Name: imaging neurons within thick brain sections using the golgi-cox method
Uploaded: 2017-04-18T13:00:00
Description: Watch this Scientific Journal Video about Imaging Neurons within Thick Brain Sections Using the Golgi-Cox Method at JoVE.com

This work was supported by a Discovery Grant to CDCB from the Natural Sciences and Engineering Research Council of Canada (NSERC), a John R. Evans Leaders Fund research infrastructure grant to CDCB from Canada Foundation for Innovation (CFI project number 30381), and by a Discovery Grant to NJM from NSERC. ELL was supported by an Ontario Graduate Scholarship and by an OVC Scholarship from the Ontario Veterinary College at the University of Guelph. CDS was supported by an Undergraduate Student Research Assistantship from NSERC. ALM was supported by an Alexander Graham Bell Scholarship from NSERC and by an OVC Scholarship from the Ontario Veterinary College at the University of Guelph.

Adult female CD1-strain mice were used in this study. Similar staining can be accomplished using both sexes at various ages. Experimental animals were cared for according to the principles and guidelines of the Canadian Council on Animal Care, and the experimental protocol was approved by the University of Guelph Animal Care Committee.
1. Golgi-Cox Staining
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	<li>Golgi-Cox Impregnation of Brains
	<ol>
		<li>Make the Golgi-Cox solution of 1% (w/v) potassium dichromate, 0....

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We describe here a Golgi-Cox staining protocol along with two useful variants for visualizing neurons within thick brain sections. As shown in the Representative Results, the use of a high-resolution objective that has a long 800 &#956;m working distance allows for the reliable visualization of entire neurons throughout the depth of brain sections cut at 500 &#956;m. This study of relatively thick brain sections increases the probability that stained neurons of any type are fully contained within the slice, which is espe...

The visualization of individual neurons within tissue samples allows for the in situ analysis of neuron morphological characteristics, which has significantly advanced our understanding of the brain and how it may be influenced by endogenous disease or exogenous environmental factors. The Golgi-Cox staining method is a cost-effective, relatively simple means of staining a random sample of neurons within the brain. First developed by Golgi1 and modified by Cox2 in the 1800s, researchers have further refined this technique over the years to produce clear, well-stained neurons that can be used to visualize and quantify both dendritic tree morphology and spine density3,4,5,6,7,8,9.
A major technical consideration for the visualization of stained neurons within brain sections is the maximum slice thickness, which is limited by the working distance of available high-magnification/high-resolution microscope objectives. Common oil-immersion objectives in the 60 - 100X range provide excellent resolution, but are limited by their working distances that are typically no greater than 200 &#956;m. Brain sections cut at the 200 &#956;m range may be adequate to visualize certain neuron types that can be contained within this slice thickness, for example pyramidal neurons in shallow layers of the cerebral cortex10,11,12, pyramidal neurons in the CA1 region of the hippocampus13,14, and granule cells in the dentate gyrus of the hippocampus15. Neurons with relatively longer dendritic&#160;trees, such as pyramidal neurons within deep layers of the cerebral cortex that for mouse can extend more than 800 &#956;m from the cell body16, provide a greater challenge because brains would need to be sectioned at a perfect angle to contain the entire dendrite tree within 200 &#956;m slices. This may not even be feasible if a dendrite or any of its branches extend in the rostral or caudal direction. While it is possible to address this limitation by tracing a neuron across multiple adjacent brain sections, this approach introduces a significant technical challenge in aligning the sections accurately for tracing17. A more practical approach would be to visualize entire neurons contained within brain sections that are cut at a greater thickness.
We report here a technique to stain neurons within 400 - 500 &#956;m thick brain sections of mice using the Golgi-Cox method, and to visualize their morphology using a high-resolution silicon oil-immersion objective that has an 800 &#956;m working distance. The Golgi-Cox impregnation and processing protocol that we describe is modified from one of the most-cited modern protocols in the literature6. Our approach with thick brain sections provides the advantage of increasing the probability of identifying neurons of any type that are fully contained within the section. In addition to the main protocol, we also present two variations that provide unique advantages: (1) Golgi-Cox staining with the cresyl violet counterstain on the surface of mounted sections, in order to define boundaries of brain regions and to identify layers of the cerebral cortex, and (2) Golgi-Cox staining with an intermediate freezing step for the long-term storage of impregnated whole brains prior to sectioning and final processing.

