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.

<table><tbody><tr><td>26G x 1/2 needle</td><td>Henke Sass Wolf</td><td>4710004512</td><td></td></tr><tr><td>Aluminium specimen mounts</td><td>EM Sciences</td><td>75220</td><td></td></tr><tr><td>Cacodylate buffer</td><td>EM Sciences</td><td>11652</td><td></td></tr><tr><td>Carbon steel surgial blades</td><td>Swann-Morton</td><td>0210</td><td>size: 0.45 mm x 12 mm</td></tr><tr><td>Carbon adhesive tabs -12 mm</td><td>EM Sciences</td><td>77825-12</td><td></td></tr><tr><td>Critical point dryer&amp;nbsp;</td><td>Bal-Tec&amp;nbsp;</td><td>CPD030</td><td></td></tr><tr><td>Crossbeam 540</td><td>Zeiss</td><td></td><td>SEM system</td></tr><tr><td>Forceps</td><td>Fine Science Tools GmbH&amp;nbsp;</td><td>91197-00</td><td></td></tr><tr><td>Glutaraldehyde</td><td>EM Sciences</td><td>16220</td><td></td></tr><tr><td>Heparin</td><td>Sigma-Aldrich</td><td>H-3125</td><td></td></tr><tr><td>Ismatec Reglo ICC Digital Peristaltic pump 2-channel</td><td>Metrohm Belgium N.V</td><td>CPA-7800160</td><td></td></tr><tr><td>Osmium Tetroxide&amp;nbsp;</td><td>EM Sciences</td><td>19170</td><td></td></tr><tr><td>Paraformaldehyde</td><td>Sigma-Aldrich</td><td>P6148</td><td></td></tr><tr><td>Phosphate buffered saline (PBS)</td><td>Lonza</td><td>BE17-516F</td><td></td></tr><tr><td>Platinum&amp;nbsp;</td><td>Quorum&amp;nbsp;</td><td>Q150T ES</td><td>PBS without Ca++ Mg++ or phenol red; sterile filtered</td></tr><tr><td>Sodium pentobarbital</td><td>Kela NV</td><td>514</td><td></td></tr><tr><td>Specimen Basket Stainless Steel</td><td>EM Sciences</td><td>70190-01</td><td></td></tr><tr><td>Stemi DV4 Stereo microscope</td><td>Zeiss</td><td></td><td></td></tr><tr><td>Surgical scissors</td><td>Fine Science Tools GmbH&amp;nbsp;</td><td>91460-11</td><td></td></tr></tbody></table>

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...

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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

3.9K 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

Intravital Microscopy of the Mouse Brain Microcirculation using a Closed Cranial Window

This experimental model was designed to assess the mouse pial microcirculation during acute and chronic, physiological and pathophysiological hemodynamic, inflammatory and metabolic conditions, using in vivo fluorescence microscopy. A closed cranial window is placed over the left parieto-occipital cortex of the mice. Local microcirculation is recorded in real time through the window using epi and fluorescence illumination, and measurements of vessels diameters and red blood cell (RBC) velocities are performed. RBC velocity is measured using real-time cross-correlation and/or fluorescent-labeled erythrocytes. Leukocyte and platelet adherence to pial vessels and assessment of perfusion and vascular leakage are made with the help of fluorescence-labeled markers such as Albumin-FITC and anti-CD45-TxR antibodies. Microcirculation can be repeatedly video-recorded over several days. We used for the first time the close window brain intravital microscopy to study the pial microcirculation to follow dynamic changes during the course of Plasmodium berghei ANKA infection in mice and show that expression of CM is associated with microcirculatory dysfunctions characterized by vasoconstriction, profound decrease in blood flow and eventually vascular collapse.

Intravital microscopy to follow temporal and spatial hemodynamic and inflammatory events in the pial microcirculation.

This experimental model was designed to assess the mouse pial microcirculation during acute and chronic, physiological and pathophysiological hemodynamic, ...

