This work was supported by the Belgian Foundation of Alzheimer&#39;s Research (SAO; project number: 20200032), the Research Foundation Flanders (FWO Vlaanderen; project numbers: 1268823N, 11D0520N, 1195021N) and the Baillet Latour Fund. We thank the VIB BioImaging Core for training, support, and access to the instrument park.

All animal experiments described in this study were conducted according to the national (Belgian Law 14/08/1986 and 22/12/2003, Belgian Royal Decree 06/04/2010) and European legislation (EU Directives 2010/63/EU, 86/609/EEC). All experiments on mice and animal protocols were approved by the ethics committee of Ghent University (permit numbers LA1400091 and EC 2017-026).
1. Preparation
<ol>
	<li>Anesthetics: Prepare a terminal anesthetic. For instance, a sodium pentobarbital (&#8805;100 mg/kg) solution in phosphate-buffered saline (PBS) can be prepared.</li>
	<li>Transcardial perfusion solution: Prepare 10 mL (per mouse) of PBS/heparin (containing 0.2% heparin) solution supplemented with 0.5% bromophenol blue. 
	&#8203;NOTE: If the downstream processing steps allow (e.g., in the case of RNA and protein analysis), the choroid plexus (CP) can be visualized via transcardial perfusion with bromophenol blue. If bromophenol blue is not compatible with the further processing steps (e.g., for imaging), use PBS/heparin to perfuse.</li>
	<li>Prepare the solutions required for SEM analysis.
	<ol>
		<li>Prepare 0.1 M Na-cacodylate buffer (pH 7.4). Prepare a solution of 2% paraformaldehyde and 2.5% glutaraldehyde in 0.1 M Na-cacodylate buffer. Make 20 mL of this solution per sample.</li>
		<li>Prepare 2% osmium tetroxide in 0.1 M Na-cacodylate buffer (5 mL per sample).</li>
		<li>Prepare 50%, 70%, 85%, and 100% EtOH solutions. Prepare 5 mL of each EtOH solution per sample.</li>
	</ol>
	</li>
</ol>
2. Microdissection of the choroid plexus out of the lateral and fourth ventricle
NOTE: Female, 9-week-old C57BL/6 mice were used in this study. However, the described isolation technique is independent of the strain, sex, and age of the adult mouse.
<ol>
	<li>Isolate the mouse brain.
	<ol>
		<li>Intraperitoneally inject a lethal dose of a short-acting barbiturate (&#62;100 mg/kg, prepared in step 1.1) using a 26G needle to terminally anesthetize the mouse. Check the foot reflex of the mouse by pinching its rear paw with forceps.</li>
		<li>When there is no foot reflex, place the terminally anesthetized animal in the dorsal decubitus position and fix the mouse by pinning the limbs on a plate. 
		NOTE: If transcardial perfusion of the mice is not needed, depending on the downstream analysis method (e.g., for SEM imaging), proceed to step 2.1.7. However, if perfusion is needed to remove blood cells or other components in the blood, the CP can be visualized as a blue structure floating in the brain ventricles via perfusion with bromophenol blue.</li>
		<li>Disinfect the chest by spraying 70% ethanol. Place a sterile drape around the surgical area. Make an incision of ~4 cm just under the diaphragm using a carbon steel surgical blade (see Table of Materials).</li>
		<li>Open the skin and expose the chest using surgical scissors. Cut the diaphragm completely open.</li>
		<li>Separate the thorax to expose the lungs and the beating heart.</li>
		<li>Transcardially perfuse the mice with 10 mL of the perfusion solution at a rate of 4.5 mL/min, using a perfusion pump (see Table of Materials). The perfusion will take ~2 min. Insert a 26G needle in the left ventricle to pump the solution into the systemic circuit. In addition, make a cut with surgical scissors in the right atrium so that blood can go out of the circulation. 
		NOTE: A perfusion pump is preferred over manual administration of the fluid, as it will deliver the fluids at a precisely programmed rate and ensure that the shearing forces caused by the perfusion are not too strong. Excessive shearing forces will compromise the viability and structure of cells within the CP. Additionally, the perfusion flow rate recommended here is optimal to perfuse adult mice from 7 weeks onward. If younger mice are used, a lower flow rate should be used. It is also important to dab away the blood that comes out of the right atrium in order to preserve a clean surgical site.</li>
		<li>Decapitate the mouse. 
		NOTE: Pay caution to cut off the head as low as possible to the shoulder to preserve the brain structure. If the cut is too high, the cerebellum can be damaged.</li>
		<li>After decapitation, cut open the scalp by a making an incision from between the ears to superior to the eyes. Pull the skin laterally to expose the skull. Then, cut open the skull by following the squamosal sutures, toward the nasal part. 
		NOTE: It is important to gently remove the skull to maintain the brain integrity.</li>