<table border="0" cellpadding="0" cellspacing="0" width="775">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">potassium dichromate</td>
			<td style="width:147px;">Fisher Scientific</td>
			<td style="width:175px;">P188-100</td>
			<td style="width:167px;">Hazardous</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">potassium chromate</td>
			<td style="width:147px;">Fisher Scientific</td>
			<td style="width:175px;">P220-100</td>
			<td>Hazardous</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">mercuric chloride</td>
			<td style="width:147px;">Fisher Scientific</td>
			<td style="width:175px;">S25423</td>
			<td>Hazardous</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">Whatman grade 1 filter paper</td>
			<td style="width:147px;">Fisher Scientific</td>
			<td style="width:175px;">1001-185</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:288px;">isoflurane</td>
			<td style="width:147px;">Pharmaceutical Partners of Canada</td>
			<td style="width:175px;">CP0406V2</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">20 mL scintillation vial</td>
			<td style="width:147px;">Fisher Scientific</td>
			<td style="width:175px;">03-337-4</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">sucrose</td>
			<td style="width:147px;">Bioshop Canada</td>
			<td style="width:175px;">SUC700.1</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">sodium phosphate monobasic</td>
			<td style="width:147px;">Sigma Aldrich</td>
			<td style="width:175px;">S5011-500G</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">sodium phosphate dibasic</td>
			<td style="width:147px;">Sigma Aldrich</td>
			<td style="width:175px;">S9390-500G</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">50 mL conical tube</td>
			<td style="width:147px;">Fisher Scientific</td>
			<td style="width:175px;">12-565-271</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">isopentane</td>
			<td style="width:147px;">Fisher Scientific</td>
			<td style="width:175px;">AC126470010</td>
			<td>also known as 2-methylbutane. Hazardous</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">agar</td>
			<td style="width:147px;">Sigma Aldrich</td>
			<td style="width:175px;">A1296-100G</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">small weigh dish</td>
			<td style="width:147px;">Fisher Scientific</td>
			<td style="width:175px;">02-202-100</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">vibratome</td>
			<td style="width:147px;">Leica</td>
			<td style="width:175px;">VT1000 S</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">6 well tissue culture plates</td>
			<td style="width:147px;">Fisher Scientific</td>
			<td style="width:175px;">08-772-1b</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">mesh bottom tissue culture inserts</td>
			<td style="width:147px;">Fisher Scientific</td>
			<td style="width:175px;">07-200-214</td>
			<td></td>
		</tr>
		<tr height="68">
			<td height="68" style="height:68px;width:288px;">paraformadelhyde, 16%</td>
			<td style="width:147px;">Electron Microscope Sciences</td>
			<td style="width:175px;">15710-S</td>
			<td>Hazardous</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">ammonium hydroxide</td>
			<td style="width:147px;">Fisher Scientific</td>
			<td style="width:175px;">A669S-500</td>
			<td>Hazardous</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">Kodak Fixative A</td>
			<td style="width:147px;">Sigma Aldrich</td>
			<td style="width:175px;">P7542</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">superfrost plus slides</td>
			<td style="width:147px;">Fisher Scientific</td>
			<td style="width:175px;">12-550-15</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">CitroSolv clearing agent</td>
			<td style="width:147px;">Fisher Scientific</td>
			<td style="width:175px;">22-143-975</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">anhydrous ethyl alcohol</td>
			<td style="width:147px;">Commercial Alcohols</td>
			<td style="width:175px;">N/A</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">cresyl violet</td>
			<td style="width:147px;">Sigma Aldrich</td>
			<td style="width:175px;">C1791</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">permount</td>
			<td style="width:147px;">Fisher Scientific</td>
			<td style="width:175px;">SP15-100</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">upright microscope</td>
			<td style="width:147px;">Olympus</td>
			<td style="width:175px;">BX53 model</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:288px;">colour camera, 12 bit</td>
			<td style="width:147px;">MBF Biosciences</td>
			<td style="width:175px;">DV-47d</td>
			<td style="width:167px;">QImaging part 01-MBF-2000R-F-CLR-12</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:43px;width:288px;">three-dimensional motorized microscope stage, controller and enoders</td>
			<td style="width:147px;">MBF Biosciences</td>
			<td style="width:175px;">N/A</td>
			<td>Supplied and integrated with microscope by MBF Biosciences</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">4x microscope objective</td>
			<td style="width:147px;">Olympus</td>
			<td style="width:175px;">4x 0.16 N.A. UplanSApo</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">10x microscope objective</td>
			<td style="width:147px;">Olympus</td>
			<td style="width:175px;">10x 0.3 N.A. UPlan FL N&#160;</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">30x microscope objective</td>
			<td style="width:147px;">Olympus</td>
			<td>30x 1.05 N.A. UPlanSApo&#160;</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">60x microscope objective</td>
			<td style="width:147px;">Olympus</td>
			<td>60x 1.42 N.A. PlanAPO N</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">silicone immersion oil</td>
			<td style="width:147px;">Olympus</td>
			<td style="width:175px;">Z-81114</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">Neurolucida software</td>
			<td style="width:147px;">MBF Biosciences</td>
			<td style="width:175px;">Version 10</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:288px;">ImageJ software</td>
			<td style="width:147px;">U. S. National Institutes of Health</td>
			<td style="width:175px;">Current version</td>
			<td>With the OME Bio-Formats plugin installed</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:288px;">Photoshop software</td>
			<td style="width:147px;">Adobe</td>
			<td style="width:175px;">version CS6</td>
			<td></td>
		</tr>
	</tbody>
</table>

