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

Summary

We present a protocol for using the Golgi-Cox staining method in thick brain sections, in order to visualize neurons with long dendritic 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.

Abstract

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 μm 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 μm 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 trees and dendrite spines may be reliably visualized and quantified.

Introduction

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 density^{3,4,5,6,7,8,9}.

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 μm. Brain sections cut at the 200 μm 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 cortex^10,11,12, pyramidal neurons in the CA1 region of the hippocampus^13,14, and granule cells in the dentate gyrus of the hippocampus¹⁵. Neurons with relatively longer dendritic trees, such as pyramidal neurons within deep layers of the cerebral cortex that for mouse can extend more than 800 μm 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 μm 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 μm 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 μm 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.

Protocol

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

1. Golgi-Cox Impregnation of Brains
   1. Make the Golgi-Cox solution of 1% (w/v) potassium dichromate, 0.8% (w/v) potassium chromate and 1% (w/v) mercuric chloride by dissolving the potassium dichromate and potassium chromate into high-quality water separately. Mix the solutions, add the mercuric chloride and filter the final solution using grade 1 filter paper. Store the solution in the dark for up to one month.
   2. Anesthetize the mouse using 5% isoflurane.
   3. Euthanize the mouse by decapitation and quickly remove the brain.
   4. Place the brain into a 20 mL glass scintillation vial containing 17 mL of Golgi-Cox solution and incubate in the dark at RT for 25 d.
   5. Cryoprotect the brain by placing it into a 50 mL conical tube containing 40 mL of sucrose cryoprotectant (30% (w/v) sucrose in 0.1 M phosphate buffer, pH 7.4) in the dark at 4 °C for 24 h.
      NOTE: The following optional three steps may be employed as an alternative to the main protocol, in order to freeze the brain at this stage for long-term storage.
   6. Freeze the whole brain by immersing it into 200 mL isopentane that has been precooled on dry ice.
   7. Place the frozen brain into a 50 mL conical tube and store in the dark at -80 °C.
   8. When ready to proceed, thaw the brain by placing it into a 50 mL conical tube containing 40 mL of sucrose cryoprotectant in the dark at 4 °C for 24 h.
2. Brain Sectioning
   1. Remove the brain from the sucrose and block it for sectioning by cutting off the cerebellum using a razor blade and leaving a flat edge at the remaining caudal end of the brain.
   2. Heat agar (3% (w/v) in water) until it is melted and let cool until it is slightly above its melting point.
   3. Place the brain in a small disposable weigh dish with its caudal end face-down and add a sufficient amount of melted agar to cover the brain.
   4. Once the agar has solidified, trim excess agar leaving approximately 2 - 4 mm surrounding the brain and glue the brain to the stage of a vibratome with its caudal end face-down using a small amount of ethyl cyanoacrylate glue.
   5.  Fill the stage area of the vibratome with a sufficient amount of sucrose cryoprotectant to cover the brain, and section the brain at a slice thickness of 400 - 500 µm (depending on the brain region to be examined) using a vibration frequency of 86 Hz and a blade advancement speed of 0.125 mm/s.
   6. Using a small paint brush, place brain sections into a well of a 6-well tissue culture plate containing commercially-available mesh-bottom inserts and pre-filled with 10 mL of 6% (w/v) sucrose in 0.1 M phosphate buffer, pH 7.4.
   7. Incubate sections in the dark at 4 °C O/N.
3. Developing Brain Sections
   1. Using the mesh-bottom inserts, transfer brain sections into a new well containing 5 mL of 2% (w/v) paraformaldehyde (PBD) in 0.1 M phosphate buffer, pH 7.4. Incubate on a rocker moving at slow speed in the dark at RT for 15 min.
   2. Wash sections twice by transferring them to new wells containing 5 mL of water. Wash on a rocker moving at a moderate speed in the dark at RT for 5 min.
   3. Transfer sections into a new well containing 5 mL of 2.7% (v/v) ammonium hydroxide. Incubate on a rocker moving at a slow speed in the dark at RT for 15 min.
   4. Wash sections twice by transferring them to new wells containing 5 mL of water. Wash on a rocker moving at a moderate speed in the dark at RT for 5 min.
   5. Transfer sections into a new well containing 5 mL of Fixative A (see Materials Table) that has been diluted in water 10x from its original purchased concentration. Incubate on a rocker moving at a slow speed in the dark at RT for 25 min.
   6. Wash sections twice by transferring them to new wells containing 5 mL of water. Wash on a rocker moving at a moderate speed in the dark at RT for 5 min.
4. Mounting Brain Sections
   1. Using a small paint brush, mount sections onto microscope slides. Remove excess water and agar using tweezers and a small tissue. Ensure that all agar is removed before proceeding with dehydration.
   2. Allow sections to air dry at RT for approximately 45 min (400 μm sections) or 90 min (500 μm sections).
      NOTE: The timing and proper level of dryness is critical, and may need to be determined in each laboratory depending on ambient temperature and humidity level. Too short a drying time leads to sections falling off of slides during subsequent dehydration steps, and too long a drying time leads to sections cracking. Sections will still appear to be shiny at the appropriate level of dryness.
   3. Stain sections with cresyl violet by placing slides into Coplin staining jars as indicated.
      NOTE: This optional step may be employed for sections that had never been frozen, in order to stain neuronal nuclei with cresyl violet. We have found that clearing and rehydrating sections before incubation in cresyl violet leads to even staining and low background across sections.
      1. Place in clearing agent for 5 min. Repeat once.
      2. Place in 100% ethanol for 5 min. Repeat once.
      3. Place in 95% ethanol in water, then 75% ethanol in water, and then 50% ethanol in water for 2 min each.
      4. Place in water for 5 min.
      5. Place in 0.5% (w/v) cresyl violet in water for 7 min.
      6. Place in water for 2 min. Repeat once.
   4. Dehydrate sections by placing slides into Coplin staining jars as indicated.
      1. Place in 50% ethanol in water, then 75% ethanol in water, and then 95% ethanol in water for 2 min each.
      2. Place in 100% ethanol for 5 min. Repeat once.
      3. Place in clearing agent for 5 min. Repeat once.
         NOTE: These dehydration and clearing times are sufficient to process mouse brains as described in this manuscript. However, we have observed for other species including rat and cowbird, that the final clearing step may need to be extended up to 15 min total.
   5. Coverslip sections using an anhydrous mounting medium.
   6. Allow slides to dry horizontally in the dark at RT for at least 5 d.

