This study was supported by the China Academy of Chinese Medical Sciences Innovation Fund (Project Code no. CI2021A03404) and the National Traditional Chinese Medicine Interdisciplinary innovation Fund (Project Code no. ZYYCXTD-D-202202).

The present study was approved by the Ethics Committee of the Institute of Acupuncture and Moxibustion, China Academy of Chinese Medical Sciences (reference number D2018-04-13-1). All procedures were conducted following the National Institutes of Health Guide for the Care and Use of Laboratory Animals (National Academy Press, Washington, D.C., 1996). Three adult male rats (Sprague-Dawley, weight 230 &#177; 15 g) were used in this study. All animals were housed in a 12 h light/dark cycle with controlled temperature and humidity and allowed free access to food and water.
1. Perfusion and sample preparation
<ol>
	<li>Inject 250 mg/kg of tribromoethanol solution (see Table of Materials) intraperitoneally into the rat to induce euthanasia.</li>
	<li>Once breathing stops, use stainless steel surgical scissors to open the thoracic cavity of the rat. Use 20 G needles to perfuse via the left cardiac ventricle13 at a rate of 3 mL/min with 100 mL of 0.9% normal saline followed by 250-300 mL of 4% paraformaldehyde in 0.1 M phosphate buffer (PB, pH 7.4) (Figure 1A,B).</li>
	<li>After perfusion, use a scalpel to remove the skin of the dorsum and sole from the hind paw.
	<ol>
		<li>For the samples requiring frozen section, post-fix the tissues in 4% paraformaldehyde in 0.1 M PB for 2 h; then cryoprotect the tissues in 25% sucrose in 0.1 M PB for more than 24 h at 4 &#176;C</li>
		<li>For the samples with the thick section that requires tissue clearing, post-fix the tissues in 4% paraformaldehyde in 1x phosphate-buffered saline (1x PBS) for 2 h; then cryoprotect the tissues in 1x PBS at 4 &#176;C (Figure 1C&#8211;E).</li>
	</ol>
	</li>
</ol>
2. Triple fluorescent staining with CGRP, phalloidin, and LYVE1 followed by tissue clearing treatment
NOTE: Triple fluorescent staining with CGRP, phalloidin and LYVE1 was applied to reveal the nerve fibers, blood vessels, and lymphatic vessels in hairy and glabrous skin on tissue sections with a thickness of 300 &#181;m following tissue clearing treatment.
<ol>
	<li>Prepare 2% agarose in 1x&#160;PBS by heating in a micro-oven for 2 min until the agarose dissolves (see&#160;Table of Materials).</li>
	<li>After washing, embed the skin tissues into 2% agarose at 37 &#176;C, and place them inside the icebox for cooling (Figure 1F,G).</li>
	<li>Fix the mounted tissue on the vibratory microtome with the ice water and slice them at a thickness of 500 &#181;m in the transversal direction. Use the following parameters to set the vibration slicer and finish slicing: speed, 0.50 mm/s, and amplitude, 1.5 mm (Figure 1H-K).</li>
	<li>After slicing, remove the agarose from the sections, and store them in a six-well plate with 1x&#160;PBS (pH 7.4) (Figure 1L).</li>
	<li>Incubate the tissue sections in a solution of 2% Triton X-100 in 1x&#160;PBS (TriX-PB) overnight at 4 &#176;C. See Figure 2 for the following procedure.</li>
	<li>Place the tissue sections into the blocking buffer and rotate at 72 rpm on the shaker overnight at 4 &#176;C. 
	NOTE: The blocking buffer composition is 10% normal donkey serum, 1% Triton-X 100, and 0.2% sodium azide in 1x&#160;PBS (see Table of Materials).</li>
	<li>Transfer the tissue sections into the solution containing the primary antibodies of mouse monoclonal anti-CGRP (1:500) and sheep polyclonal anti-LYVE1 antibody (1:500) in dilution buffer in the microcentrifuge tube (see Table of Materials), and rotate on the shaker for 2 days at 4 &#176;C. 
	NOTE: The dilution buffer composition is 1% normal donkey serum, 0.2% Triton-X 100, and 0.2% sodium azide in 1x&#160;PBS.</li>
	<li>Wash the tissue sections twice with washing buffer at room temperature, then keep them on the shaker overnight at 4 &#176;C in washing buffer. 
	NOTE: The washing buffer composition is 0.2% Triton-X 100 in 0.1 M PB (pH 7.4).</li>