		<li>Place the brain in an ice-cold Petri dish and add 1 mL of ice-cold 1x PBS on the brain tissue. 
		NOTE:Coating of the dish is not needed.</li>
	</ol>
	</li>
	<li>Isolate the CP floating in the fourth ventricle.
	<ol>
		<li>Gently cut off the cerebellum from the cerebrum with a scalpel. The cerebrum is the largest part of the brain, while the cerebellum is a much smaller part at the back of the brain. If necessary, remove remaining brain stem tissue parts from the cerebellum.</li>
		<li>Rotate the brain so that the cutting line is facing upward. Ensure that the fourth ventricle cavity is now visible in the middle of the section, with the CP floating at the dorsal site. If necessary, open up the ventricle a bit by pulling open the connective tissue with sharp forceps. In this way, the CP will be more visible and easier to reach.</li>
		<li>Gently tear the CP out of the ventricle wall using tiny sharp forceps. 
		NOTE It is important to only touch and take out the CP in this step in order to not contaminate the sample with the surrounding tissue. Opening up the ventricles as described in step 2.2.2 will facilitate step 2.2.3.</li>
	</ol>
	</li>
	<li>Isolate the CP protruding out of the lateral ventricle.
	<ol>
		<li>Use tiny sharp forceps to sagittally cut the brain into two hemispheres.</li>
		<li>Rotate the brain so that the cutting line is facing upward. To reveal the lateral ventricle, gently pull away the cortex from the thalamus.</li>
		<li>Retract the hippocampus to the midsagittal cutting line. The lateral ventricle is now visible with the CP lying at the bottom of the ventricle. If necessary, open up the ventricle a bit by pulling open the connective tissue with sharp forceps. In this way, the CP will be more visible and easier to reach.</li>
		<li>Use tiny sharp forceps to gently tear the CP out of the ventricle wall. 
		&#8203;NOTE: It is important to only touch and take out the CP in this step in order to not contaminate the sample with the surrounding tissue. Opening up the ventricles as described in step 2.3.4 will facilitate step 2.3.5. Use necessary caution in this step to preserve the structure of the CP as much as possible. It is easiest to start pulling the CP in the rostral to caudal direction.</li>
	</ol>
	</li>
</ol>
3. Morphological analysis of CP tissue using scanning electron microscopy (SEM)
CAUTION: Toxic solutions are used in the following processing steps. It is recommended to perform the sample preparation in a fume hood.
<ol>
	<li>Perform sample preparation for SEM as described below.
	<ol>
		<li>Transfer the fresh isolated CP into freshly made fixation solution containing 2% paraformaldehyde and 2.5% glutaraldehyde in 0.1 M Na-cacodylate buffer (pH 7.4). Incubate overnight at 4 &#176;C. 
		NOTE: As the CP tissue is very fragile and thin, the tissue should be put in small specimen baskets (see Table of Materials) to transfer it between buffers. In this way, the structure of the tissue will be better preserved. A cacodylate-based fixative is used over a PBS-based one, as the strong cacodylate buffer can counter the low pH of glutaraldehyde in the EM fixative.</li>
		<li>Wash the sample 3x for 5 min each with 3-5 mL of 0.1 M Na-cacodylate buffer (pH 7.4).</li>
		<li>Post-fix the samples in 3-5 mL of 2% osmium tetroxide in 0.1 M Na-cacodylate buffer for 30 min. Wash the samples 3x for 5 min each with 3-5 mL of ultrapure water.</li>
		<li>Dehydrate the samples in a series of ice-cold solutions of increasing EtOH concentrations (50%, 70%, 85%, 100%), for 15 min per EtOH solution. Use a critical point dryer to properly point dry the sample. 
		NOTE Changing from liquid to gas phase affects the surface tension, causing damage to the surface structure. The critical point drying step allows for the preservation of the surface structure.</li>
		<li>Position the sample carefully on a specimen mount provided with a carbon sticker (see Table of Materials).</li>
		<li>Coat the samples with a thin layer (2-5 nm) of platinum. To do so, mount a platinum source in a vacuum system between two high-current electrical terminals and heat the platinum to its evaporation temperature. A fine stream of platinum is deposited on the sample. 
		NOTE: A more detailed protocol for this step is provided in reference30. Besides platinum, gold or gold/palladium can also be used.</li>
	</ol>
	</li>
	<li>Visualize the CP samples with SEM (see Table of Materials). The visualization procedure of CP tissue via SEM is similar to that of other types of tissues and depends on the used software and instruments. A step by step SEM protocol with an accompanying video is available in reference31 or reference30.</li>
</ol>

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The authors have nothing to disclose.

