surgery
Caution: PFA and acrylamide are toxic irritants and thus should be handled in a fume hood with appropriate PPE.
<ol>
	<li>Anesthetize the animal in a pathogen-free room with 30 mg/kg of zoletil using a 1 mL syringe with a 26-gauge needle. Monitor the animal for 5 -10 min.</li>
	<li>Once the animal has reached a surgical plane of anesthesia, use the toe-pinch response method to determine the depth of anesthesia. Confirm unresponsiveness. 
	NOTE: The following steps involving mouse and rat surgery follow a similar protocol used in a previous study11.</li>
	<li>Make a 5-6 cm incision beneath the rib cage, through the integument and abdominal wall.</li>
	<li>Use curved, blunt scissors to make an incision in the diaphragm, palpating the thoracic area to locate the site of incision beforehand.</li>
	<li>Extend the incision across the entire length of the ribcage to expose the pleural cavity.</li>
	<li>Carefully displace the lungs. Make a cut through the ribcage up to the collarbone. Using iris scissors, make a small incision to the posterior end of the left ventricle. Insert an 18-gauge blunt- or olive-tipped perfusion needle into the ascending aorta, passing through the cut ventricle.</li>
	<li>Secure the needle and prevent leakage by clamping the heart with a hemostat. Alternatively, use a modified hemostat to clamp the aorta around the needle tip.</li>
	<li>Make a large incision in the right atrium, taking care to ensure minimal damage to the descending aorta. The rat is now ready for perfusion.</li>
</ol>
3. Whole Perfusion and Dissection of Rat
Caution: PFA and acrylamide are toxic irritants and thus should be handled in a fume hood with appropriate PPE.
NOTE: The following whole perfusion steps are a similar to a protocol used in a previous study by Woo et al. (2016)10,11.
<ol>
	<li>Attach the right atrium of heart to an 18-gauge needle, taking care to not introduce any air bubbles.</li>
	<li>Using a 50 mL syringe, quickly and evenly pump 50 mL of cold 0.1 M PBS solution containing heparin (10 units/mL).</li>
	<li>Connect the 18-gauge needle to the tube of the peristaltic pump.</li>
	<li>Wash with 200 mL of cold 0.1 M PBS solution containing heparin (10 units/mL) at a circulation velocity of 10 mL/min.</li>
	<li>Fix with 250 mL of cold 4% PFA solution at a circulation velocity of 10 mL/min. The rat should be stiff at this stage.</li>
	<li>Collect and store the remaining PFA solution for disposal.</li>
	<li>For PACT, perfuse tissues with a chilled PACT cocktail solution of 4% PFA, 4% acrylamide, and 0.25% VA-044 powder with an 18-gauge needle, and then remove the head and spine. Expose the skull by making a midline incision from the neck to the nose. This additional perfusion step is unnecessary for the psPACT and mPACT methods; after fixation, immediately isolate the head and spine and expose the skull.</li>
	<li>Expose the base of the skull by removing any residual neck and spine muscle.</li>
	<li>Use rongeurs and scissors to peel away the skull. Remove the brain and spinal cord.</li>
	<li>Store the tissue in 4% PFA at 4 &#176;C; tissues can be stored for up to 1 week.</li>
</ol>
4. Hydrogel Monomer Infusion and Polymerization of the Rat and Mouse CNS
Caution: Acrylamide, SDS, &#945;-thioglycerol and PFA are irritants and thus should be handled in a fume hood and with appropriate PEE.
<ol>
	<li>PACT (Passive Clearing Technique)
	<ol>
		<li>Isolate and culture the whole CNS (brain and spinal cord) from fixed mice and rats to a 50 mL tube containing a chilled PACT cocktail solution of 4% PFA, 4% acrylamide, and 0.25% VA-044 powder and store at 4 &#176;C for 24 h. Ensure that the tissue is completely immersed in the solution.</li>
		<li>Embed the sample in nitrogen gas for 10 min using a tissue gel hybridization system connected to a nitrogen tank: Set the system to 37 &#176;C. Transfer tissues to a 50 mL tube containing fresh cold (4 &#176;C) PACT cocktail solution and connect with tube cap, then turn on the vacuum.</li>
		<li>To polymerize the hydrogel, place the tube containing the sample in a shaking incubator (150 rpm, 37 &#176;C) for 3 h, or until polymerization is complete.</li>