imaging neurons within thick brain sections using the golgi-cox method

The Golgi-Cox method of neuron staining has been employed for more than two hundred years to advance our understanding of neuron morphology within histological brain samples. While it is preferable from a practical perspective to prepare brain sections at the greatest thickness possible, in order to increase the probability of identifying stained neurons that are fully contained within single sections, this approach is limited from a technical perspective by the working distance of high-magnification microscope objectives. We report here a protocol to stain neurons using the Golgi-Cox method in mouse brain sections that are cut at 500 &#956;m thickness, and to visualize neurons throughout the depth of these sections using an upright microscope fitted with a high-resolution 30X 1.05 N.A. silicone oil-immersion objective that has an 800 &#956;m working distance. We also report two useful variants of this protocol that may be employed to counterstain the surface of mounted brain sections with the cresyl violet Nissl stain, or to freeze whole brains for long-term storage prior to sectioning and final processing. The main protocol and its two variants produce stained thick brain sections, throughout which full neuron dendritic&#160;trees and dendrite spines may be reliably visualized and quantified.

This Golgi-Cox staining protocol and its two described optional variants may be employed to visualize individual neurons within 400 - 500 &#956;m thick brain sections. Representative image montages of two-dimensional Z-projections captured using a 10X&#160;objective and 5 &#956;m steps in the Z axis are shown in Figure 1: A1 - C1&#160;for a large area of coronal brain sections that includes the anterior cingulate cortex area 1 and the secondary motor cortex<sup class="xre...

Watch this Scientific Journal Video about Imaging Neurons within Thick Brain Sections Using the Golgi-Cox Method at JoVE.com

Imaging Neurons within Thick Brain Sections Using the Golgi-Cox Method

We present a protocol for using the Golgi-Cox staining method in thick brain sections, in order to visualize neurons with long dendritic&#160;trees contained within single tissue samples. Two variants of this protocol are also presented that involve cresyl violet counterstaining, and the freezing of unprocessed brains for long-term storage.

The Golgi-Cox method of neuron staining has been employed for more than two hundred years to advance our understanding of neuron morphology within ...