	<li>The next day, transfer the tissue sections into the mixed solution containing the secondary antibodies of donkey anti-mouse IgG H&#38;L Alexa-Flour488 (1:500) and donkey anti-sheep IgG H&#38;L Alexa-Flour405 (1:500), as well as Alexa-Flour594 phalloidin (1:1000) (see Table of Materials) in a microcentrifuge tube and rotate at 72 rpm on the shaker for 5 h at 4 &#176;C.</li>
	<li>Wash the tissue sections in a six-well plate with washing buffer for 1 h, twice, at room temperature. Keep the tissue sections in washing buffer on the shaker overnight at 4 &#176;C.</li>
	<li>Transfer the tissue sections into the tissue clearing reagent (see Table of Materials, its volume was five times more than the sample volume) and rotate at 60 rpm on the shaker gently for 1 h at room temperature. 
	NOTE: After this treatment, the tissue sections become clear (Figure 2B).</li>
	<li>Mount the cleared tissues on the slide, circle the tissues with a spacer, and stick the gap with fresh tissue clearing reagent and coverslip.</li>
</ol>
3. Triple fluorescent staining with CGRP, phalloidin, and LYVE1 following the conventional approach
NOTE: As a comparison, the same staining was performed on tissue sections at a thickness of 30 &#181;m following conventional techniques.
<ol>
	<li>Slice the tissue sections at 30 &#181;m on a microtome in the transversal direction and mount them on the slide.</li>
	<li>Add blocking solution with 3% normal donkey serum and 0.3% Triton X-100 in 0.1 M PB and incubate the sections for 30 min at room temperature.</li>
	<li>Remove the blocking solution and incubate the sections with the solution containing the primary antibodies of mouse monoclonal anti-CGRP (1:500) and sheep polyclonal anti-LYVE1 antibody (1:500) in dilution buffer overnight at 4 &#176;C.</li>
	<li>The next day, wash the tissue sections with 0.1 M PB (pH 7.4) three times, add the mixed solution containing the secondary antibodies of donkey anti-mouse IgG H&#38;L Alexa-Flour488 (1:500) and donkey anti-sheep IgG H&#38;L Alexa-Flour405 (1:500), as well as Alexa-Flour594 phalloidin (1:1000) on the sections and incubate for 1 h at room temperature.</li>
	<li>Before microscopic observation, wash the sections in 0.1 M PB (pH 7.4) three times, then cover them with coverslips in 50% glycerin.</li>
</ol>
4. Imaging and analyses
<ol>
	<li>Observe the stained samples under a fluorescent microscope, and then take the images using a confocal microscope.</li>
	<li>Capture 30 images (Z-stacks), each in 10 &#181;m frames, from each 300 &#181;m thick section and integrate a single in-focus image using an image processor of the confocal microscopy system (see Table of Materials).
	<ol>
		<li>Perform the following steps in the image processing software of the confocal setup for three-dimensional (3D) analysis: Set Start Focal Plane | Set End focal plane | Set step size | Choose Depth Pattern | Image Capture | Z series. 
		NOTE: The excitation and emission wavelengths of blue fluorescence signals were 401 nm and 421 nm, respectively. Excitation and emission wavelengths of green fluorescence signals were 499 nm and 519 nm, respectively. Excitation and emission wavelengths of red fluorescence signals were 591 nm and 618 nm, respectively. 152 &#181;m is the diameter of the confocal pinhole (10x objective lens, NA: 0.4). The image capture resolution was 1024 &#215; 1024 pixels.</li>
	</ol>
	</li>
	<li>For the conventional samples (step 3), capture 30 images (Z-stacks) in 1 &#181;m frames from each 30 &#181;m thick section and further treat these images as mentioned above (steps 4.1-4.2).</li>
	<li>Demonstrate the images in the pattern of 3D reconstruction with the image processing system.</li>
	<li>Perform 3D reconstructions by importing Z-stack confocal images into Imaris 9.0 (cell imaging software, see Table of Materials) and create surface renderings based on stain intensities: Add new surfaces | Choose Source Channel | Determine the parameters in the smooth option of Surfaces Detail | Determine the parameters in the threshold option of Absolute Intensity | Classify Surfaces | Finish.</li>
	<li>Acquire data in the surface area of positive nerve fibers with the cell imaging software. Select the rendered image | Statistics | Detailed | Specific Values | Volume.</li>
</ol>