Here, a method to isolate the choroid plexus (CP) out of the lateral ventricle and the fourth ventricle of a mouse brain is described. This whole mounting method of the CP facilitates further analysis using a repertoire of techniques to get a complete view of the CP morphology, cellular composition, transcriptome, proteome, and secretome. Such analyses are crucial to gain a better understanding of this remarkable structure protruding from the ventricles of the brain. This knowledge is of immense research interest, as it ...

Tight barriers separate the central nervous system (CNS) from the periphery, including the blood-brain barrier (BBB) and the blood-cerebrospinal fluid (CSF) barrier. These barriers protect the CNS against external insults and ensure a balanced and controlled microenvironment1,2,3. While the BBB has been extensively studied over time, the blood-CSF barrier located at the choroid plexus (CP) has only gained increasing research interest during the last decade. This latter barrier can be found in the four ventricles of the brain (Figure 1A, B)&#160;and is characterized by a single layer of choroid plexus epithelial (CPE) cells surrounding a central stroma, leaky capillaries, fibroblasts, and a lymphoid and myeloid cell population (Figure 1C)4,5,6. The CPE cells are firmly interconnected by tight junctions, thus preventing leakage from the underlying fenestrated blood capillaries into the CSF and brain. Additionally, transport across the CPE cells is regulated by a number of inward and outward transport systems that manage the influx of beneficial compounds (e.g., nutrients and hormones) from the blood to the CSF and the efflux of harmful molecules (e.g., metabolic waste, excess neurotransmitters) in the other direction1,6. To be able to exert their active transport function, the CPE cells contain numerous mitochondria in their cytoplasm7. Moreover, the CP is the main source of CSF and acts as the gatekeeper of the brain by the presence of resident inflammatory cells1. Due to its unique location between the blood and the brain, the CP is also perfectly positioned to carry out immune surveillance8.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64733/64733fig01.jpg" /> 
Figure 1: Schematic overview of the location and composition of the choroid plexus (CP). (A,B)&#160;CP tissue is found within the two lateral, the third, and the fourth ventricles of (A) human and (B) mouse brains. (C) The CP tissue consists of a single layer of tightly connected cuboidal CP epithelium (CPE) cells surrounding fenestrated capillaries, loose connective tissue, and lymphoid and myeloid cells, and forms the blood-cerebrospinal fluid barrier (adapted and modified from reference23). Figure created with Biorender.com. <a href="https://www.jove.com/files/ftp_upload/64733/64733fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Over the past decade, increasing evidence, including several reports from our research group, have revealed that the CP plays a central role in health and disease9,10,11,12,13,14,15,16,17,18. For example, it is known that the aging blood-CSF barrier displays morphological alterations in, among others, the nuclei, microvilli, and the basement membrane1,19. Additionally, in the context of Alzheimer&#39;s disease, the overall barrier integrity is compromised and all of these age-related changes appear to be even more pronounced1,8,20. In addition to morphological changes, the transcriptome, proteome, and secretome of the CP are altered during disease12,21,22,23. Thus, advanced knowledge of the CP is essential to better understand its role in neurological diseases and potentially develop new therapeutic strategies.
An efficient method for accurate microdissection of the CP out of the brain ventricles is the first invaluable step to allow proper investigation of this tiny brain structure. On account of its highly vascularized nature (Figure 2B), the CP floating inside the ventricular cavities of the brain can be identified using a binocular microscope. However, transcardial perfusion is often required for downstream analysis, complicating the proper identification and isolation of the CP tissue (Figure 2C). If the further processing steps allow (e.g., in the case of RNA and protein analysis), the CP can be visualized via transcardial perfusion with bromophenol blue (Figure 2A). Several publications already describe the isolation of the CP from rat24 and mouse pup brains25. Here, a microdissection isolation technique is described to isolate the CP from adult mice. Importantly, this isolation technique preserves the viability, function, and structure of the cells within the CP. The isolation of the CP floating in the fourth and lateral ventricles is described here. In short, the mice are terminally anesthetized&#160;and, if necessary, transcardially perfused. However, it should be noted that perfusion can damage the structure of the cells within the CP. Consequently, if the sample is to be analyzed using transmission electron microscopy (TEM), serial block face scanning electron microscopy (SBF-SEM), or focused ion beam SEM (FIB-SEM), perfusion should not be performed. Next, the whole brain is isolated, and forceps are used to sagittally hemisect the brain. From here, the CPs floating in the lateral ventricles can be identified and dissected, while the CP from the fourth ventricle can be isolated from the cerebellar side of the brain.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64733/64733fig02.jpg" /> 
Figure 2: Visualization of the (A-C) fourth and (D-F) lateral ventricle choroid plexus (CP) after (A,D) bromophenol blue perfusion, (B,E) no perfusion, and (C,F) perfusion with PBS/heparin. The images are taken with a stereo microscope (8x-32x magnification). <a href="https://www.jove.com/files/ftp_upload/64733/64733fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Once the CP is properly dissected out of the brain ventricles, a whole repertoire of techniques can be applied to gain further understanding on the function of this structure. For example, flow cytometry or single cell RNA sequencing can be performed to quantify and phenotypically analyze the infiltrating inflammatory cells under certain disease conditions26,27. In addition to the cellular composition, the molecular composition of the CP can be analyzed to assess the presence of cytokines and chemokines via enzyme-linked immunosorbent assay (ELISA), immunoblot, or through simultaneous analysis of multiple cytokines using the cytokine bead array28. Moreover, transcriptome, vascular, immune cell histology, and secretome analyses can be performed on the microdissected CP explants29. Here, scanning electron microscopy (SEM) on whole mount CP is used to obtain an overall view of the CP structure. SEM uses a beam of focused electrons to scan over the surface and create an image of the surface&#39;s topography and composition. Since the wavelength of electrons is much smaller than that of light, the resolution of SEM is in the nanometer range and superior to that of a light microscope. Consequently, morphological studies on the subcellular level can be performed via SEM. Briefly, the dissected CP is immediately transferred into a glutaraldehyde-containing fixative for an overnight fixation, followed by osmication and uranyl acetate staining. The samples are then treated with lead aspartate stain, dehydrated, and ultimately embedded for imaging.
Thus, this protocol facilitates the efficient isolation of the CP from the mouse brain ventricles, which can be further analyzed using a variety of downstream techniques to investigate its structure and function.