The overall goal of this procedure is to determine the effective treatment on the morphology of neurons within the rodent brain. This method can help answer key questions in the neuroscience field, such as the normal and experimentally manipulated morphology of neurons within various regions of the rodent brain. This technique has the main advantage of increasing the probability of identifying and visualizing labeled neurons that are fully contained within single histological samples.Begin Golgi-Cox staining by placing a fresh mouse brain into a 20 milliliter glass scintillation vial containing 17 milliliters of Golgi-Cox solution. Protect from light and incubate at room temperature for 25 days. Once the incubation period has elapsed, Cryo protect the brain by placing it into 40 milliliters of sucrose cryoprotectant, and incubate in the dark at 4 degrees celsius for 24 hours.After the incubation in sucrose, the brain can be frozen for long term storage, or immediately blocked for sectioning as shown in the next section of the video. If freezing, immerse the whole brain in 200 milliliters of isopentane that has been precooled on dry ice. Then place the frozen brain into a 50 milliliter conical tube and store in the dark at minus 80 degrees celsius.To begin, remove the brain from the sucrose. And use a razor blade to block it for sectioning by cutting off the cerebellum, leaving a flat edge at the remaining caudal end of the brain. The brain must be blocked at a precise angle to ensure that neurons of interest are fully contained within the resulting sections.For example, we use a coronal plane that is perfectly perpendicular to the rostral caudal axis for cerebral cortex pyramidal neurons. Next, heat agar until it is melted. And let it cool until it is slightly above it's melting point.Then place the brain in a small disposable weigh dish with the caudal end face down. Add melted agar to cover the brain. Once the agar has solidified, trim the excess leaving approximately two to four milliliters surrounding the brain.Then use a small amount of ethyl cyanoacrylate to glue the brain to the stage of a vibratome with it's caudal end face down. Fill the stage area of the vibratome with enough sucrose cryoprotectant to cover the brain. Then section the brain at a slice thickness of 400 to 500 microns depending on the brain region to be examined.Using a small paintbrush, place brain sections into wells of a six well tissue culture plate containing mesh bottom inserts and pre filled with sucrose M phosphate buffer. Protect from light during sectioning. When finished sectioning, incubate sections in the dark at four degrees celsius overnight.Using the mesh bottom inserts, transfer the brain sections into a new well containing five milliliters of paraformaldehyde in phosphate buffer. Incubate on a rocker moving at slow speed at room temperature in the dark for 15 minutes. Wash the sections twice by transferring them to new wells containing five milliliters of water.Protect from light and wash at moderate speed at room temperature for five minutes. Next, transfer the sections into a new well containing five milliliters of ammonium hydroxide. Rock slowly in the dark at room temperature for 15 minutes.Wash the sections twice by transferring them to new wells containing five milliliters of water. Wash on a rocker moving at moderate speed in the dark at room temperature for five minutes. Transfer the sections into a new well containing five milliliters of diluted fixative A.Incubate on a rocker moving at slow speed in the dark at room temperature for 25 minutes.Wash sections in water twice as before. Use a small paintbrush to place the sections onto a microscope slide, then use tweezers and a small tissue to remove excess water and agar. Ensure that all agar is removed before proceeding with the dehydration steps.Allow the sections to air dry at room temperature protected from light. The appropriate drying time is critical, and should be determined for each laboratory. Too short of a time may result in sections falling off of slides during dehydration steps, and too long of a time may result in sections cracking.After drying, first place the slides in clearing agent for five minutes. Place the slides in 100%ethanol for five minutes. Then move through a series of decreasing ethanol concentrations for two minutes each.After the alcohol series, place the slides in water for five minutes. Then stain in 0.5%cresyl violet in water for seven minutes. Following the cresyl violet stain, place the slides in water for two minutes.Then dehydrate the sections through a series of increasing alcohol concentrations for two minutes each. Then two washes in 100%ethanol for five minutes. Place in clearing agent for five minutes.Finally add an anhydrous mounting medium and place a cover slip over the sections. Allow slides to dry horizontally in the dark at room temperature for at least five days. To capture high resolution image stacks of the area containing the neuron of interest, first select a 30X 1.05 NA silicone oil immersion objective on the microscope.Then apply three to four drops of silicone immersion oil to the slide. Place the objective over the slide, ensuring contact between the objective and oil. Focus the image and adjust the camera settings including the exposure and white balance in the imaging software.Next set the upper and lower boundaries for the image stack by focusing to the top of the section, and then selecting set next to top of stack within the image acquisition window. Then focus to the bottom of the section, and select set next to bottom of stack. Set the step distance to one micron by entering one micron next to distance between images within the image acquisition window.Lastly capture the image stack by selecting acquire image stack within the image acquisition window. Repeat the previous steps to capture the entire area of interest making sure that all image stacks overlap by at least 10%in the X and Y axes. This image of the cerebral cortex is a merged Z projection of image stacks acquired from a 500 micron thick section using a 10X objective.The tissue in this image was processed using the main all fresh protocol. The area indicated by the red square was imaged using a 30X silicon oil immersion objective and a higher resolution merged Z projection image stack was acquired. The red arrow shows the two dimensional Z projection of a neuron that was traced from the high resolution 30X image stack.Here the fresh protocol variant with cresyl violet staining was used. This higher resolution image shows the cresyl violet stained section. And this is the neuron tracing obtained.Finally this tissue was processed using the protocol variant in which brains are frozen following Golgi-Cox impregnation. Here the previously frozen tissue is seen at higher resolution. And once again a high quality two dimensional Z projection of the neuron tracing is obtained.This technique can be completed within one month with the processing and imaging steps completed within the final week. Brains can alternatively be frozen following Golgi-Cox impregnation allowing for the processing and imaging to be completed at a later date. Following this procedure, neuroscientists can use the captured images to perform detailed morphological analyses such as dendrite complexity or spine density in order to determine how an experimental manipulation can alter the structure of neurons within the brain.

imaging-neurons-within-thick-brain-sections-using-the-golgi-cox-method

Research

JoVE Journal

Neuroscience

A Rapid Approach to High-Resolution Fluorescence Imaging in Semi-Thick Brain Slices