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The authors have nothing to disclose.

The present study provides a detailed demonstration of the cutaneous nerve fibers in the hairy and glabrous skin by using immunofluorescence on thicker tissue sections with clearing treatment and a 3D view to understand the skin innervation better. The antibody incubation time of up to 1-2 days and an overnight cleaning process are important. These two key steps directly affect the immunofluorescence staining effect of thick sections. A further problem was raised from the choice of antibodies, not all of which are suitab...

The skin, the largest organ in the body, serving as a key interface to the environment, is densely innervated by many nerve fibers1,2,3. Although skin innervation has been widely studied previously with various histological methods, such as staining on whole-mount skin and tissue sections4,5,6, the detailed effective demonstration of cutaneous nerve fibers is still a challenge7,8. Given this, the present protocol developed a unique technique to exhibit cutaneous nerve fibers more clearly in the thick tissue section.
Because of the limit by the thickness of sections, the observation of innervated skin nerve fibers is not precise enough to accurately depict the relationship between calcitonin gene-related peptide (CGRP) nerve fibers and local tissues and organs from the acquired image information. The emergence of 3D tissue clearing technology provides a feasible method to solve this problem9,10. The fast development of tissue-clearing approaches has offered many tools for studying tissue structures, entire organs, neuronal projections, and whole animals in recent times11. The transparent skin tissue could be imaged in a much thicker section by confocal microscopy to obtain the data to visualize cutaneous nerve fibers.
In the current study, the plantar and dorsal skin of a rat hindfoot was selected as the two target sites of hairy and glabrous skin3,4,7. To trace the cutaneous nerve fibers at a longer distance, the skin tissue was sliced at the thickness of 300 &#181;m for immunohistochemical and histochemical staining, followed by tissue clearing treatment. CGRP was used to label the sensory nerve fibers12,13. In addition, to highlight the cutaneous nerve fibers on the tissue background, phalloidin and lymphatic vessel endothelial hyaluronan receptor 1 (LYVE1) were further used to label the blood vessels and lymphatic vessels, respectively14,15.
These approaches provided a straightforward method that can be applied to demonstrate a high-resolution view of the cutaneous nerve fibers and also to visualize the spatial correlation among the nerve fibers, blood vessels, and lymphatic vessels in the skin, which may provide much more information to understand the homeostasis of the normal skin and the cutaneous alteration under the pathological conditions.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1x phosphate-buffered saline</td>
			<td>Solarbio Life Sciences</td>
			<td>P1020</td>
			<td>pH 7.2-7.4, 0.01 Mol</td>
		</tr>
		<tr>
			<td>2,2,2-Tribromoethanol</td>
			<td>Sigma Life Science</td>
			<td>T48402-5G</td>
			<td></td>
		</tr>
		<tr>
			<td>Confocal fluorescence microscopy</td>
			<td>Olympus Corporation</td>
			<td>Fluoview FV1200</td>
			<td></td>
		</tr>
		<tr>
			<td>Donkey anti-mouse IgG H&#38;L Alexa-Flour488</td>
			<td>Abcam plc.</td>
			<td>ab150105</td>
			<td></td>
		</tr>
		<tr>
			<td>Donkey anti-sheep IgG H&#38;L Alexa-Flour405</td>
			<td>Abcam plc.</td>
			<td>ab175676</td>
			<td></td>
		</tr>
		<tr>
			<td>EP tube</td>
			<td>Wuxi NEST Biotechnology Co.</td>
			<td>615001</td>
			<td>1.5 mL</td>
		</tr>
		<tr>
			<td>Freezing stage sliding microtome system</td>
			<td>Leica Biosystems</td>
			<td>CM1860</td>
			<td></td>
		</tr>
		<tr>
			<td>Imaris Software</td>
			<td>Oxford Instruments</td>
			<td>v.9.0.1</td>
			<td></td>
		</tr>
		<tr>