Whole-mount Confocal Microscopy for Adult Ear Skin: A Model System to Study Neuro-vascular Branching Morphogenesis and Immune Cell Distribution

Here, we present a protocol of a whole-mount adult ear skin imaging technique to study comprehensive three-dimensional neuro-vascular branching morphogenesis and patterning, as well as immune cell distribution at a cellular level. The analysis of peripheral nerve and blood vessel anatomical structures in adult tissues provides some insights into the understanding of functional neuro-vascular wiring and neuro-vascular degeneration in pathological conditions such as wound healing. As a highly informative model system, we have focused our studies on adult ear skin, which is readily accessible for dissection. Our simple and reproducible protocol provides an accurate depiction of the cellular components in the entire skin, such as peripheral nerves (sensory axons, sympathetic axons, and Schwann cells), blood vessels (endothelial cells and vascular smooth muscle cells), and inflammatory cells. We believe this protocol will pave the way to investigate morphological abnormalities in peripheral nerves and blood vessels as well as the inflammation in the adult ear skin under different pathological conditions.

Here, we describe a high resolution whole-mount imaging method in the entire adult mouse ear skin, which enables us to visualize branching morphogenesis and patterning of peripheral nerves and blood vessels, as well as immune cell distribution.

Here, we present a protocol of a whole-mount adult ear skin imaging technique to study comprehensive three-dimensional neuro-vascular branching ...

Developmental Biology

Visualizing the Node and Notochordal Plate In Gastrulating Mouse Embryos Using Scanning Electron Microscopy and Whole Mount Immunofluorescence

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is the formation of the transient organizers, the node and notochordal plate, from cells that have passed through the primitive streak. The proper formation of these signaling centers is essential for the development of the body plan and techniques to visualize them are of high interest to mouse developmental biologists. The node and notochordal plate lie on the ventral surface of gastrulating mouse embryos around embryonic day (E) 7.5 of development. The node is a cup-shaped structure whose cells possess a single slender cilium each. The proper subcellular localization and rotation of the cilia in the node pit determines left-right asymmetry. The notochordal plate cells also possess single cilia albeit shorter than those of the node cells. The notochordal plate forms the notochord which acts as an important signaling organizer for somitogenesis and neural patterning. Because the cells of the node and notochordal plate are transiently present on the surface and possess cilia, they can be visualized using scanning electron microscopy (SEM). Among other techniques used to visualize these structures at the cellular level is whole mount immunofluorescence (WMIF) using the antibodies against the proteins that are highly expressed in the node and notochordal plate. In this report, we describe our optimized protocols to perform SEM and WMIF of the node and notochordal plate in developing mouse embryos to help in the assessment of tissue shape and cellular organization in wild-type and gastrulation mutant embryos.

The node and notochordal plate are transient signaling organizers in developing mouse embryos that can be visualized using several techniques. Here, we describe in detail how to perform two of the techniques to study their structure and morphogenesis: 1) scanning electron microscopy (SEM); and 2) whole mount immunofluorescence (WMIF).

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is ...

Micron-scale Resolution Optical Tomography of Entire Mouse Brains with Confocal Light Sheet Microscopy

Understanding the architecture of mammalian brain at single-cell resolution is one of the key issues of neuroscience. However, mapping neuronal soma and projections throughout the whole brain is still challenging for imaging and data management technologies. Indeed, macroscopic volumes need to be reconstructed with high resolution and contrast in a reasonable time, producing datasets in the TeraByte range. We recently demonstrated an optical method (confocal light sheet microscopy, CLSM) capable of obtaining micron-scale reconstruction of entire mouse brains labeled with enhanced green fluorescent protein (EGFP). Combining light sheet illumination and confocal detection, CLSM allows deep imaging inside macroscopic cleared specimens with high contrast and speed. Here we describe the complete experimental pipeline to obtain comprehensive and human-readable images of entire mouse brains labeled with fluorescent proteins. The clearing and the mounting procedures are described, together with the steps to perform an optical tomography on its whole volume by acquiring many parallel adjacent stacks. We showed the usage of open-source custom-made software tools enabling stitching of the multiple stacks and multi-resolution data navigation. Finally, we illustrated some example of brain maps: the cerebellum from an L7-GFP transgenic mouse, in which all Purkinje cells are selectively labeled, and the whole brain from a thy1-GFP-M mouse, characterized by a random sparse neuronal labeling.

In this article we describe the full experimental procedure to reconstruct, with high resolution, the fine brain anatomy of fluorescently labeled mouse brains. The described protocol includes sample preparation and clearing, specimen mounting for imaging, data post-processing and multi-scale visualization.

Understanding the architecture of mammalian  brain at single-cell resolution is one of the key issues of neuroscience.  However, mapping neuronal soma and ...

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. ...