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microdissection and whole mount scanning electron microscopy visualization of mouse choroid plexus

The choroid plexus (CP), a highly vascularized structure protruding into the ventricles of the brain, is one of the most understudied tissues in neuroscience. As it is becoming increasingly clear that this tiny structure plays a crucial role in health and disease of the central nervous system (CNS), it is of utmost importance to properly dissect the CP out of the brain ventricles in a way that allows downstream processing, ranging from functional to structural analysis. Here, isolation of the lateral and fourth brain ventricle mouse CP without the need for specialized tools or equipment is described. This isolation technique preserves the viability, function, and structure of cells within the CP. On account of its high vascularization, the CP can be visualized floating inside the ventricular cavities of the brain using a binocular microscope. However, transcardial perfusion required for downstream analysis can complicate the identification of the CP tissue. Depending on the further processing steps (e.g., RNA and protein analysis), this can be solved by visualizing the CP via transcardial perfusion with bromophenol blue. After isolation, the CP can be processed using several techniques, including RNA, protein, or single cell analysis, to gain further understanding on the function of this special brain structure. Here, scanning electron microscopy (SEM) on whole mount CP is used to get an overall view of the structure.

The described protocol facilitates the efficient isolation of the CP from the mouse brain lateral (Figure 2A-C) and fourth (Figure 2D-F)&#160;ventricles. After isolating the whole brain, forceps are used to sagittally hemisect the brain and identify the CPs floating in the lateral ventricles. The CP from the fourth ventricle can be isolated from the cerebellar side of the brain. Perfusion with b...

Watch this Scientific Journal Video about Microdissection and Whole Mount Scanning Electron Microscopy Visualization of Mouse Choroid Plexus at JoVE.com

Microdissection and Whole Mount Scanning Electron Microscopy Visualization of Mouse Choroid Plexus

The choroid plexus (CP), an understudied tissue in neuroscience, plays a key role in health and disease of the central nervous system. This protocol describes a microdissection technique for isolating the CP and the use of scanning electron microscopy to obtain an overall view of its cellular structure.