			<td>IRIS standard scissor</td>
			<td>WPI (World Precision Instruments Inc.)</td>
			<td>503242</td>
			<td></td>
		</tr>
		<tr>
			<td>iSpacer</td>
			<td>SunJin Lab co.</td>
			<td>IS005</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro forceps-Str</td>
			<td>RWD</td>
			<td>F11020-11</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse monoclonal anti-CGRP antibody</td>
			<td>Santa cruz biotechnology, Inc.</td>
			<td>sc-57053</td>
			<td></td>
		</tr>
		<tr>
			<td>Neutral buffered Formalin</td>
			<td>Solarbio Life Sciences</td>
			<td>G2161</td>
			<td>10%</td>
		</tr>
		<tr>
			<td>Normal donkey serum</td>
			<td>Jackson ImmunoResearch Laboratories</td>
			<td>017-000-12</td>
			<td>10 mL</td>
		</tr>
		<tr>
			<td>Peristaltic pump</td>
			<td>Longer Precision Pump Co., Ltd</td>
			<td>BT300-2J</td>
			<td></td>
		</tr>
		<tr>
			<td>Phalloidin Alexa-Fluor 594</td>
			<td>Thermo Fisher Scientific</td>
			<td>A12381</td>
			<td></td>
		</tr>
		<tr>
			<td>RapiClear 1.52 solution</td>
			<td>SunJin Lab co.</td>
			<td>RC152001</td>
			<td>10 mL</td>
		</tr>
		<tr>
			<td>Regular agarose</td>
			<td>Gene Company Limited</td>
			<td>G-10</td>
			<td></td>
		</tr>
		<tr>
			<td>SEMKEN 1 x 2 Teeth Tissue Forceps-Str</td>
			<td>RWD</td>
			<td>F13038-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Sheep polyclonal anti-LYVE1 antibody</td>
			<td>R&#38;D Systems, Inc.</td>
			<td>AF7939</td>
			<td></td>
		</tr>
		<tr>
			<td>Six-well plate</td>
			<td>Corning Incorporated</td>
			<td>3335</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium azide</td>
			<td>Sigma Life Science</td>
			<td>S2002</td>
			<td>25 g</td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sigma Life Science</td>
			<td>V900116</td>
			<td>500 g</td>
		</tr>
		<tr>
			<td>Super Glue</td>
			<td>Henkel AG &#38; Co.</td>
			<td>Pattex 502</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical Handles</td>
			<td>RWD</td>
			<td>S32003-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Solarbio Life Sciences</td>
			<td>9002-93-1</td>
			<td>100 mL</td>
		</tr>
		<tr>
			<td>Urethane</td>
			<td>Sigma Life Science</td>
			<td>U2500</td>
			<td>500 g</td>
		</tr>
		<tr>
			<td>VANNAS spring scissors</td>
			<td>RWD</td>
			<td>S1014-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Vibratory microtome</td>
			<td>Leica Biosystems</td>
			<td>VT1200S</td>
			<td></td>
		</tr>
	</tbody>
</table>

demonstrating hairy and glabrous skin innervation in a 3d pattern using multiple fluorescent staining and tissue clearing approaches

Skin innervation is an important part of the peripheral nervous system. Although the study of the cutaneous nerve fibers has progressed rapidly, most of the understanding of their distributional and chemical characteristics comes from conventional histochemical and immunohistochemical staining on thin tissue sections. With the development of the tissue clearing technique, it has become possible to view the cutaneous nerve fibers on thicker tissue sections. The present protocol describes multiple fluorescent staining on tissue sections at a thickness of 300 &#181;m from the plantar and dorsal skin of rat hindfoot, the two typical hairy and glabrous skin sites. Here, the calcitonin gene-related peptide labels the sensory nerve fibers, while phalloidin and lymphatic vessel endothelial hyaluronan receptor 1 label the blood and lymphatic vessels, respectively. Under a confocal microscope, the labeled sensory nerve fibers were followed completely at a longer distance, running in bundles in the deep cutaneous layer and freestyle in the superficial layer. These nerve fibers ran in parallel to or surrounded the blood vessels, and lymphatic vessels formed a three-dimensional (3D) network in the hairy and glabrous skin. The current protocol provides a more effective approach to studying skin innervation than the existing conventional methods from the methodology perspective.