		<li>Using blotting paper (see Table of Materials), remove the remaining polymerized hydrogel surrounding the tissues.</li>
		<li>Transfer the tissue to a 50 mL tube containing clearing solution (8% SDS in 0.1 M PBS, pH 8.0).</li>
		<li>Place the sample in a shaking incubator set to 37 &#176;C and 150 rpm until the tissue has been cleared. On average, it takes about 20 days for the mouse CNS to achieve full clarity.</li>
	</ol>
	</li>
	<li>psPACT (process separate Passive Clearing Technique)
	<ol>
		<li>Isolate the whole CNS (brain and spinal cord) with bone cutter and scissors from PFA-fixed mice and rats on a clean bench.</li>
		<li>Transfer tissues to a 50 mL tube containing 4% PFA, and store at 4 &#176;C for 24 h. Ensure that the tissue is completely immersed in the fixative.</li>
		<li>Wash the fixed tissue for 1 h in 0.1 M PBS, and then transfer to A4P0 solution (4% acrylamide in 0.1 M PBS). Store at 37 &#176;C for 24 h.</li>
		<li>Wash the tissue for 5 min in 0.1 M PBS.</li>
		<li>Immerse the tissue in 0.25% VA-044 in 0.1 M PBS. Store at 37 &#176;C for 6-24 h, and then transfer to fresh 0.25% VA-044/PBS solution.</li>
		<li>Embed the sample in nitrogen gas for 10 min using a tissue gel hybridization system (see Table of Materials) connected to a nitrogen tank: Transfer tissues to a 50 mL tube containing fresh cold (4 &#176;C) 0.25% VA-044/PBS solution, and connect with tube cap. Turn on the vacuum.</li>
		<li>Transfer the tissue to clearing solution (8% SDS in 0.1 M PBS, pH 8.0).</li>
		<li>Incubate the sample in a shaking incubator set to 37 &#176;C and 150 rpm until the tissue has been cleared. On average, it takes approximately 17 days for mouse CNS to achieve full clarity.</li>
	</ol>
	</li>
	<li>mPACT (modified Passive Clearing Technique)
	<ol>
		<li>Isolate the whole CNS (brain and spinal cord) with bone cutter and scissors from PFA-fixed mice and rats on a clean bench.</li>
		<li>Follow steps 4.2.2 - 4.2.8 of the psPACT protocol.</li>
		<li>Transfer the tissue to clearing solution. Note that unlike the PACT and psPACT protocols, mPACT requires a clearing solution consisting of 0.5% &#945;-thioglycerol in addition to 8% SDS in 0.1 M PBS, pH 8.0. &#945;-thioglycerol is an un-browning agent that helps clear tissue more rapidly and effectively. Ensure that the tissue is fully immersed in the solution.</li>
		<li>Place the sample in a shaking incubator set to 37 &#176;C and 150 rpm until the tissue has been cleared. On average, it takes approximately 2 weeks for mouse CNS to achieve full clarity.</li>
	</ol>
	</li>
</ol>
5. Refractive Index Matching and Immunostaining of Cleared CNS
NOTE: nRIMS (Nycodenz-based Refractive Index Matching Solution) consists of 0.8 g/mL Nycodenz powder dissolved in 30 mL base buffer (0.01% sodium azide and Tween-20 in 0.1 M PBS, pH 8.0). It is recommended that the solution is placed in a 37 &#176;C shaking incubator to allow for proper solvation of the powder.
<ol>
	<li>Incubate tissues in 0.1% Triton X-100 in 0.1 M PBS for 2 h, then block with 2% bovine serum albumin (BSA) in 0.1 M PBS for 6 h.</li>
	<li>Wash sections three times in PBST (0.1% Tween-20 in 0.1 M PBS) for 2 h. Stain sections with primary antibodies (in this case, an anti-rabbit PECAM-CD31 antibody, which stains blood vessels) for 2 days.</li>
	<li>Stain sections with secondary antibodies (in this case, a goat anti-rabbit IgG Cy3 fluorescent conjugate) in 2% BSA for 2 days.</li>
	<li>Wash labeled tissues three times in PBST for 2 h, and store in 15 mL nRIMS for 2 - 10 days.</li>
	<li>Before imaging, move labeled tissues to a small amount of nRIMS on 35- or 60-mm tissue culture dishes. Fix with silicone around the bottom edge of the dish.</li>
	<li>Add 1.5 - 2 mL of fresh nRIMS.</li>
</ol>
6. Image processing
<ol>
	<li>Acquire images of cleared tissues with tile scanning using a confocal laser-scanning microscopewith 10x magnification (see Table of Materials). For of Cy3-labeled tissue, use wavelengths of 550-600 nm.</li>
</ol>

