A Method for Exploring Adult Neurogenesis in the Songbird Brain

BrdU Injections

• The number of injections per bird and the survival times depend on the question addressed. To analyze the number of mitotically active cells in the ventricular zone, perform a single injection, followed by a short survival time (2–4 h). For quantification of the number of new cells incorporated into a particular brain region, especially in adults where relatively few cells are labeled per day, perform multiple injections over several days (i.e., inject each bird twice daily, at the beginning and end of the light cycle, for 4 d) and use a survival time of roughly 30 d to allow for labeled cells to migrate to their final destination and differentiate (Alvarez-Buylla and Nottebohm 1988; Kirn et al. 1999). For birds larger than the zebra finch, osmotic minipumps can be implanted, which can deliver cell birth markers for a week or longer (see Thompson and Brenowitz 2005), ensuring that the marker is available for uptake by dividing cells that vary in the timing of S phase.

• 1. Measure the weight of the bird by weighing a small cardboard box with a lid and then subtracting its weight from the weight of the bird inside the same box.

• 2. In one hand, gently restrain the bird on its back, using the thumb and fingers to move its wings and feet out of the way to maximize exposure of the breast muscle (musculus pectoralis) to be injected. If desired, use a small amount of 70% ethanol to moisten feathers and spread them for a better view of the muscle. Inject the bird intramuscularly with 6.4 µL of freshly prepared BrdU solution per gram of body weight using a glass microsyringe and 28-gauge needle.

  • This dosage equals roughly 100 mg BrdU/kg body weight. Adult zebra finches weigh 12–15 g, so 1 mL of BrdU solution is sufficient to inject roughly 10 birds.

  • If multiple injections are performed, alternate sides to minimize bruising.

• 3. Allow the bird to survive for an appropriate period postinjection.

Perfusion

• 4. Euthanize each bird with a lethal intramuscular injection of 0.175 mL each of ketamine (10 mg/mL) and xylazine (20 mg/mL). Confirm complete anesthesia by ensuring that the birds do not respond to a toe pinch.

• 5. Place the birds on their backs in a perfusion tray in a fume hood with their wings and legs taped down. Use 70% ethanol to moisten the feathers over the breast muscle. Under a dissecting microscope, expose the intraperitoneal cavity by making an incision with fine iris scissors just below the ribcage, while pulling up on the ribcage to avoid cutting internal organs. Enlarge the incision following the edge of the ribcage on either side toward the neck, creating a “flap” that, when lifted, exposes the heart. Rupture the bird's right atrium and perform perfusion using a 28-gauge needle through the bird's left ventricle. Perfuse first with 20 mL of 0.1 m PBS (pH 7.4) followed by 50 mL of 4% paraformaldehyde. Use both solutions at room temperature.

• 6. Decapitate birds and expose the skull by sliding the skin from the neck region toward the beak. Beginning at the back of the skull near the cerebellum, use fine forceps or a scraping tool to chip away the two layers of skull surrounding the sides and top of the brain. Cut and remove the meninges using two pairs of fine forceps, one pair to lift the meninges off the brain and the other to simultaneously pull on them until they tear. Work along the midline between hemispheres to pull the meninges laterally, fully exposing the brain, and remove them.

• 7. Gently lift the brain away from the base of the skull. Sever the cranial nerves, and carefully remove the brain.

• 8. Cut the brain into two hemispheres with a single-edged razor blade.

  • We routinely use sagittal sections, but any plane of sectioning can be used.

• 9. Place the brain in fresh 4% paraformaldehyde for 1 h at room temperature.

• 10. Remove the brain from the fixative and incubate for 1 h in 0.1 m PBS with two to three changes of PBS on a shaker at room temperature.

Dehydration

• 11. Dehydrate the brains separately in plastic vials with increasing ethanol concentrations at room temperature on a shaker. Ethanol concentrations are as follows: 50% for 1 h, 70% for 1 h, 95% for 40 min, and 100% for 35 min.

  • While performing this step, put plastic ICC trays in the incubator to warm. One 4× 6-well ICC tray can be used for processing three brains (six hemispheres).

Infiltration/Embedding

• While not essential, a vacuum oven can be used to shorten infiltration times.

• 12. Use a warmed plastic pipette to fill wells of 4 × 6-well ICC trays with the following PEG solutions. Incubate the dehydrated brain tissue at 47°C in the solutions for the times indicated. Use a suitable glass or metal weight to keep the brain tissue submerged and prevent it from floating. Use warmed forceps to transfer the tissues to the new wells during solution changes.

  • i. PEG 1000 for 1 h

  • ii. PEG 1000/1450 (3:1 ratio) for 3 h

  • iii. PEG 1000/1450 (3:1 ratio) for 8–12 h (overnight)

  • iv. PEG 1450 for 20 min.

    • At the time of the final PEG change, place the plastic embedding molds on a small tray in the incubator to warm, in preparation for embedding.

• 13. Embed the tissue as follows.

  • i. Fill each plastic embedding mold at least halfway with PEG 1450. Transfer the brain tissue to the plastic embedding mold with the medial surface facing down. Cover the tissue with PEG 1450 using a warmed plastic pipette, making sure the medial surface of the hemisphere remains as close as possible to the bottom of the embedding mold. Add PEG until the molds are ∼¾ full.