After triple fluorescent staining, the nerve fibers, blood vessels, and lymphatic vessels were clearly labeled with CGRP, phalloidin, and LYVE1, respectively, in the hairy and glabrous skin (Figure 3,4). With the clearing treatment, the CGRP-positive nerve fibers, phalloidin-positive blood vessels, and LYVE1-positive lymphatic vessels can be imaged at a greater depth to acquire the complete structural information of the skin (Figure 3). When the...

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Demonstrating Hairy and Glabrous Skin Innervation in a 3D Pattern Using Multiple Fluorescent Staining and Tissue Clearing Approaches

The thickness of tissue sections limited the morphological study of the skin innervation. The present protocol describes a unique tissue clearing technique to visualize cutaneous nerve fibers in thick 300 &#181;m tissue sections under confocal microscopy.

Skin innervation is an important part of the peripheral nervous system. Although the study of the cutaneous nerve fibers has progressed rapidly, most of ...

demonstrating-hairy-glabrous-skin-innervation-3d-pattern-using

Research

JoVE Journal

Neuroscience

2.5K Views.  China Academy of Chinese Medical Sciences. The thickness of tissue sections limited the morphological study of the skin innervation. The present protocol describes a unique tissue clearing technique to visualize cutaneous nerve fibers in thick 300 µm tissue sections under confocal microscopy.

Video: Demonstrating Hairy and Glabrous Skin Innervation in a 3D Pattern Using Multiple Fluorescent Staining and Tissue Clearing Approaches

Cross-Modal Multivariate Pattern Analysis

Multivariate pattern analysis (MVPA) is an increasingly popular method of analyzing functional magnetic resonance imaging (fMRI) data1-4. Typically, the method is used to identify a subject's perceptual experience from neural activity in certain regions of the brain. For instance, it has been employed to predict the orientation of visual gratings a subject perceives from activity in early visual cortices5 or, analogously, the content of speech from activity in early auditory cortices6.
Here, we present an extension of the classical MVPA paradigm, according to which perceptual stimuli are not predicted within, but across sensory systems. Specifically, the method we describe addresses the question of whether stimuli that evoke memory associations in modalities other than the one through which they are presented induce content-specific activity patterns in the sensory cortices of those other modalities. For instance, seeing a muted video clip of a glass vase shattering on the ground automatically triggers in most observers an auditory image of the associated sound; is the experience of this image in the "mind's ear" correlated with a specific neural activity pattern in early auditory cortices? Furthermore, is this activity pattern distinct from the pattern that could be observed if the subject were, instead, watching a video clip of a howling dog?
In two previous studies7,8, we were able to predict sound- and touch-implying video clips based on neural activity in early auditory and somatosensory cortices, respectively. Our results are in line with a neuroarchitectural framework proposed by Damasio9,10, according to which the experience of mental images that are based on memories - such as hearing the shattering sound of a vase in the "mind's ear" upon seeing the corresponding video clip - is supported by the re-construction of content-specific neural activity patterns in early sensory cortices.

Classical multivariate pattern analysis predicts sensory stimuli a subject perceives from neural activity in the corresponding cortices (e.g. visual stimuli from activity in visual cortex). Here, we apply pattern analysis cross-modally and show that sound- and touch-implying visual stimuli can be predicted from activity in auditory and somatosensory cortices, respectively.

Multivariate pattern analysis (MVPA) is an increasingly popular method of analyzing functional magnetic resonance imaging (fMRI) data1-4. Typically, the ...