A fundamental goal to both basic and clinical neuroscience is to better understand the identities, molecular makeup, and patterns of connectivity that are characteristic to neurons in both normal and diseased brain. Towards this, a great deal of effort has been placed on building high-resolution neuroanatomical maps1-3. With the expansion of molecular genetics and advances in light microscopy has come the ability to query not only neuronal morphologies, but also the molecular and cellular makeup of individual neurons and their associated networks4. Major advances in the ability to mark and manipulate neurons through transgenic and gene targeting technologies in the rodent now allow investigators to 'program' neuronal subsets at will5-6. Arguably, one of the most influential contributions to contemporary neuroscience has been the discovery and cloning of genes encoding fluorescent proteins (FPs) in marine invertebrates7-8, alongside their subsequent engineering to yield an ever-expanding toolbox of vital reporters9. Exploiting cell type-specific promoter activity to drive targeted FP expression in discrete neuronal populations now affords neuroanatomical investigation with genetic precision.
Engineering FP expression in neurons has vastly improved our understanding of brain structure and function. However, imaging individual neurons and their associated networks in deep brain tissues, or in three dimensions, has remained a challenge. Due to high lipid content, nervous tissue is rather opaque and exhibits auto fluorescence. These inherent biophysical properties make it difficult to visualize and image fluorescently labelled neurons at high resolution using standard epifluorescent or confocal microscopy beyond depths of tens of microns. To circumvent this challenge investigators often employ serial thin-section imaging and reconstruction methods10, or 2-photon laser scanning microscopy11. Current drawbacks to these approaches are the associated labor-intensive tissue preparation, or cost-prohibitive instrumentation respectively.
Here, we present a relatively rapid and simple method to visualize fluorescently labelled cells in fixed semi-thick mouse brain slices by optical clearing and imaging. In the attached protocol we describe the methods of: 1) fixing brain tissue in situ via intracardial perfusion, 2) dissection and removal of whole brain, 3) stationary brain embedding in agarose, 4) precision semi-thick slice preparation using new vibratome instrumentation, 5) clearing brain tissue through a glycerol gradient, and 6) mounting on glass slides for light microscopy and z-stack reconstruction (Figure 1).
For preparing brain slices we implemented a relatively new piece of instrumentation called the 'Compresstome' VF-200 (http://www.precisionary.com/products_vf200.html). This instrument is a semi-automated microtome equipped with a motorized advance and blade vibration system with features similar in function to other vibratomes. Unlike other vibratomes, the tissue to be sliced is mounted in an agarose plug within a stainless steel cylinder. The tissue is extruded at desired thicknesses from the cylinder, and cut by the forward advancing vibrating blade. The agarose plug/cylinder system allows for reproducible tissue mounting, alignment, and precision cutting. In our hands, the 'Compresstome' yields high quality tissue slices for electrophysiology, immunohistochemistry, and direct fixed-tissue mounting and imaging. Combined with optical clearing, here we demonstrate the preparation of semi-thick fixed brain slices for high-resolution fluorescent imaging.

Here we describe a rapid and simple method to image fluorescently labeled cells in semi-thick brain slices. By fixing, slicing, and optically clearing brain tissue we describe how standard epifluorescent or confocal imaging can be used to visualize individual cells and neuronal networks within intact nervous tissue.

A fundamental goal to both basic and clinical neuroscience is to better understand the identities, molecular makeup, and patterns of connectivity that are ...

Morphometric Analyses of Retinal Sections

Morphometric analyses of retinal sections have been used in examining retinal diseases. For examples, neuronal cells were significantly lost in the retinal ganglion cell layer (RGCL) in rat models with N-methyl-D-aspartate (NMDA)&#x2013;induced excitotoxicity1, retinal ischemia-reperfusion injury2 and glaucoma3. Reduction of INL and inner plexiform layer (IPL) thicknesses were reversed with citicoline treatment in rats' eyes subjected to kainic acid-mediated glutamate excitotoxicity4. Alteration of RGC density and soma sizes were observed with different drug treatments in eyes with elevated intraocular pressure3,5,6. Therefore, having objective methods of analyzing the retinal morphometries may be of great significance in evaluating retinal pathologies and the effectiveness of therapeutic strategies.
The retinal structure is multi-layers and several different kinds of neurons exist in the retina. The morphometric parameters of retina such as cell number, cell size and thickness of different layers are more complex than the cell culture system. Early on, these parameters can be detected using other commercial imaging software. The values are normally of relative value, and changing to the precise value may need further accurate calculation. Also, the tracing of the cell size and morphology may not be accurate and sensitive enough for statistic analysis, especially in the chronic glaucoma model. The measurements used in this protocol provided a more precise and easy way. And the absolute length of the line and size of the cell can be reported directly and easy to be copied to other files. For example, we traced the margin of the inner and outer most nuclei in the INL and formed a line then using the software to draw a 90 degree angle to measure the thickness. While without the help of the software, the line maybe oblique and the changing of retinal thickness may not be repeatable among individual observers. In addition, the number and density of RGCs can also be quantified. This protocol successfully decreases the variability in quantitating features of the retina, increases the sensitivity in detecting minimal changes.
 	This video will demonstrate three types of morphometric analyses of the retinal sections. They include measuring the INL thickness, quantifying the number of RGCs and measuring the sizes of RGCs in absolute value. These three analyses are carried out with Stereo Investigator (MBF Bioscience &mdash; MicroBrightField, Inc.). The technique can offer a simple but scientific platform for morphometric analyses.