The choroid plexus (CP), a highly vascularized structure protruding into the ventricles of the brain, is one of the most understudied tissues in ...

microdissection-whole-mount-scanning-electron-microscopy

Research

JoVE Journal

Neuroscience

3.8K Views.  VIB. The choroid plexus (CP), an understudied tissue in neuroscience, plays a key role in health and disease of the central nervous system. This protocol describes a microdissection technique for isolating the CP and the use of scanning electron microscopy to obtain an overall view of its cellular structure.

Video: Microdissection and Whole Mount Scanning Electron Microscopy Visualization of Mouse Choroid Plexus

Visualization of the Embryonic Nervous System in Whole-mount Drosophila Embryos

The Drosophila embryo is an attractive model system for investigating the cellular and molecular basis of neuronal development. Here we describe the procedure for the visualization of Drosophila embryonic nervous system using antibodies to neuronal proteins. Since the entire embryonic peripheral nervous and central nervous systems are well characterized at the level of individual cells (Dambly-Chaudi&egrave;re et al., 1986; Bodmer et al., 1987; Bodmer et al., 1989), any aberrations to these systems can be easily identified using antibodies to different neuronal proteins. The developing embryos are collected at certain times to ensure that the embryos are in the proper developmental stages for visualization. After collection, the outer layers of the embryo, the chorion membrane and the vitelline envelope that surrounds the embryo, are removed before fixation. Embryos are then incubated with neuronal antibodies and visualized using fluorescently labeled secondary antibodies. Embryos at stages 12-17 are visualized to access the embryonic nervous system. At stage 12 the CNS germ band starts shortening and by stage 15 the definitive pattern of the commissure has been achieved. By stage 17 the CNS contracts and the PNS is fully developed (Campos-Ortega et al. 1985). Thus changes in the pattern of the PNS and CNS can be easily observed during these developmental stages.

Visualization of the Embryonic Nervous System in Whole-mount Drosophila Embryos

We describe the procedure to prepare staged Drosophila embryos for the visualization of the embryonic nervous system during embryogenesis.

The Drosophila embryo is an attractive model system for investigating the cellular and molecular basis of neuronal development. Here we describe the ...

Dissection of a Mouse Eye for a Whole Mount of the Retinal Pigment Epithelium

The retinal pigment epithelium (RPE) lies at the back of the mammalian eye, just under the neural retina, which contains the photoreceptors (rods and cones). The RPE is a monolayer of pigmented cuboidal cells and associates closely with the neural retina just above it. This association makes the RPE of great interest to researchers studying retinal diseases. The RPE is also the site of an in vivo assay of homology-directed DNA repair, the pun assay. The mouse eye is particularly difficult to dissect due to its small size (about 3.5mm in diameter) and its spherical shape. This article demonstrates in detail a procedure for dissection of the eye resulting in a whole mount of the RPE. In this procedure, we show how to work with, rather than against, the spherical structure of the eye. Briefly, the connective tissue, muscle, and optic nerve are removed from the back of the eye. Then, the cornea and lens are removed. Next, strategic cuts are made that result in significant flattening of the remaining tissue. Finally, the neural retina is gently lifted off, revealing an intact RPE, which is still attached to the underlying choroid and sclera. This whole mount can be used to perform the pun assay or for immunohistochemistry or immunofluorescent assessment of the RPE tissue.

A formal demonstration of the dissection of a mouse eye, resulting in a whole mount of the retinal pigment epithelium.

The retinal pigment epithelium (RPE) lies at the back of the mammalian eye, just under the neural retina, which contains the photoreceptors (rods and ...

Focussed Ion Beam Milling and Scanning Electron Microscopy of Brain Tissue

This protocol describes how biological samples, like brain tissue, can be imaged in three dimensions using the focussed ion beam/scanning electron microscope (FIB/SEM). The samples are fixed with aldehydes, heavy metal stained using osmium tetroxide and uranyl acetate. They are then dehydrated with alcohol and infiltrated with resin, which is then hardened. Using a light microscope and ultramicrotome with glass knives, a small block containing the region interest close to the surface is made. The block is then placed inside the FIB/SEM, and the ion beam used to roughly mill a vertical face along one side of the block, close to this region. Using backscattered electrons to image the underlying structures, a smaller face is then milled with a finer ion beam and the surface scrutinised more closely to determine the exact area of the face to be imaged and milled. The parameters of the microscope are then set so that the face is repeatedly milled and imaged so that serial images are collected through a volume of the block. The image stack will typically contain isotropic voxels with dimenions as small a 4 nm in each direction. This image quality in any imaging plane enables the user to analyse cell ultrastructure at any viewing angle within the image stack.