Tissue Preparation and Immunostaining of Mouse Sensory Nerve Fibers Innervating Skin and Limb Bones

Detection and primary processing of physical, chemical and thermal sensory stimuli by peripheral sensory nerve fibers is key to sensory perception in animals and humans. These peripheral sensory nerve fibers express a plethora of receptors and ion channel proteins which detect and initiate specific sensory stimuli. Methods are available to characterize the electrical properties of peripheral sensory nerve fibers innervating the skin, which can also be utilized to identify the functional expression of specific ion channel proteins in these fibers. However, similar electrophysiological methods are not available (and are also difficult to develop) for the detection of the functional expression of receptors and ion channel proteins in peripheral sensory nerve fibers innervating other visceral organs, including the most challenging tissues such as bone. Moreover, such electrophysiological methods cannot be utilized to determine the expression of non-excitable proteins in peripheral sensory nerve fibers. Therefore, immunostaining of peripheral/visceral tissue samples for sensory nerve fivers provides the best possible way to determine the expression of specific proteins of interest in these nerve fibers. So far, most of the protein expression studies in sensory neurons have utilized immunostaining procedures in sensory ganglia, where the information is limited to the expression of specific proteins in the cell body of specific types or subsets of sensory neurons. Here we report detailed methods/protocols for the preparation of peripheral/visceral tissue samples for immunostaining of peripheral sensory nerve fibers. We specifically detail methods for the preparation of skin or plantar punch biopsy and bone (femur) sections from mice for immunostaining of peripheral sensory nerve fibers. These methods are not only key to the qualitative determination of protein expression in peripheral sensory neurons, but also provide a quantitative assay method for determining changes in protein expression levels in specific types or subsets of sensory fibers, as well as for determining the morphological and/or anatomical changes in the number and density of sensory fibers during various pathological states. Further, these methods are not confined to the staining of only sensory nerve fibers, but can also be used for staining any types of nerve fibers in the skin, bones and other visceral tissue.

Immunocytochemical identification of peripheral sensory nerve fiber subtypes (and detection of protein expression therein) are key to the understanding of molecular mechanisms underlying peripheral sensation. Here we describe methods for preparation of peripheral/visceral tissue samples, such as skin and limb bones, for specific immunostaining of peripheral sensory nerve fibers.

Detection and primary processing of physical, chemical and thermal sensory stimuli by peripheral sensory nerve fibers is key to sensory perception in ...

3D Modeling of the Lateral Ventricles and Histological Characterization of Periventricular Tissue in Humans and Mouse

The ventricular system carries and circulates cerebral spinal fluid (CSF) and facilitates clearance of solutes and toxins from the brain. The functional units of the ventricles are ciliated epithelial cells termed ependymal cells, which line the ventricles and through ciliary action are capable of generating laminar flow of CSF at the ventricle surface. This monolayer of ependymal cells also provides barrier and filtration functions that promote exchange between brain interstitial fluids (ISF) and circulating CSF. Biochemical changes in the brain are thereby reflected in the composition of the CSF and destruction of the ependyma can disrupt the delicate balance of CSF and ISF exchange. In humans there is a strong correlation between lateral ventricle expansion and aging. Age-associated ventriculomegaly can occur even in the absence of dementia or obstruction of CSF flow. The exact cause and progression of ventriculomegaly is often unknown; however, enlarged ventricles can show regional and, often, extensive loss of ependymal cell coverage with ventricle surface astrogliosis and associated periventricular edema replacing the functional ependymal cell monolayer. Using MRI scans together with postmortem human brain tissue, we describe how to prepare, image and compile 3D renderings of lateral ventricle volumes, calculate lateral ventricle volumes, and characterize periventricular tissue through immunohistochemical analysis of en face lateral ventricle wall tissue preparations. Corresponding analyses of mouse brain tissue are also presented supporting the use of mouse models as a means to evaluate changes to the lateral ventricles and periventricular tissue found in human aging and disease. Together, these protocols allow investigations into the cause and effect of ventriculomegaly and highlight techniques to study ventricular system health and its important barrier and filtration functions within the brain.