This video demonstrates three types of morphometric analyses of the retina, which include measuring the inner nuclear layer thickness, quantifying the number of retinal ganglion cells (RGCs) and measuring the sizes of RGCs. The technique can offer a simple but scientific platform for morphometric analyses.

Morphometric analyses of retinal sections have been used in examining retinal diseases. For examples, neuronal cells were significantly lost in the ...

Imaging Calcium Responses in GFP-tagged Neurons of Hypothalamic Mouse Brain Slices

Despite an enormous increase in our knowledge about the mechanisms underlying the encoding of information in the brain, a central question concerning the precise molecular steps as well as the activity of specific neurons in multi-functional nuclei of brain areas such as the hypothalamus remain. This problem includes identification of the molecular components involved in the regulation of various neurohormone signal transduction cascades. Elevations of intracellular Ca2+ play an important role in regulating the sensitivity of neurons, both at the level of signal transduction and at synaptic sites.
New tools have emerged to help identify neurons in the myriad of brain neurons by expressing green fluorescent protein (GFP) under the control of a particular promoter. To monitor both spatially and temporally stimulus-induced Ca2+ responses in GFP-tagged neurons, a non-green fluorescent Ca2+ indicator dye needs to be used. In addition, confocal microscopy is a favorite method of imaging individual neurons in tissue slices due to its ability to visualize neurons in distinct planes of depth within the tissue and to limit out-of-focus fluorescence. The ratiometric Ca2+ indicator fura-2 has been used in combination with GFP-tagged neurons1. However, the dye is excited by ultraviolet (UV) light. The cost of the laser and the limited optical penetration depth of UV light hindered its use in many laboratories. Moreover, GFP fluorescence may interfere with the fura-2 signals2. Therefore, we decided to use a red fluorescent Ca2+ indicator dye. The huge Stokes shift of fura-red permits multicolor analysis of the red fluorescence in combination with GFP using a single excitation wavelength. We had previously good results using fura-red in combination with GFP-tagged olfactory neurons3. The protocols for olfactory tissue slices seemed to work equally well in hypothalamic neurons4. Fura-red based Ca2+ imaging was also successfully combined with GFP-tagged pancreatic &beta;-cells and GFP-tagged receptors expressed in HEK cells5,6. A little quirk of fura-red is that its fluorescence intensity at 650 nm decreases once the indicator binds calcium7. Therefore, the fluorescence of resting neurons with low Ca2+ concentration has relatively high intensity. It should be noted, that other red Ca2+-indicator dyes exist or are currently being developed, that might give better or improved results in different neurons and brain areas.

In this protocol, we update recent progress in imaging Ca2+ signals of GFP-tagged neurons in brain tissue slices using a red fluorescent Ca2+ indicator dye.

Despite an enormous increase in our knowledge about the mechanisms underlying the encoding of information in the brain, a central question concerning the ...

Visualizing the Effects of a Positive Early Experience, Tactile Stimulation, on Dendritic Morphology and Synaptic Connectivity with Golgi-Cox Staining

To generate longer-term changes in behavior, experiences must be producing stable changes in neuronal morphology and synaptic connectivity. Tactile stimulation is a positive early experience that mimics maternal licking and grooming in the rat. Exposing rat pups to this positive experience can be completed easily and cost-effectively by using highly accessible materials such as a household duster. Using a cross-litter design, pups are either stroked or left undisturbed, for 15 min, three times per day throughout the perinatal period. To measure the neuroplastic changes related to this positive early experience, Golgi-Cox staining of brain tissue is utilized. Owing to the fact that Golgi-Cox impregnation stains a discrete number of neurons rather than all of the cells, staining of the rodent brain with Golgi-Cox solution permits the visualization of entire neuronal elements, including the cell body, dendrites, axons, and dendritic spines. The staining procedure is carried out over several days and requires that the researcher pay close attention to detail. However, once staining is completed, the entire brain has been impregnated and can be preserved indefinitely for ongoing analysis. Therefore, Golgi-Cox staining is a valuable resource for studying experience-dependent plasticity.