This protocol describes how resin embedded brain tissue can be prepared and imaged in the three dimensions in the focussed ion beam, scanning electron microscope.

This protocol describes how biological samples, like brain tissue, can be imaged in three dimensions using the focussed ion beam/scanning electron ...

Reproducible Mouse Sciatic Nerve Crush and Subsequent Assessment of Regeneration by Whole Mount Muscle Analysis

Regeneration in the peripheral nervous system (PNS) is widely studied both for its relevance to human disease and to understand the robust regenerative response mounted by PNS neurons thereby possibly illuminating the failures of CNS regeneration1. Sciatic nerve crush (axonotmesis) is one of the most common models of peripheral nerve injury in rodents2. Crushing interrupts all axons but Schwann cell basal laminae are preserved so that regeneration is optimal3,4. This allows the investigator to study precisely the ability of a growing axon to interact with both the Schwann cell and basal laminae4. Rats have generally been the preferred animal models for experimental nerve crush. They are widely available and their lesioned sciatic nerve provides a reasonable approximation of human nerve lesions5,4. Though smaller in size than rat nerve, the mouse nerve has many similar qualities. Most importantly though, mouse models are increasingly valuable because of the wide availability of transgenic lines now allows for a detailed dissection of the individual molecules critical for nerve regeneration6, 7. Prior investigators have used multiple methods to produce a nerve crush or injury including simple angled forceps, chilled forceps, hemostatic forceps, vascular clamps, and investigator-designed clamps8,9,10,11,12. Investigators have also used various methods of marking the injury site including suture, carbon particles and fluorescent beads13,14,1. We describe our method to obtain a reproducibly complete sciatic nerve crush with accurate and persistent marking of the crush-site using a fine hemostatic forceps and subsequent carbon crush-site marking. As part of our description of the sciatic nerve crush procedure we have also included a relatively simple method of muscle whole mount we use to subsequently quantify regeneration.

In this report we describe a method to crush mouse sciatic nerve. This method uses readily available hemostatic forceps and easily and reproducibly produces complete sciatic nerve crush. In addition, we describe a method to prepare muscle whole mounts suitable for analysis of nerve regeneration after sciatic nerve crush.

Regeneration in the peripheral nervous system (PNS) is widely studied both for its relevance to human disease and to understand the robust regenerative ...

A Visual Description of the Dissection of the Cerebral Surface Vasculature and Associated Meninges and the Choroid Plexus from Rat Brain

This video presentation was created to show a method of harvesting the two most important highly vascular structures, not residing within the brain proper, that support forebrain function. They are the cerebral surface (superficial) vasculature along with associated meninges (MAV) and the choroid plexus which are necessary for cerebral blood flow and cerebrospinal fluid (CSF) homeostasis. The tissue harvested is suitable for biochemical and physiological analysis, and the MAV has been shown to be sensitive to damage produced by amphetamine and hyperthermia 1,2. As well, the major and minor cerebral vasculatures harvested in MAV are of potentially high interest when investigating concussive types of head trauma. The MAV dissected in this presentation consists of the pial and some of the arachnoid membrane (less dura) of the meninges and the major and minor cerebral surface vasculature. The choroid plexus dissected is the structure that resides in the lateral ventricles as described by Oldfield and McKinley3,4,5,6. The methods used for harvesting these two tissues also facilitate the harvesting of regional cortical tissue devoid of meninges and larger cerebral surface vasculature, and is compatible with harvesting other brain tissues such as striatum, hypothalamus, hippocampus, etc. The dissection of the two tissues takes from 5 to 10 min total. The gene expression levels for the dissected MAV and choroid plexus, as shown and described in this presentation can be found at GSE23093 (MAV) and GSE29733 (choroid plexus) at the NCBI GEO repository. This data has been, and is being, used to help further understand the functioning of the MAV and choroid plexus and how neurotoxic events such as severe hyperthermia and AMPH adversely affect their function.

This video presentation shows a method of harvesting the two most important highly vascular structures that support forebrain function. They are the cerebral surface (superficial) vasculature along with associated meninges (MAV) and the choroid plexus which are necessary for cerebral blood flow and cerebrospinal fluid (CSF) homeostasis.

This video presentation was created to show a method of harvesting the two most important highly vascular structures, not residing within the brain ...