Using MRI scans (human), 3D imaging software, and immunohistological analysis, we document changes to the brain&#8217;s lateral ventricles. Longitudinal 3D mapping of lateral ventricle volume changes and characterization of periventricular cellular changes that occur in the human brain due to aging or disease are then modeled in mice.

The ventricular system carries and circulates cerebral spinal fluid (CSF) and facilitates clearance of solutes and toxins from the brain. The functional ...

Analysis of Developing Tooth Germ Innervation Using Microfluidic Co-culture Devices

Innervation plays a key role in the development, homeostasis and regeneration of organs and tissues. However, the mechanisms underlying these phenomena are not well understood yet. In particular, the role of innervation in tooth development and regeneration is neglected.
Several in vivo studies have provided important information about the patterns of innervation of dental tissues during development and repair processes of various animal models. However, most of these approaches are not optimal to highlight the molecular basis of the interactions between nerve fibres and target organs and tissues.
Co-cultures constitute a valuable method to investigate and manipulate the interactions between nerve fibres and teeth in a controlled and isolated environment. In the last decades, conventional co-cultures using the same culture medium have been performed for very short periods (e.g., two days) to investigate the attractive or repulsive effects of developing oral and dental tissues on sensory nerve fibres. However, extension of the culture period is required to investigate the effects of innervation on tooth morphogenesis and cytodifferentiation.
Microfluidics systems allow co-cultures of neurons and different cell types in their appropriate culture media. We have recently demonstrated that trigeminal ganglia (TG) and teeth are able to survive for a long period of time when co-cultured in microfluidic devices, and that they maintain in these conditions the same innervation pattern that they show in vivo.
On this basis, we describe how to isolate and co-culture developing trigeminal ganglia and tooth germs in a microfluidic co-culture system.This protocol describes a simple and flexible way to co-culture ganglia/nerves and target tissues and to study the roles of specific molecules on such interactions in a controlled and isolated environment.

Co-cultures represent a valuable method to study the interactions between nerves and target tissues and organs. Microfluidic systems allow co-culturing ganglia and whole developing organs or tissues in different culture media, thus representing a valuable tool for the in vitro study of the crosstalk between neurons and their targets.

Innervation plays a key role in the development, homeostasis and regeneration of organs and tissues. However, the mechanisms underlying these phenomena ...

Quantification of Endosome and Lysosome Motilities in Cultured Neurons Using Fluorescent Probes

In the brain, membrane trafficking systems play important roles in regulating neuronal functions, such as neuronal morphology, synaptic plasticity, survival, and glial communications. To date, numerous studies have reported that defects in these systems cause various neuronal diseases. Thus, understanding the mechanisms underlying vesicle dynamics may provide influential clues that could aid in the treatment of several neuronal disorders. Here, we describe a method for quantifying vesicle motilities, such as motility distance and rate of movement, using a software plug-in for the ImageJ platform. To obtain images for quantification, we labeled neuronal endosome-lysosome structures with EGFP-tagged vesicle marker proteins and observed the movement of vesicles using a time-lapse microscopy. This method is highly useful and simplify measuring vesicle motility in neurites, such as axons and dendrites, as well as in the soma of both neurons and glial cells. Furthermore, this method can be applied to other cell lines, such as fibroblasts and endothelial cells. This approach could provide a valuable advancement of our understanding of membrane trafficking.

The investigation of membrane trafficking is crucial for understanding neuronal functions. Here, we introduce a method for quantifying vesicle motility in neurons. This is a convenient method that can be adapted to the quantification of membrane trafficking in the nervous system.

In the brain, membrane trafficking systems play important roles in regulating neuronal functions, such as neuronal morphology, synaptic plasticity, ...

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

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

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

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

A Rat Model of Central Fatigue Using a Modified Multiple Platform Method

In this article, we introduced a rat model of central fatigue using the modified multiple platform method (MMPM). The Multiple Platform box was designed as a water tank with narrow platforms on the bottom. The model rats were put into the tank and stood on the platforms for 14 h (18:00 - 8:00) per day for a consecutive 21 days, with a blank control group set for contrast. At the end of modeling, rats in the model group showed an obvious fatigued appearance. To assess the model, we performed several behavioral tests, including the open field test (OFT), the elevated plus maze (EPM) test, and the exhaustive swimming (ES) test. The results showed that anxiety, spatial cognition impairment, poor muscle performance, and declined voluntary activity presented in model rats confirm the diagnosis of&#160;central fatigue. Changes of the central neurotransmitters also verified the result. In conclusion, the model successfully simulated central fatigue, and future study with the model may help reveal the pathological mechanism of the disease.