This paper describes the procedures for tactile stimulation of rat pups and subsequent Golgi-Cox staining of neuronal morphology. Tactile stimulation is a positive experience that is administered in the perinatal period by stroking pups with a household duster. Golgi-cox staining is a reliable procedure permitting the visualization of entire neurons.

To generate longer-term changes in behavior, experiences must be producing stable changes in neuronal morphology and synaptic connectivity. Tactile ...

Juxtacellular Monitoring and Localization of Single Neurons within Sub-cortical Brain Structures of Alert, Head-restrained Rats

There are a variety of techniques to monitor extracellular activity of single neuronal units. However, monitoring this activity from deep brain structures in behaving animals remains a technical challenge, especially if the structures must be targeted stereotaxically. This protocol describes convenient surgical and electrophysiological techniques that maintain the animal&#8217;s head in the stereotaxic plane and unambiguously isolate the spiking activity of single neurons. The protocol combines head restraint of alert rodents, juxtacellular monitoring with micropipette electrodes, and iontophoretic dye injection to identify the neuron location in post-hoc histology. While each of these techniques is in itself well-established, the protocol focuses on the specifics of their combined use in a single experiment. These neurophysiological and neuroanatomical techniques are combined with behavioral monitoring. In the present example, the combined techniques are used to determine how self-generated vibrissa movements are encoded in the activity of neurons within the somatosensory thalamus. More generally, it is straightforward to adapt this protocol to monitor neuronal activity in conjunction with a variety of behavioral tasks in rats, mice, and other animals. Critically, the combination of these methods allows the experimenter to directly relate anatomically-identified neurophysiological signals to behavior.

This protocol describes the design and surgical implantation of a head-restraining mechanism to monitor neuronal activity in sub-cortical brain structures in alert rats. It delineates procedures to isolate single neurons in the juxtacellular configuration and to efficiently identify their anatomical locations.

There are a variety of techniques to monitor extracellular activity of single neuronal units. However, monitoring this activity from deep brain structures ...

Imaging Serotonergic Fibers in the Mouse Spinal Cord Using the CLARITY/CUBIC Technique

Long descending fibers to the spinal cord are essential for locomotion, pain perception, and other behaviors. The fiber termination pattern in the spinal cord of the majority of these fiber systems have not been thoroughly investigated in any species. Serotonergic fibers, which project to the spinal cord, have been studied in rats and opossums on histological sections and their functional significance has been deduced based on their fiber termination pattern in the spinal cord. With the development of CLARITY and CUBIC techniques, it is possible to investigate this fiber system and its distribution in the spinal cord, which is likely to reveal previously unknown features of serotonergic supraspinal pathways. Here, we provide a detailed protocol for imaging the serotonergic fibers in the mouse spinal cord using the combined CLARITY and CUBIC techniques. The method involves perfusion of a mouse with a hydrogel solution and clarification of the tissue with a combination of clearing reagents. Spinal cord tissue was cleared in just under two weeks, and the subsequent immunofluorescent staining against serotonin was completed in less than ten days. With a multi-photon fluorescent microscope, the tissue was scanned and a 3D image was reconstructed using Osirix software.

Supraspinal projections are important for pain perception and other behaviors, and serotonergic fibers are one of these fiber systems. The present study focused on the application of the combined CLARITY/CUBIC protocol to the mouse spinal cord in order to investigate the termination of these serotonergic fibers.

Long descending fibers to the spinal cord are essential for locomotion, pain perception, and other behaviors. The fiber termination pattern in the spinal ...

Preparation of Acute Brain Slices Using an Optimized N-Methyl-D-glucamine Protective Recovery Method

This protocol is a practical guide to the N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. Numerous recent studies have validated the utility of this method for enhancing neuronal preservation and overall brain slice viability. The implementation of this technique by early adopters has facilitated detailed investigations into brain function using diverse experimental applications and spanning a wide range of animal ages, brain regions, and cell types. Steps are outlined for carrying out the protective recovery brain slice technique using an optimized NMDG artificial cerebrospinal fluid (aCSF) media formulation and enhanced procedure to reliably obtain healthy brain slices for patch clamp electrophysiology. With this updated approach, a substantial improvement is observed in the speed and reliability of gigaohm seal formation during targeted patch clamp recording experiments while maintaining excellent neuronal preservation, thereby facilitating challenging experimental applications. Representative results are provided from multi-neuron patch clamp recording experiments to assay synaptic connectivity in neocortical brain slices prepared from young adult transgenic mice and mature adult human neurosurgical specimens. Furthermore, the optimized NMDG protective recovery method of brain slicing is compatible with both juvenile and adult animals, thus resolving a limitation of the original methodology. In summary, a single media formulation and brain slicing procedure can be implemented across various species and ages to achieve excellent viability and tissue preservation.