Whole-mount Imaging of Mouse Embryo Sensory Axon Projections

The visualization of full-length neuronal projections in embryos is essential to gain an understanding of how mammalian neuronal networks develop. Here we describe a method to label in situ a subset of dorsal root ganglion (DRG) axon projections to assess their phenotypic characteristics using several genetically manipulated mouse lines. The TrkA-positive neurons are nociceptor neurons, dedicated to the transmission of pain signals. We utilize a TrkAtaulacZ mouse line to label the trajectories of all TrkA-positive peripheral axons in the intact mouse embryo. We further breed the TrkAtaulacZ line onto a Bax null background, which essentially abolishes neuronal apoptosis, in order to assess growth-related questions independently of possible effects of genetic manipulations on neuronal survival. Subsequently, genetically modified mice of interest are bred with the TrkAtaulacZ/Bax null line and are then ready for study using the techniques described herein. This presentation includes detailed information on mouse breeding plans, genotyping at the time of dissection, tissue preparation, staining and clearing to allow for visualization of full-length axonal trajectories in whole-mount preparation.

We present here an optimized protocol to genotype, stain and prepare fetal mice for the imaging of peripheral nociceptor axon projections in the whole animal, as an effective method to assess sensory axon growth phenotypes in developing genetically engineered mice.

The visualization of full-length neuronal projections in embryos is essential to gain an understanding of how mammalian neuronal networks develop. Here we ...

Observation of the Ciliary Movement of Choroid Plexus Epithelial Cells Ex Vivo

The choroid plexus is located in the ventricular wall of the brain, the main function of which is believed to be production of cerebrospinal fluid. Choroid plexus epithelial cells (CPECs) covering the surface of choroid plexus tissue harbor multiple unique cilia, but most of the functions of these cilia remain to be investigated. To uncover the function of CPEC cilia with particular reference to their motility, an ex vivo observation system was developed to monitor ciliary motility during embryonic, perinatal and postnatal periods. The choroid plexus was dissected out of the brain ventricle and observed under a video-enhanced contrast microscope equipped with differential interference contrast optics. Under this condition, a simple and quantitative method was developed to analyze the motile profiles of CPEC cilia for several hours ex vivo. Next, the morphological changes of cilia during development were observed by scanning electron microscopy to elucidate the relationship between the morphological maturity of cilia and motility. Interestingly, this method could delineate changes in the number and length of cilia, which peaked at postnatal day (P) 2, while the beating frequency reached a maximum at P10, followed by abrupt cessation at P14. These techniques will enable elucidation of the functions of cilia in various tissues. While related techniques have been published in a previous report1, the current study focuses on detailed techniques to observe the motility and morphology of CPEC cilia ex vivo.

Observation of the Ciliary Movement of Choroid Plexus Epithelial Cells Ex Vivo

In this study, a detailed light microscopic technique was optimized for real-time observation and analysis of the motion of CPEC cilia ex vivo together with an electron microscopic method for ultrastructural analysis.

The choroid plexus is located in the ventricular wall of the brain, the main function of which is believed to be production of cerebrospinal fluid. ...

Whole Mount Dissection and Immunofluorescence of the Adult Mouse Cochlea

The organ of Corti, housed in the cochlea of the inner ear, contains mechanosensory hair cells and surrounding supporting cells which are organized in a spiral shape and have a tonotopic gradient for sound detection. The mouse cochlea is approximately 6 mm long and often divided into three turns (apex, middle, and base) for analysis. To investigate cell loss, cell division, or mosaic gene expression, the whole mount or surface preparation of the cochlea is useful. This dissection method allows visualization of all cells within the organ of Corti when combined with immunostaining and confocal microscopy to image cells at different planes in the z-axis. Multiple optical cross-sections can also be obtained from these z-stack images. In addition, the whole mount dissection method can be used for scanning electron microscopy, although a different fixation method is needed. Here, we present a method to isolate the organ of Corti as three intact cochlear turns (apex, middle, and base). This method can be used for mice ranging from one week of age through adulthood and differs from the technique used for neonatal samples where calcification of the cochlea is incomplete. A slightly modified version can be used for dissection of the rat cochlea. We also demonstrate a procedure for immunostaining with fluorescently tagged antibodies.

We present a method to isolate the adult organ of Corti as three intact cochlear turns (apex, middle, and base). We also demonstrate a procedure for immunostaining with fluorescently tagged antibodies. Together these techniques allow the study of hair cells, supporting cells, and other cell types found in the cochlea.

