Assessment of Dendritic Arborization in the Dentate Gyrus of the Hippocampal Region in Mice

NOTE: Experiments were conducted in accordance with the ethical standards approved by the Committee on Animal Research at the Veterans Affairs Palo Alto Health Care System.

1. Golgi-Cox Staining

1. Brain extraction and staining
   1. On the 1st day, deeply anesthetize mice with 100 mg/kg ketamine and 10 mg/kg xylazine before euthanizing via exsanguination.
   2. Carefully remove the calvarium and dissect out the brain.
      1. First remove the skin on the top of the skull, placing a curved scissor on the top of the cerebellum and gently cut through the calvarium parallel to the central sulcus with the tip of the scissor pointed outward, moving in the direction toward the olfactory bulb. Repeat this process on both sides.
      2. Use a surgical pincet to lift the calvarium while turning it toward the olfactory bulbs and break it off. Then, flip the skull upside down and use a pick to cut through the optic nerves and loosen the brain. Finally, use the pick to separate the brain from the spinal cord at the level of the foramen magnum.
   3. Immediately immerse the brain in 15 ml of commercially-available, undiluted Golgi-Cox solution. Store the brain in Golgi solution in a capped, 20 ml glass bottle at room temperature in the dark.
   4. On the 2nd day, slowly pour out the solution and replace it with 15 ml fresh Golgi-Cox solution. Recap the bottle containing the brains and leave undisturbed for another 9 days in the dark.
2. Sectioning and embedment
   1. On the 11th day, place the brains in 30% sucrose solution, prepared in ddH₂O for dehydration. Change the solution with freshly prepared 30% sucrose after 12 hours. Incubate the brains in a 30% sucrose solution for 3 days at 4 °C.
   2. On the 14th day, fill the reservoir of the vibratome with a 30% sucrose solution until the blade is covered. Set the speed to 1 mm/s with amplitude of 0.95 mm.
   3. Blot the brains with a Kimwipe to remove excess moisture and remove the cerebellum using a razor blade to cut coronally.
   4. Firmly mount the whole brain onto the vibratome platform using superglue and allow it to set for 2-4 min. Insert the platform with the adhered brain into the reservoir and adjust the blade to the top of the brain.
   5. Using the vibratome, slice the brain into 150 µm thick sections at room temperature, remembering to change the blade after every 3rd brain.
   6. Pick up the sections from the reservoir using a small tipped brush and immerse them into  a petri dish containing 0.3% gelatin made in ddH₂O. While the sections remain in the petri dish, add 0.5 ml of 0.3% gelatin on each slide and use a brush to spread and coat the superfrost + slides.
   7. Gently pick up the immersed sections from the petri dish and place them onto the gelatinized slides. Orient the brain sections by touching them only on their sides and not on the tops.
   8. After about ten min, coat the sections with 30% sucrose before the tissue completely dries. Then air dry the prepared slides in a dark place in a slide rack for 3 days.
   9. Dilute commercially-available 10x Developing solution to 1x using ddH₂O. Immerse the slides in developing solution for 7-10 min. Rinse the slides 3 times in ddH₂O.
3. Dehydration
   1. Prepare 20%, 30%, 40%, 50%, 60%, 70%, 80%, 90%, and 95% ethanol solutions using distilled ddH₂O.
   2. Soak the slides in increasingly higher ethanol solutions for 10 min each.
   3. Soak the slides in 100% ethanol solution 3 times for 10 min each time.
   4. Soak the slides in 100% xylene 3 consecutive times for 5 min each, keeping the sections in the final solution until they are cover-slipped.
   5. Place a generous amount of DPX (Di-n-butyl phthalate in xylene) mounting medium (about 1-1.5 ml) on each slide using a plastic transfer pipet. Carefully place coverslips to avoid bubbles.
   6. Air dry cover slipped-brain sections horizontally for 3 days.

2. Doublecortin Staining

1. Deeply anesthetize (see step 1.1.1) the animals.
2. Perform cardiac perfusion with freshly prepared 4% paraformaldehyde made in phosphate buffer¹².
3. Extract the brains (see 1.1.2) and store in 45 ml of 4% paraformaldehyde at 4 °C overnight on a rocking shaker.
4. Transfer the brains to a 30% sucrose solution and wait for complete dehydration for 48 hr at 4 °C. Remove the brains from sucrose and place them directly on copper blocks placed on dry ice.
5. Fill the block with optimum cutting temperature (OCT) compound and mark the orientation of the brain using the olfactory bulb as a landmark.
6. Using a cryostat, cut 70 micron-thick sections at -20 ºC and place them in cryoprotectant solution (25% ethylene glycol, 25% glycerol, in 0.05 M sodium phosphate buffer, pH 7.4) and keep at -20 °C until use.
7. Make a solution of 50% cryoprotectant and 50% methanol. To this solution add 0.5% hydrogen peroxide (vol/vol). Incubate floating sections for 30 min at room temperature.
8. Warm sections in 37 ºC in TBS (Tris-buffered saline) for 30-40 min by placing them in a histological oven.
9. Incubate the sections with 0.3% triton and 3% normal horse serum for 45 min at room temperature prior to immunostaining. Incubate the sections at 4 ºC overnight in 3 ml solution of the DCX antibody diluted 1:500 in TBS with 0.3% triton and 1% normal horse serum.
10. The following day wash the sections 3 consecutive times in TBS (10 min each).
11. Incubate sections with a biotinylated horse anti-goat (1:200 in TBS) for two hr at room temperature. Wash the sections 3 consecutive times in TBS (10 min each).
12. Incubate them with ABC Lite (1:1,000) for 1.5 hr at room temperature.
13. Add 5 µl of 30% H₂O₂ to one tablet of 10 mg of DAB (3,3”-Diaminobenzidine) solution dissolved in 15 ml of 1 M Tris-HCl (pH 7.6). Incubate the sections immediately in this solution for 5 min.
14. Terminate the reaction by washing them with ice-cold TBS three times followed by one wash in TBS at room temperature.
15. Dehydrate sections using a series of ethanol solutions (50%, 60%, 70%, 80%, 90%, 95%, 100%, 5 min each), clear in xylene, and then cover slip using DPX as in step 1.3.