Preparation of Acute Brain Slices Using an Optimized N-Methyl-D-glucamine Protective Recovery Method

This protocol demonstrates the implementation of an optimized N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. A single media formulation is used to reliably obtain healthy brain slices from animals of any age and for diverse experimental applications.

This protocol is a practical guide to the N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. Numerous recent studies have ...

Assessment of Hippocampal Dendritic Complexity in Aged Mice Using the Golgi-Cox Method

Dendritic spines are the protuberances from the neuronal dendritic shafts that contain&#160; excitatory synapses. The morphological and branching variations of the neuronal dendrites within the hippocampus are implicated in cognition and memory formation. There are several approaches to Golgi staining, all of which have been useful for determining the morphological characteristics of dendritic arbors and produce a clear background. The present Golgi-Cox method, (a slight variation of the protocol that is provided with a commercial Golgi staining kit), was designed to assess how a relatively low dose of the chemotherapeutic drug 5-flurouracil (5-Fu) would affect dendritic morphology, the number of spines, and the complexity of arborization within the hippocampus. The 5-Fu significantly modulated the dendritic complexity and decreased the spine density throughout the hippocampus in a region-specific manner. The data presented show that the Golgi staining method effectively stained the mature neurons in the CA1, the CA3, and the dentate gyrus (DG) of the hippocampus. This protocol reports the details for each step so that other researchers can reliably stain tissue throughout the brain with high quality results and minimal troubleshooting.

Here we present a Golgi-Cox protocol in extensive detail. This reliable tissue stain method allows for a high-quality assessment of the cytoarchitecture in the hippocampus, and throughout the entire brain, with minimal troubleshooting.

Dendritic spines are the protuberances from the neuronal dendritic shafts that contain&#160; excitatory synapses. The morphological and branching ...

Determining the Functional Status of the Corticospinal Tract Within One Week of Stroke

High interindividual variability in the recovery of upper limb (UL) function after stroke means it is difficult to predict an individual&#39;s potential for recovery based on clinical assessments alone. The functional integrity of the corticospinal tract is an important prognostic biomarker for recovery of UL function, particularly for those with severe initial UL impairment. This article presents a protocol for evaluating corticospinal tract function within 1 week of stroke. This protocol can be used to select and stratify patients in trials of interventions designed to improve UL motor recovery and outcomes after stroke. The protocol also forms part of the PREP2 algorithm, which predicts UL function for individual patients 3 months poststroke. The algorithm sequentially combines a UL strength assessment, age, transcranial magnetic stimulation, and stroke severity, within a few days of the stroke. The benefits of using PREP2 in clinical practice are described elsewhere. This article focuses on the use of a UL strength assessment and transcranial magnetic stimulation to evaluate corticospinal tract function.

This protocol is for evaluating corticospinal tract function within 1 week of stroke. It can be used to select and stratify patients in trials of interventions designed to improve upper limb motor recovery and outcomes and in clinical practice for predicting upper limb functional outcomes 3 months after stroke.

High interindividual variability in the recovery of upper limb (UL) function after stroke means it is difficult to predict an individual&#39;s potential ...

Investigation of Spatial Interaction Between Astrocytes and Neurons in Cleared Brains

Combining viral vector transduction and tissue clearing using the CLARITY method makes it possible to simultaneously investigate several types of brain cells and their interactions. Viral vector transduction enables the marking of diverse cell types in different fluorescence colors within the same tissue. Cells can be identified genetically by activity or projection. Using a modified CLARITY protocol, the potential sample size of astrocytes and neurons has grown by 2-3 orders of magnitude. The use of CLARITY allows the imaging of complete astrocytes, which are too large to fit in their entirety in slices, and the examination of the somata with all their processes. In addition, it provides the opportunity to investigate the spatial interaction between astrocytes and different neuronal cell types, namely, the number of pyramidal neurons in each astrocytic domain or the proximity between astrocytes and specific inhibitory neuron populations. This paper describes, in detail, how these methods are to be applied.

Combining viral vector transduction and brain clearing using the CLARITY method allows the investigation of a large number of neurons and astrocytes simultaneously.

Combining viral vector transduction and tissue clearing using the CLARITY method makes it possible to simultaneously investigate several types of brain ...