Here, we present a protocol to introduce a rat model of central fatigue using the modified multiple platform method (MMPM).

In this article, we introduced a rat model of central fatigue using the modified multiple platform method (MMPM). The Multiple Platform box was designed ...

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 ...

In vivo Imaging of Fully Active Brain Tissue in Awake Zebrafish Larvae and Juveniles by Skull and Skin Removal

Understanding the ephemeral changes that occur during brain development and maturation requires detailed high-resolution imaging in space and time at cellular and subcellular resolution. Advances in molecular and imaging technologies have allowed us to gain numerous detailed insights into cellular and molecular mechanisms of brain development in the transparent zebrafish embryo. Recently, processes of refinement of neuronal connectivity that occur at later larval stages several weeks after fertilization, which are for example control of social behavior, decision making or motivation-driven behavior, have moved into focus of research. At these stages, pigmentation of the zebrafish skin interferes with light penetration into brain tissue, and solutions for embryonic stages, e.g., pharmacological inhibition of pigmentation, are not feasible anymore.
Therefore, a minimally invasive surgical solution for microscopy access to the brain of awake zebrafish is provided that is derived from electrophysiological&#160;approaches. In teleosts, skin and soft skull cartilage can be carefully removed by micro-peeling these layers, exposing underlying neurons and axonal tracts without damage. This allows for recording neuronal morphology, including synaptic structures and their molecular contents, and the observation of physiological changes such as Ca2+ transients or intracellular transport events. In addition, interrogation of these processes by means of pharmacological inhibition or optogenetic manipulation is feasible. This brain exposure approach provides information about structural and physiological changes in neurons as well as the correlation and interdependence of these events in live brain tissue in the range of minutes or hours. The technique is suitable for in vivo brain imaging of zebrafish larvae up to 30 days post fertilization, the latest developmental stage tested so far. It, thus, provides access to such important questions as synaptic refinement and scaling, axonal and dendritic transport, synaptic targeting of cytoskeletal cargo or local activity-dependent expression. Therefore, a broad use for this mounting and imaging approach can be anticipated.

Here we present a method to image the zebrafish embryonic brain in vivo upto larval and juvenile stages. This microinvasive procedure, adapted from electrophysiological approaches, provides access to cellular and subcellular details of mature neuron and can be combined with optogenetics and neuropharmacological studies for characterizing brain function and drug intervention.

Understanding the ephemeral changes that occur during brain development and maturation requires detailed high-resolution imaging in space and time at ...

Imaging and Quantification of Intact Neuronal Dendrites via CLARITY Tissue Clearing

Brain activity, the electrochemical signals passed between neurons, is determined by the connectivity patterns of neuronal networks, and from the morphology of processes and substructures within these neurons. As such, much of what is known about brain function has arisen alongside developments in imaging technologies that allow further insight into how neurons are organized and connected in the brain. Improvements in tissue clearing have allowed for high-resolution imaging of thick brain slices, facilitating morphological reconstruction and analyses of neuronal substructures, such as dendritic arbors and spines. In tandem, advances in image processing software provide methods of quickly analyzing large imaging datasets. This work presents a relatively rapid method of processing, visualizing, and analyzing thick slices of labeled neural tissue at high-resolution using CLARITY tissue clearing, confocal microscopy, and image analysis. This protocol will facilitate efforts toward understanding the connectivity patterns and neuronal morphologies that characterize healthy brains, and the changes in these characteristics that arise in diseased brain states.

Neuronal dendritic morphology often underlies function. Indeed, many disease processes that affect the development of neurons manifest with a morphological phenotype. This protocol describes a simple and powerful method for analyzing intact dendritic arbors and their associated spines.