3. Visualization

1. Following complete drying of the sections, remove excess DPX on top of the slides with a sharp blade, and place them on the scanning stage of the microscope. The system is composed of a microscope equipped with scanning stage, joystick, and a color 12-bit camera.
2. Start the program. Open a new data file.
3. Place a reference point on the screen by clicking with the mouse pointer at any place. This will activate all the icons from the tool panel of program window.
4. Click on the “Joystick Free” icon on the toolbar and then use the joystick to locate the dentate gyrus of the hippocampus in the first section.
5. In the “Tools” tab of the program window, select “Serial Section Manager”. Click on the “New Section” icon on the lower left corner of the window. This will open the “Serial Section Setup” window.
6. Select the Serial Section Setup window and then enter the total number of sections that contain hippocampal regions. Select the evaluation interval. Enter the section cut thickness (70 µm for DCX and 150 µm for Golgi-stained sections).
7. Start tracing the dentate gyrus region by clicking on the “Freehand Contour Drawing” icon in the tool bar. Outline the dentate granular cell layer.
8. Select “100X” from the "Magnification" menu. Add a drop of immersion oil on the section and switch the objective to 100X. Locate the dentate granule cell bodies and dendrites under this objective.
9. Focus on a selected neuron and click on the “Neuron Tracing” icon on the toolbar. Trace the circumference of the dentate granule cell body. When finished tracing, right click to select “Finish Cell Body”.
10. Start tracing the dendrites manually in x, y, and z directions and follow each branch using the joystick and z-motor knob. At a bifurcation or trifurcation node, select the respective option from the dropdown menu upon right-click. Trace each of the branches that arise from these nodes. At the end of each branch right click and select “Ending” from the dropdown menu.
11. Using the arrow key icons in the software window randomly select another dentate granule cell and follow the same procedure as described in step 3.9-3.10. Save the entire tracing.
12. Start the neuronal tracing software.
13. Open the first NRX data file from the first mouse of the experimental group. Append the NRX file of the second mouse of the experimental group. Continue until all the NRX files from this group are appended.
14. Under the Analysis tab, select “Fan In-diagram”. This opens the Fan In-analysis window. Check mark “dendrites” and click Display, which depicts the lengths and branching patterns of dendrites for this group of mice (Figure 1).

4. EDFI

NOTE: This method enables the user to rapidly record a large number of images through the z-axis of each field and generate a 2D image that contains all the focused pixels throughout the z-axis. The result will be a composite image that contains all focused pixels from collected images.

1. Connect the microscope to a digital camera. Place the sections first on the scanning stage connected to the microscope and then switch to a 10X objective. Move the stage to the area of interest.
2. Start a video capturing program.
3. Press the record button. As soon as the record button is pressed, use the macro-focus knob to move from the top to the bottom of the section for a total of 4 sec. Save the resulting video file.
4. Use ImageJ freeware to convert the AVI video files into an uncompressed format by opening the file and saving them in uncompressed file format.
5. Start an image analysis program and open the AVI file. Go to “Process” menu and click on “Extended Depth of Field”. Select the corresponding video file. In “Output” options, select “Generate Composite Best-Focus Image”.
6. In “Focus Analysis” options, select “Normalized Illumination” and “Max Local Contrast” options. Then, select “Create”. The resulting image represents all the focused pixels throughout the z-axis (Figure 2).
7. Save the resulting image.

5. Generating Extremely-high Resolution Images

NOTE: This method allows the user to automatically generate low magnification and extremely high-resolution images from high-magnification-high resolution images in an automatized fashion.

1. Place the sections first on the scanning stage connected to the microscope. The microscope is connected to a digital camera. Move the stage to the area of interest.
2. Start an image acquisition program.
3. Use a 10X objective and acquire and save high-resolution (4076 x 3116) images from the region of interest making sure to have at least 10% overlap.
4. Run Image Composite Editor to stitch images.
5. Go to File menu, and click on “New Panorama”. Select the folder in which the images are stored. Select the images that belong to the same region and press “ok”.
6. Press “Export to Disk” button and save the image. This image represents an extremely high-resolution picture of the area of interest that could be used for analysis and/or demonstration (Figure 3).