The organ of Corti, housed in the cochlea of the inner ear, contains mechanosensory hair cells and surrounding supporting cells which are organized in a ...

Analysis of Brain Mitochondria Using Serial Block-Face Scanning Electron Microscopy

Human brain is a high energy consuming organ that mainly relies on glucose as a fuel source. Glucose is catabolized by brain mitochondria via glycolysis, tri-carboxylic acid (TCA) cycle and oxidative phosphorylation (OXPHOS) pathways to produce cellular energy in the form of adenosine triphosphate (ATP). Impairment of mitochondrial ATP production causes mitochondrial disorders, which present clinically with prominent neurological and myopathic symptoms. Mitochondrial defects are also present in neurodevelopmental disorders (e.g. autism spectrum disorder) and neurodegenerative disorders (e.g. amyotrophic lateral sclerosis, Alzheimer&#39;s and Parkinson&#39;s diseases). Thus, there is an increased interest in the field for performing 3D analysis of mitochondrial morphology, structure and distribution under both healthy and disease states. The brain mitochondrial morphology is extremely diverse, with some mitochondria especially those in the synaptic region being in the range of &#60;200 nm diameter, which is below the resolution limit of traditional light microscopy. Expressing a mitochondrially-targeted green fluorescent protein (GFP) in the brain significantly enhances the organellar detection by confocal microscopy. However, it does not overcome the constraints on the sensitivity of detection of relatively small sized mitochondria without oversaturating the images of large sized mitochondria. While serial transmission electron microscopy has been successfully used to characterize mitochondria at the neuronal synapse, this technique is extremely time-consuming especially when comparing multiple samples. The serial block-face scanning electron microscopy (SBFSEM) technique involves an automated process of sectioning, imaging blocks of tissue and data acquisition. Here, we provide a protocol to perform SBFSEM of a defined region from rodent brain to rapidly reconstruct and visualize mitochondrial morphology. This technique could also be used to provide accurate information on mitochondrial number, volume, size and distribution in a defined brain region. Since the obtained image resolution is high (typically under 10 nm) any gross mitochondrial morphological defects may also be detected.

Mitochondrial visualization and analysis from mammalian brain tissue is a challenging task. Here, we describe how three dimensional (3D) reconstruction analysis from the serial block-face scanning electron microscopy (SBFSEM) can be used to gain insights on the morphological and volumetric analysis of this critical energy generating organelle.

Human brain is a high energy consuming organ that mainly relies on glucose as a fuel source. Glucose is catabolized by brain mitochondria via glycolysis, ...

Whole-Mount Staining, Visualization, and Analysis of Fungiform, Circumvallate, and Palate Taste Buds

Taste buds are collections of taste-transducing cells specialized to detect subsets of chemical stimuli in the oral cavity. These transducing cells communicate with nerve fibers that carry this information to the brain. Because taste-transducing cells continuously die and are replaced throughout adulthood, the taste-bud environment is both complex and dynamic, requiring detailed analyses of its cell types, their locations, and any physical relationships between them. Detailed analyses have been limited by tongue-tissue heterogeneity and density that have significantly reduced antibody permeability. These obstacles require sectioning protocols that result in splitting taste buds across sections so that measurements are only approximated, and cell relationships are lost. To overcome these challenges, the methods described herein involve collecting, imaging, and analyzing whole taste buds and individual terminal arbors from three taste regions: fungiform papillae, circumvallate papillae, and the palate. Collecting whole taste buds reduces bias and technical variability and can be used to report absolute numbers for features including taste-bud volume, total taste-bud innervation, transducing-cell counts, and the morphology of individual terminal arbors. To demonstrate the advantages of this method, this paper provides comparisons of taste bud and innervation volumes between fungiform and circumvallate taste buds using a general taste-bud marker and a label for all taste fibers. A workflow for the use of sparse-cell genetic labeling of taste neurons (with labeled subsets of taste-transducing cells) is also provided. This workflow analyzes the structures of individual taste-nerve arbors, cell type numbers, and the physical relationships between cells using image analysis software. Together, these workflows provide a novel approach for tissue preparation and analysis of both whole taste buds and the complete morphology of their innervating arbors.

This paper describes methods for tissue preparation, staining, and analysis of whole fungiform, circumvallate, and palate taste buds that consistently yield whole and intact taste buds (including the nerve fibers that innervate them) and maintain the relationships between structures within taste buds and the surrounding papilla.

Taste buds are collections of taste-transducing cells specialized to detect subsets of chemical stimuli in the oral cavity. These transducing cells ...