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The authors have nothing to disclose.

While the passive, non-electrophoretic extraction methods employed in PACT significantly improved the consistency achieved with previous tissue clearing methods such as CLARITY2,3,4,7,8, the technique still bears several shortcomings, the most pressing of which is the length of time required to achieve maximal tissue clarity12. In the c...

A fundamental objective of scientific and clinical inquiry involves attaining a complete understanding of organ structure and function; however, the exceedingly complex nature of mammalian organs often serves as a barrier to fully achieving this aim1. CLARITY (Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tisssue-hYdrogel)2,3,4, which involves building an acrylamide-based hydrogel hybrid from intact tissues, achieves optical clearance of a variety of organs, including the brain, liver, and spleen, while preserving their structural integrity5. CLARITY has thus enabled not only visualization but also the opportunity to finely dissect complex cellular networks and tissue morphologies without the need for sectioning.
In order to achieve tissue clearance, CLARITY employs electrophoretic methods to remove the lipid content of the sample at hand. While CLARITY has been noted for producing physically stable tissue-hydrogel hybrids, studies have shown that its use of electrophoretic tissue clearing (ETC) methods yields variable results in terms of tissue quality, including browning, epitope damage, and protein loss5,6. To address these issues, modified protocols such as PACT (PAssive Clarity Technique), which replaces the ETC treatment with a passive, ionic-detergent based delipidation technique, have been developed7,8,9. Despite achieving a greater consistency in results, however, PACT requires more time to obtain maximal clearance. Furthermore, none of these techniques have yet been applied to the whole CNS form, or in larger rodent models such as rats and guinea pigs.
The present study seeks to address these limitations by proposing novel methodologies, psPACT (process-separate PACT) and mPACT (modified PACT), for facilitating the fast clearance of the whole CNS and internal organs in both mouse and rat models10. Specifically, psPACT processes tissues in 4% acrylamide and 0.25% VA-044 in two separate steps during hydrogel formation; mPACT essentially involves the same steps, but supplements the SDS-based clearing solution with 0.5% &#945;-thioglycerol as a key reagent. Both techniques harness the endogenous systemic and cerebrospinal circulatory systems to significantly reduce the time needed to produce optical clearance. As a proof of principle, we demonstrate the use of confocal microscopy to analyze blood vessel patterns in the cleared tissues10.