A Custom Multiphoton Microscopy Platform for Live Imaging of Mouse Cornea and Conjunctiva

Conventional histological analysis and cell culture systems are insufficient to simulate in vivo physiological and pathological dynamics completely. Multiphoton microscopy (MPM) has become one of the most popular imaging modalities for biomedical study at cellular levels in vivo, advantages include high resolution, deep tissue penetration and minimal phototoxicity. We have designed an MPM imaging platform with a customized mouse eye holder and a stereotaxic stage for imaging ocular surface in vivo. Dual fluorescent protein reporter mouse enables visualization of cell nuclei, cell membranes, nerve fibers, and capillaries within the ocular surface. In addition to multiphoton fluorescence signals, acquiring second harmonic generation (SHG) simultaneously allows for the characterization of collagenous stromal architecture. This platform can be employed for intravital imaging with accurate positioning across the entire ocular surface, including cornea and conjunctiva.

Presented here is a multiphoton microscopic platform for live mouse ocular surface imaging. Fluorescent transgenic mouse enables the visualization of cell nuclei, cell membranes, nerve fibers and capillaries within the ocular surface. Non-linear second harmonic generation signals derived from collagenous structures provide label-free imaging for stromal architectures.

Conventional histological analysis and cell culture systems are insufficient to simulate in vivo physiological and pathological dynamics completely. ...

Bioengineering

Microsurgical Dissection and Tissue Clearing for High Resolution Intact Whole Retina and Vitreous Imaging

Neuronal and vascular structures of the retina in physiologic and pathologic conditions can be better visualized and characterized by using intact whole retina imaging techniques compared to conventional retinal flat mount preparations and sections. However, immunofluorescent imaging of intact whole retina is hindered by the opaque coatings of the eyeball, i.e., sclera, choroid, and retinal pigment epithelium (RPE) and the light scattering properties of retinal layers that prevent full thickness high resolution optical imaging. Chemical bleaching of the pigmented layers and tissue clearing protocols have been described to address these obstacles; however, currently described methods are not suitable for imaging endogenous fluorescent molecules such as green fluorescent protein (GFP) in intact whole retina. Other approaches bypassed this limitation by surgical removal of pigmented layers and the anterior segment of the eyeball allowing intact eye imaging, though the peripheral retina and hyaloid structures were disrupted. Presented here is an intact whole retina and vitreous immunofluorescent imaging protocol that combines surgical dissection of the sclera/choroid/retina pigment epithelium (RPE) layers with a modified tissue clearing method and light sheet fluorescent microscopy (LSFM). The new approach offers an unprecedented view of&#160;unperturbed vascular and neuronal elements of the retina as well as the vitreous and hyaloid vascular system in pathologic conditions.

Presented here is a protocol for intact whole retina imaging in which the outer opaque/pigmented layers of the eyeball are surgically removed, and optical clearing is applied to render retina transparent enabling the visualization of the peripheral retina and hyaloid vasculature in intact retina using light sheet fluorescent microscopy.

Neuronal and vascular structures of the retina in physiologic and pathologic conditions can be better visualized and characterized by using intact whole ...

Microdissection of the Rodent Eye

The ocular micro-dissection of the rodent eye involves the segmentation of the enucleated eyeball with the attached nictitating membrane, or third eyelid, to obtain the anterior and posterior eyecups. With this technique, the sub-parts of the eye, including the corneal tissue, neural tissue, retinal pigment epithelial (RPE) tissue, and lens, can be obtained for wholemounts, cryo-sectioning, and/or single-cell suspensions of a specific ocular tissue. The presence of the third eyelid presents unique and significant advantages, as it benefits the maintenance of the orientation of the eye, which is important for understanding eye physiology following any localized intervention or in studies involving ocular analysis relating to the eye&#39;s spatial topography.
In this method, we enucleated the eyeball at the socket along with the third eyelid by carefully and slowly cutting through the extraocular muscles and severing the optic nerve. The eyeball was pierced through the corneal limbus using a microblade.&#160;The incision was used as the point of entry, allowing for cutting along the corneal-scleral junction by inserting micro-scissors through the incision point. Small and continuous cuts along the circumference were made until the cups separated. These could be further dissected by gently peeling the translucent layer of the neural retina using Colibri suturing forceps to obtain the neural retina and RPE layers. Further, three/four equidistant cuts were made from the periphery perpendicularly to the optic center until the optic nerve was reached. This opened the hemispherical cups into a floret shape so that they fell flat and could be easily mounted. This technique has been used in our lab for corneal wholemounts and retinal sections. The presence of the third eyelid delineates the nasal-temporal orientation, which allows for the study of various cell therapy interventions post-transplantation and, thus, the targeted physiological validation vital for visualization and accurate representation in such studies.