2. Imaging Stained Neurons within Thick Brain Sections

1. Capturing Image Stacks
   1. Turn on the microscope light bulb, camera, and stage controller.
   2. Open the microscope software (e.g., Neurolucida).
   3. Place a slide on the microscope stage.
   4. Capture a 2D wide-view image of the brain section using a low magnification objective such as 1.25X 0.4 N.A. PlanAPO or 4X 0.16 N.A.
      1. Focus image and adjust camera settings including the exposure time and white balance.
      2. Create a reference point by left-clicking anywhere on the section
      3. Capture the image by selecting "acquire single image" within the image acquisition window.
   5. Capture mid-resolution image stacks of the area containing the neurons(s) of interest, using a 10X 0.3 N.A. UPlan FL N objective.
      1. Focus the image and adjust camera settings including the exposure and white balance.
      2. Set the upper and lower boundaries for the image stack by focusing to the top of the section and selecting "set" next to "top of stack" within the image acquisition window, and then focusing to the bottom of the section and selecting "set" next to "bottom of stack" within the image acquisition window.
      3. Set the step distance to 5 μm by entering "5 μm" next to "distance between images" within the image acquisition window.
      4. Capture the image stack by selecting "acquire image stack" within the image acquisition window.
      5. Repeat the above 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.
   6. Capture high-resolution image stacks of the area containing the neuron of interest, using a 30X 1.05 N.A. silicone oil-immersion objective.
      1. Apply 3 - 4 drops of silicone immersion oil to the slide and place the objective over the slide, ensuring to make contact between the objective and oil.
      2. Focus the image and adjust camera settings including the exposure and white balance.
      3. Set the upper and lower boundaries for the image stack by focusing to the top of the section and selecting "set" next to "top of stack" within the image acquisition window, and then focusing to the bottom of the section and selecting "set" next to "bottom of stack" within the image acquisition window.
      4. Set the step distance to 1 μm by entering "1 μm" next to "distance between images" within the image acquisition window.
      5. Capture the image stack by selecting "acquire image stack" within the image acquisition window.
      6. Repeat the above 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.
   7. Save the data file and save all image files in TIFF format for external processing.
2. Creating Z-projection Images and Image Montages
   1. Create Z-projection images in ImageJ
      1. Open ImageJ software that has the Bio-Formats plugin installed.
      2. Select "Plugins" -> "Bio-Formats" -> "Bio-Formats Importer".
      3. Select the image stack file to be opened.
      4. Once the file is opened, change the format to RGB by selecting "Image" -> "Type" -> "RGB Color".
      5. Create the Z-projection by selecting "Image" -> "Stacks" -> "Z Project…".
      6. Save two-dimensional Z-projection image as a TIFF file.
   2. Create a two-dimensional image montage of entire area of interest.
      1. Open the software (e.g., Adobe Photoshop).
      2. Select "File" -> "Automate" -> "Photomerge".
      3. Select "Browse" and then add all images files to be merged.
      4. Ensure that "Blend Images Together" is selected and then select "OK".
      5. Save the resulting montage image of the entire area of interest as a TIFF file.