<table>
	<tbody>
		<tr>
			<td>Sodium Dodecyl Sulfate (SDS)</td>
			<td>Affymetrix, Inc.</td>
			<td>75819</td>
			<td>Clearing solution</td>
		</tr>
		<tr>
			<td>Nycodenz</td>
			<td>Axia-Shield</td>
			<td>1002424</td>
			<td>nRIMS solution</td>
		</tr>
		<tr>
			<td>40% Acrylamide Solution</td>
			<td>Bio Rad Laboratories, Inc.</td>
			<td>161-0140</td>
			<td>Polymerization (A4P0)</td>
		</tr>
		<tr>
			<td>2,2&#180;-Azobis[2-(2-imidazolin-2-yl)propane] Dihydrochloride</td>
			<td>Wako Pure Chemical Industries, Ltd.</td>
			<td>017-19362</td>
			<td>Polymerization (VA-044)</td>
		</tr>
		<tr>
			<td>1-Thioglycerol</td>
			<td>Sigma-Aldrich</td>
			<td>M1753-100ML</td>
			<td>Clearing solution (mPACT)</td>
		</tr>
		<tr>
			<td>Tween-20</td>
			<td>Georgiachem</td>
			<td>9005-64-5</td>
			<td>nRIMS solution</td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Sigma-Aldrich</td>
			<td>T8787-50ML</td>
			<td>Immuno Staining</td>
		</tr>
		<tr>
			<td>Bovine serum albumin (BSA)</td>
			<td>Bovogen</td>
			<td>BSA100</td>
			<td>Immuno Staining</td>
		</tr>
		<tr>
			<td>Heparin</td>
			<td>Merck Millipore</td>
			<td>375095</td>
			<td>Perfusion (PBS)</td>
		</tr>
		<tr>
			<td>Sodium azide</td>
			<td>Sigma-Aldrich</td>
			<td>S2002-25G</td>
			<td>nRIMS solution</td>
		</tr>
		<tr>
			<td>PECAM-CD31 antibody</td>
			<td>Santa Cruz Biotechnology Inc.</td>
			<td>sc-28188</td>
			<td>Immuno Staining</td>
		</tr>
		<tr>
			<td>Goat anti-rabbit-IgG Cy3 fluorescent conjugate</td>
			<td>Jackson ImmunoResearch Inc.</td>
			<td>111-165-003</td>
			<td>Immuno Staining</td>
		</tr>
		<tr>
			<td>4% Paraformaldehyde</td>
			<td>Tech &#38; Innovation</td>
			<td>BPP-9004</td>
			<td>Perfusion, Polymerization</td>
		</tr>
		<tr>
			<td>20X Phosphate Buffered Saline (pH 7.4)</td>
			<td>Tech &#38; Innovation</td>
			<td>BPB-9121</td>
			<td>Perfusion, Buffer</td>
		</tr>
		<tr>
			<td>10 mL stripette</td>
			<td>Coatar</td>
			<td>4488</td>
			<td>Solution transfer</td>
		</tr>
		<tr>
			<td>50 mL tube</td>
			<td>Falcon</td>
			<td>352070</td>
			<td>Clearing tube</td>
		</tr>
		<tr>
			<td>35 mm Cell culture dish</td>
			<td>SPL</td>
			<td>20035</td>
			<td>Imaging</td>
		</tr>
		<tr>
			<td>Confocal dish</td>
			<td>SPL</td>
			<td>211350</td>
			<td>Imaging</td>
		</tr>
		<tr>
			<td>1 mL syringe</td>
			<td>Korea vaccine Co., Ltd</td>
			<td>26G 1/2</td>
			<td>Anesthetize&#160;</td>
		</tr>
		<tr>
			<td>50 mL syringe</td>
			<td>Korea vaccine Co., Ltd</td>
			<td>21G1 1/4</td>
			<td>Perfusion</td>
		</tr>
		<tr>
			<td>Acrylamide</td>
			<td>Sigma-Aldrich</td>
			<td>A3553</td>
			<td>Polymerization (A4P0)</td>
		</tr>
		<tr>
			<td>Whatman 3MM paper</td>
			<td>Sigma-Aldrich</td>
			<td>Z270849</td>
			<td>Blotting paper for gel removal</td>
		</tr>
		<tr>
			<td>Confocal microscope</td>
			<td>Zeiss</td>
			<td>LSM780</td>
			<td>Imaging</td>
		</tr>
		<tr>
			<td>ZEN lite Software</td>
			<td>Zeiss</td>
			<td>ZEN 2012</td>
			<td>Imaging</td>
		</tr>
		<tr>
			<td>Peristaltic pump</td>
			<td>Longerpump</td>
			<td>BT100-1F</td>
			<td>Perfusion</td>
		</tr>
		<tr>
			<td>EasyGel</td>
			<td>Lifecanvas Technologies</td>
			<td>EasyGel</td>
			<td>Tissue gel hybridization system</td>
		</tr>
	</tbody>
</table>

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.

