Microscopic Imaging of Neurons in a Mouse Brain Section Using Golgi-Cox Metallic Staining

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All procedures involving animal samples have been reviewed and approved by the appropriate animal ethical review 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 separately in high-quality water. 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 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 precooled on dry ice.
   7. Place the frozen brain into a 50 mL conical tube and store it 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 and keeping it 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, then let it 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 it.
   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. 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, using the mesh-bottom inserts. Incubate on a rocker, moving slowly in the dark at RT for 15 min.
   2. Sections should be washed twice by transferring them to new wells containing 5 mL of water. They should be washed 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 slowly in the dark at RT for 15 min.
   4. Sections should be washed twice by transferring them to new wells containing 5 mL of water. They should be washed 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 diluted in water 10x from its original purchased concentration. Incubate on a rocker, moving slowly in the dark at RT for 25 min.
   6. Sections should be washed twice by transferring them to new wells containing 5 mL of water. They should be washed on a rocker moving at a moderate speed in the dark at RT for 5 min.
4. Mounting Brain Sections
   1. Using a small paintbrush, mount sections onto microscope slides. Using tweezers and a small tissue, remove excess water and agar. 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 are critical and may need to be determined in each laboratory depending on ambient temperature and humidity. 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 have never been frozen 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 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 that for other species, including rats and cowbirds, the final clearing step may need to be extended up to 15 minutes in 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 on the 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 on 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, ensuring 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 contact between the objective and the oil.
      2. Focus on 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, ensuring that all image stacks overlap by at least 10% in the X and Y axes.
   7. Save the data file and 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 the two-dimensional Z-projection image as a TIFF file.
   2. Create a two-dimensional image montage of the entire area of interest.
      1. Open the software (e.g., Adobe Photoshop).
      2. Select "File" -> "Automate" -> "Photomerge".
      3. Select "Browse" and then add all image files to be merged.
      4. Ensure that "Blend Images" is selected and then select "OK".
      5. Save the resulting montage image of the entire area of interest as a TIFF file.

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