Results

This Golgi-Cox staining protocol and its two described optional variants may be employed to visualize individual neurons within 400 - 500 μm thick brain sections. Representative image montages of two-dimensional Z-projections captured using a 10X objective and 5 μm steps in the Z axis are shown in Figure 1: A1 - C1 for a large area of coronal brain sections that includes the anterior cingulate cortex area 1 and the secondary motor cortex¹⁸. Section...

Discussion

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 μm working distance allows for the reliable visualization of entire neurons throughout the depth of brain sections cut at 500 μm. 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...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
potassium dichromate	Fisher Scientific	P188-100	Hazardous
potassium chromate	Fisher Scientific	P220-100	Hazardous
mercuric chloride	Fisher Scientific	S25423	Hazardous
Whatman grade 1 filter paper	Fisher Scientific	1001-185
isoflurane	Pharmaceutical Partners of Canada	CP0406V2
20 mL scintillation vial	Fisher Scientific	03-337-4
sucrose	Bioshop Canada	SUC700.1
sodium phosphate monobasic	Sigma Aldrich	S5011-500G
sodium phosphate dibasic	Sigma Aldrich	S9390-500G
50 mL conical tube	Fisher Scientific	12-565-271
isopentane	Fisher Scientific	AC126470010	Also known as 2-methylbutane; hazardous
agar	Sigma Aldrich	A1296-100G
small weigh dish	Fisher Scientific	02-202-100
vibratome	Leica	VT1000 S
6-well tissue culture plates	Fisher Scientific	08-772-1b
mesh bottom tissue culture inserts	Fisher Scientific	07-200-214
paraformadelhyde (PFA), 16%	Electron Microscope Sciences	15710-S	Hazardous
ammonium hydroxide	Fisher Scientific	A669S-500	Hazardous
Kodak Fixative A	Sigma Aldrich	P7542
superfrost plus slides	Fisher Scientific	12-550-15
CitroSolv clearing agent	Fisher Scientific	22-143-975
anhydrous ethyl alcohol	Commercial Alcohols	N/A
cresyl violet	Sigma Aldrich	C1791
permount	Fisher Scientific	SP15-100
upright microscope	Olympus	BX53 model
colour camera, 12 bit	MBF Biosciences	DV-47d	QImaging part 01-MBF-2000R-F-CLR-12
3D motorized microscope stage, controller and enoders	MBF Biosciences	N/A	Supplied and integrated with microscope by MBF Biosciences
4X microscope objective	Olympus	4x 0.16 N.A. UplanSApo
10X microscope objective	Olympus	10x 0.3 N.A. UPlan FL N
30X microscope objective	Olympus	30x 1.05 N.A. UPlanSApo
60X microscope objective	Olympus	60x 1.42 N.A. PlanAPO N
silicone immersion oil	Olympus	Z-81114
Neurolucida software	MBF Biosciences	Version 10
ImageJ software	U. S. National Institutes of Health	Current version	With the OME Bio-Formats plugin installed
Photoshop software	Adobe	version CS6