Generation of a transparent model of the whole CNS using optimized passive clearing techniques
Optical clearance of mouse and rat whole CNS tissues was rapidly achieved using various passive clearing techniques (Figure 1). A schematic of tissue clearing over time is shown in Figure 2A. Unlike the original PACT method, psPACT (process-separate PACT) invo...

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

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, ...

novel-passive-clearing-methods-for-rapid-production-optical

Research

JoVE Journal

Neuroscience

8.7K Views.  Yonsei University College of Medicine. 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.

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

Methods for Experimental Manipulations after Optic Nerve Transection in the Mammalian CNS

Retinal ganglion cells (RGCs) are CNS neurons that output visual information from the retina to the brain, via the optic nerve. The optic nerve can be accessed within the orbit of the eye and completely transected (axotomized), cutting the axons of the entire RGC population. Optic nerve transection is a reproducible model of apoptotic neuronal cell death in the adult CNS 1-4. This model is particularly attractive because the vitreous chamber of the eye acts as a capsule for drug delivery to the retina, permitting experimental manipulations via intraocular injections. The diffusion of chemicals through the vitreous fluid ensures that they act upon the entire RGC population. Viral vectors, plasmids or short interfering RNAs (siRNAs) can also be delivered to the vitreous chamber in order to infect or transfect retinal cells 5-12. The high tropism of Adeno-Associated Virus (AAV) vectors is beneficial to target RGCs, with an infection rate approaching 90% of cells near the injection site 6, 7, 13-15. Moreover, RGCs can be selectively transfected by applying siRNAs, plasmids, or viral vectors to the cut end of the optic nerve 16-19 or injecting vectors into their target the superior colliculus 10. This allows researchers to study apoptotic mechanisms in the injured neuronal population without confounding effects on other bystander neurons or surrounding glia. RGC apoptosis has a characteristic time-course whereby cell death is delayed 3-4 days postaxotomy, after which the cells rapidly degenerate. This provides a window for experimental manipulations directed against pathways involved in apoptosis. Manipulations that directly target RGCs from the transected optic nerve stump are performed at the time of axotomy, immediately after cutting the nerve. In contrast, when substances are delivered via an intraocular route, they can be injected prior to surgery or within the first 3 days after surgery, preceding the initiation of apoptosis in axotomized RGCs. In the present article, we demonstrate several methods for experimental manipulations after optic nerve transection.

Optic Nerve transection is a widely used model of adult CNS injury. This model is ideal for performing a number of experimental manipulations that target the retina globally or directly target the injured neuronal population of retinal ganglion cells.

Retinal ganglion cells (RGCs) are CNS neurons that output visual information from the retina to the brain, via the optic nerve. The optic nerve can be ...

Fast Micro-iontophoresis of Glutamate and GABA: A Useful Tool to Investigate Synaptic Integration