This paper presents a protocol for ocular micro-dissection in rodents. The process involves the enucleation of the eyeball along with the nictitating membrane (i.e., the third eyelid). This is then followed by the separation of the posterior and anterior eye cups.

The ocular micro-dissection of the rodent eye involves the segmentation of the enucleated eyeball with the attached nictitating membrane, or third eyelid, ...

Medicine

Cryosectioning and Immunostaining of Mouse Retina

Tissue sectioning and immunohistochemistry are essential techniques in histological and pathological studies of retinal diseases using animal models. These methods enable detailed examinations of tissue morphologies and the localization of specific proteins within the tissue, which provide valuable insights into disease processes and mechanisms. Mice are the most widely used model for this purpose. However, because mouse eyeballs are small and mouse retinas are extremely delicate tissues, obtaining high-quality retinal sections and immunostaining images from mouse eyeballs is typically challenging. This study describes an improved protocol for cryosectioning mouse retinas and performing immunohistochemistry. An essential point of this protocol involves coating the eyeball with a layer of super glue, which prevents deformation of the eyeballs during the processes of cornea removal, lens extraction, and embedding. This step ensures the integrity of retinal morphologies is well preserved. This protocol highlights critical technical considerations and optimization strategies for consistently producing high-quality retinal sections and achieving excellent immunostaining results.

A protocol for preparing mouse retinal cryosections and performing immunostaining on photoreceptors is described. This article enables researchers to consistently produce mouse retinal frozen sections with well-preserved morphology and high-quality immunostaining results.

Tissue sectioning and immunohistochemistry are essential techniques in histological and pathological studies of retinal diseases using animal models. ...

Mesoscopic Optical Imaging of Whole Mouse Heart

Both genetic and non-genetic cardiac diseases can cause severe remodeling processes in the heart. Structural remodeling, such as collagen deposition (fibrosis) and cellular misalignment, can affect electrical conduction, introduce electromechanical dysfunctions and, eventually, lead to arrhythmia. Current predictive models of these functional alterations are based on non-integrated and low-resolution structural information. Placing this framework on a different order of magnitude is challenging due to the inefficacy of standard imaging methods in performing high-resolution imaging in massive tissue. In this work, we describe a methodological framework that allows imaging of whole mouse hearts with micrometric resolution. The achievement of this goal has required a technological effort where advances in tissue transformation and imaging methods have been combined. First, we describe an optimized CLARITY protocol capable of transforming an intact heart into a nanoporous, hydrogel-hybridized, lipid-free form that allows high transparency and deep staining. Then, a fluorescence light-sheet microscope able to rapidly acquire images of a mesoscopic field of view (mm-scale) with the micron-scale resolution is described. Following the mesoSPIM project, the conceived microscope allows the reconstruction of the whole mouse heart with micrometric resolution in a single tomographic scan. We believe that this methodological framework will allow clarifying the involvement of the cytoarchitecture disarray in the electrical dysfunctions and pave the way for a comprehensive model that considers both the functional and structural data, thus enabling a unified investigation of the structural causes that lead to electrical and mechanical alterations after the tissue remodeling.

We report a method for mesoscopic reconstruction of the whole mouse heart by combining new advancements in tissue transformation and staining with the development of an axially scanned light-sheet microscope.

Both genetic and non-genetic cardiac diseases can cause severe remodeling processes in the heart. Structural remodeling, such as collagen deposition ...

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

Summary

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Chapters in this video

Intravital Microscopy of the Mouse Brain Microcirculation using a Closed Cranial Window

Whole-mount Confocal Microscopy for Adult Ear Skin: A Model System to Study Neuro-vascular Branching Morphogenesis and Immune Cell Distribution

Visualizing the Node and Notochordal Plate In Gastrulating Mouse Embryos Using Scanning Electron Microscopy and Whole Mount Immunofluorescence

Micron-scale Resolution Optical Tomography of Entire Mouse Brains with Confocal Light Sheet Microscopy

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

A Custom Multiphoton Microscopy Platform for Live Imaging of Mouse Cornea and Conjunctiva

Microsurgical Dissection and Tissue Clearing for High Resolution Intact Whole Retina and Vitreous Imaging

Microdissection of the Rodent Eye

Cryosectioning and Immunostaining of Mouse Retina

Mesoscopic Optical Imaging of Whole Mouse Heart