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of the dendritic tree, which eventually leads to neuronal output of action potentials at the axon. The influence of diverse spatial and temporal parameters of specific synaptic input on neuronal output is currently under investigation, e.g. the distance-dependent attenuation of dendritic inputs, the location-dependent interaction of spatially segregated inputs, the influence of GABAergig inhibition on excitatory integration, linear and non-linear integration modes, and many more.
With fast micro-iontophoresis of glutamate and GABA it is possible to precisely investigate the spatial and temporal integration of glutamatergic excitation and GABAergic inhibition. Critical technical requirements are either a triggered fluorescent lamp, light-emitting diode (LED), or a two-photon scanning microscope to visualize dendritic branches without introducing significant photo-damage of the tissue. Furthermore, it is very important to have a micro-iontophoresis amplifier that allows for fast capacitance compensation of high resistance pipettes. Another crucial point is that no transmitter is involuntarily released by the pipette during the experiment.
Once established, this technique will give reliable and reproducible signals with a high neurotransmitter and location specificity. Compared to glutamate and GABA uncaging, fast iontophoresis allows using both transmitters at the same time but at very distant locations without limitation to the field of view. There are also advantages compared to focal electrical stimulation of axons: with micro-iontophoresis the location of the input site is definitely known and it is sure that only the neurotransmitter of interest is released. However it has to be considered that with micro-iontophoresis only the postsynapse is activated and presynaptic aspects of neurotransmitter release are not resolved. In this article we demonstrate how to set up micro-iontophoresis in brain slice experiments.

In this article we introduce fast micro-iontophoresis of neurotransmitters as a technique to investigate integration of postsynaptic signals with high spatial and temporal precision.

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of ...

Whole Mount Immunolabeling of Olfactory Receptor Neurons in the Drosophila Antenna

Odorant molecules bind to their target receptors in a precise and coordinated manner. Each receptor recognizes a specific signal and relays this information to the brain. As such, determining how olfactory information is transferred to the brain, modifying both perception and behavior, merits investigation. Interestingly, there is emerging evidence that cellular transduction and transcriptional factors are involved in the diversification of olfactory receptor neuron. Here we provide a robust whole mount immunological labeling method to assay in vivo olfactory receptor neuron organization. Using this method, we identified all olfactory receptor neurons with anti-ELAV antibody, a known pan-neural marker and Or49a-mCD8::GFP, an olfactory receptor neuron specifically expressed in Nba neuron using anti-GFP antibody.

Whole Mount Immunolabeling of Olfactory Receptor Neurons in the Drosophila Antenna

Herein we describe the process of whole mount immunostaining of Drosophila antennae, which enables us to better understand the molecular mechanisms involved in the diversification of olfactory receptor neurons (ORN)s.

Odorant molecules bind to their target receptors in a precise and coordinated manner. Each receptor recognizes a specific signal and relays this ...

Assessment of Vascular Regeneration in the CNS Using the Mouse Retina

The rodent retina is perhaps the most accessible mammalian system in which to investigate neurovascular interplay within the central nervous system (CNS). It is increasingly being recognized that several neurodegenerative diseases such as Alzheimer&#8217;s, multiple sclerosis, and amyotrophic lateral sclerosis present elements of vascular compromise. In addition, the most prominent causes of blindness in pediatric and working age populations (retinopathy of prematurity and diabetic retinopathy, respectively) are characterized by vascular degeneration and failure of physiological vascular regrowth. The aim of this technical paper is to provide a detailed protocol to study CNS vascular regeneration in the retina. The method can be employed to elucidate molecular mechanisms that lead to failure of vascular growth after ischemic injury. In addition, potential therapeutic modalities to accelerate and restore healthy vascular plexuses can be explored. Findings obtained using the described approach may provide therapeutic avenues for ischemic retinopathies such as that of diabetes or prematurity and possibly benefit other vascular disorders of the CNS.

The rodent retina has long been recognized as an accessible window to the brain. In this technical paper we provide a protocol that employs the mouse model of oxygen-induced retinopathy to study the mechanisms that lead to failure of vascular regeneration within the central nervous system after ischemic injury. The described system can also be harnessed to explore strategies to promote regrowth of functional blood vessels within the retina and CNS.

The rodent retina is perhaps the most accessible mammalian system in which to investigate neurovascular interplay within the central nervous system (CNS). ...

Optical Clearing of the Mouse Central Nervous System Using Passive CLARITY

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

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

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

A Simple One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the molecular and cellular basis of higher brain functions. A whole-mount preparation of adult brains with well-preserved morphology is critical for such whole brain-based studies, but can be technically challenging and time-consuming. This protocol describes an easy-to-learn, one-step dissection approach of an adult fly head in less than 10 s, while keeping the intact brain attached to the rest of the body to facilitate subsequent processing steps. The procedure helps remove most of the eye and tracheal tissues normally associated with the brain that can interfere with the later imaging step, and also places less demand on the quality of the dissecting forceps. Additionally, we describe a simple method that allows convenient flipping of the mounted brain samples on a coverslip, which is important for imaging both sides of the brains with similar signal intensity and quality. As an example of the protocol, we present an analysis of dopaminergic (DA) neurons in adult brains of WT (w1118) flies. The high efficacy of the dissection method makes it particularly useful for large-scale adult brain-based studies in Drosophila.

A Simple One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

The adult Drosophila brain is a valuable system for studying neuronal circuitry, higher brain functions, and complex disorders. An efficient method to dissect whole brain tissue from the small fly head will facilitate brain-based studies. Here we describe a simple, one-step dissection protocol of adult brains with well-preserved morphology.

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the ...

Methods for the Discovery of Novel Compounds Modulating a Gamma-Aminobutyric Acid Receptor Type A Neurotransmission

This manuscript presents a step-by-step protocol for screening compounds at gamma-aminobutyric acid type A (GABAA) receptors and its use towards the identification of novel molecules active in preclinical assays from an in vitro recombinant receptor to their pharmacological effects at native receptors in rodent brain slices. For compounds binding at the benzodiazepine site of the receptor, the first step is to set up a primary screen that consists of developing radioligand binding assays on cell membranes expressing the major GABAA subtypes. Then, taking advantage of the heterologous expression of rodent and human GABAA receptors in Xenopus oocytes or HEK 293 cells, it is possible to explore, in electrophysiological assays, the physiological properties of the different receptor subtypes and the pharmacological properties of the identified compounds. The Xenopus oocyte system will be presented here, starting with the isolation of the oocytes and their microinjection with different mRNAs, up to the pharmacological characterization using two-electrode voltage clamps. Finally, recordings conducted in rodent brain slices will be described that are used as a secondary physiological test to assess the activity of molecules at their native receptors in a well-defined neuronal circuit. Extracellular recordings using population responses of multiple neurons are demonstrated together with the drug application.

Here, we present protocols to discover compounds active at GABAA receptors, from the binding to the physiology and pharmacology.

This manuscript presents a step-by-step protocol for screening compounds at gamma-aminobutyric acid type A (GABAA) receptors and its use towards the ...

Chronic Implantation of Whole-cortical Electrocorticographic Array in the Common Marmoset

Electrocorticography (ECoG) allows the monitoring of electrical field potentials from the cerebral cortex with high spatiotemporal resolution. Recent development of thin, flexible ECoG electrodes has enabled conduction of stable recordings of large-scale cortical activity. We have developed a whole-cortical ECoG array for the common marmoset. The array continuously covers almost the entire lateral surface of cortical hemisphere, from the occipital pole to the temporal and frontal poles, and it captures whole-cortical neural activity in one shot. This protocol describes a chronic implantation procedure of the array in the epidural space of the marmoset brain. Marmosets have two advantages regarding ECoG recordings, one being the homologous organization of anatomical structures in humans and macaques, including frontal, parietal, and temporal complexes. The other advantage is that the marmoset brain is lissencephalic and contains a large number of complexes, which are more difficult to access in macaques with ECoG, that are exposed to the brain surface.These features allow direct access to most cortical areas beneath the surface of the brain. This system provides an opportunity to investigate global cortical information processing with high resolutions at a sub-millisecond order in time and millimeter order in space.

We have developed a whole-cortical electrocorticographic array for the common marmoset that continuously covers almost the entire lateral surface of cortex, from the occipital pole to the temporal and frontal poles. This protocol describes a chronic implantation procedure of the array in the epidural space of the marmoset brain.

Electrocorticography (ECoG) allows the monitoring of electrical field potentials from the cerebral cortex with high spatiotemporal resolution. Recent ...

A Hydrophobic Tissue Clearing Method for Rat Brain Tissue

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

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

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

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

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

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

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