  • ii. Carefully take the tray of molds out of the incubator and bring them to room temperature so that the PEG hardens. Place molds in a −20°C freezer for 20 min to accelerate hardening, if desired.

    • The humidity of the room will influence how well PEG solidifies. Optimum relative humidity ranges are between 20% and 30%.

  • iii. Store PEG molds at 4°C in a sealable plastic bag with a Drierite desiccant pouch.

• 14. Cut the tissue as follows.

  • i. Bring PEG mold to room temperature and place it in the incubator for 30 min at 47°C.

    • This will slightly soften the top of the PEG block for a better seal with the microtome chuck.

  • ii. Use a warm plastic pipette to add a layer of fresh, molten PEG 1450 on top of the block. Place a chuck on top of the PEG mold and fill up the rest of the mold with fresh PEG 1450. Bring to room temperature.

    • This will create a block of PEG with brain tissue embedded inside, attached to a chuck for mounting in a rotating microtome.

  • iii. Section the tissue.

    • PEG sections can be cut at a range of thicknesses; however, we have had the best success with 6-µm sections using a rotary microtome.

    • Sections come off the microtome in ribbons, which can be stored in a tray made of cardboard with numbered lanes (Fig. 1). The lanes are made with foam weather stripping, which helps preserve the medial-lateral order of ribbons and provides space so that a cardboard cover can be affixed with tape without letting it touch the ribbons. The sections can be mounted onto glass slides immediately or covered trays of sections can be stored at 4°C in a sealed bag with a Drierite desiccant pouch.

• 15. Mount the tissue as follows.

  • i. If the sections are stored in a sealed bag, bring them to room temperature before unsealing the bag. Break apart ribbons of sections using a fine-tipped, slightly damp paintbrush, if necessary.

    • Wetting the brush too much may lead to dissolving the PEG around a section, which would make the section unrecoverable.

  • ii. Float sections on 0.1 m PBS for mounting. Using the paintbrush to orient the section so that the section is above and parallel with the surface of the PBS, gently lower the section onto the PBS with the paintbrush on the underside.

    • Sections should detach from the paintbrush and remain flat on the surface of the PBS.

    • It is essential that a very fine-tipped brush be used.

  • iii. Orient floating sections so that they contact the edge of a glass slide set at an angle in the PBS. Raise the slide to lift the section out of the PBS.

    • Use positively charged slides to ensure that the sections do not fall off during subsequent processing.

  • iv. Air-dry the sections before proceeding with staining. For immunohistochemistry, put the mounted sections in slide boxes and store in sealed bags with desiccant at −20°C.

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Figure 1.

PEG Ribbons on trays. An example of the cardboard storage trays for cut PEG sections. Ribbons of PEG sections can be cut and placed in lanes created using foam weather stripping. Creating lanes helps keep ribbons of sections in their proper sequence during mounting and storage. In this particular image, intact ribbons can be seen at the far right, sections having been detached from the other ribbons and mounted onto slides. Before storage, a cardboard cover is taped over the tray and then the trays are placed in sealed plastic bags with desiccant at 4°C or −20°C.

Antigen Retrieval

• 16. Lay slides flat on a raised platform in an electric vegetable steamer. Cover sections with enough 0.01 m citrate buffer to cover the surface of the slide, to prevent drying of the tissue. Incubate for 10 min at 95°C–100°C.

• 17. Heat PBS in a glass staining dish and place it in the incubator at 37°C. Place slides into a metal slide rack, and immerse the slide rack in the staining dish. Incubate slides in the PBS for 5 min.

• 18. Incubate slides with pepsin solution for 3 min at 37°C in the incubator.

  • Pepsin incubation is necessary to expose the antigen in PEG sections. When omitted, the signal is reduced substantially.

Blocking

• 19. Place drops of 0.1 m PBS directly on each slide and transfer to humid chamber for 5 min at room temperature.

  • All subsequent incubations are performed with the slides laid flat in the humid chamber.

• 20. Incubate slides with blocking buffer for 30 min at room temperature.

Incubation with Antibodies and Detection

• 21. Dilute primary antibodies for BrdU (1:100) and Hu (1:40) in blocking buffer. Incubate slides with primary antibody mixture at room temperature overnight.

  • Place rectangular cutouts of Parafilm over slides during the incubation to ensure even distribution of antibody over sections and to prevent drying out.

• 22. Wash slides with PBST three times for 5 min each at room temperature.

• 23. Dilute secondary antibodies against sheep (1:200) and mouse (1:1000) with blocking buffer. Incubate slides with secondary antibody mixture for 2 h at room temperature.

• 24. Wash slides with PBST three times for 5 min each at room temperature.

• 25. Incubate slides with fluorophore-conjugated streptavidin (1:800 in PBST) for 2 h at room temperature.

• 26. Wash slides with PBST three times for 5 min each at room temperature.

• 27. Fill a staining dish with ddH₂O and dip the slides directly into the staining dish. Add a coverslip using 1 drop of Aqua/Poly Mount per brain section on the slide.

• 28. Examine slides under fluorescence microscope with appropriate filters (Fig. 2).

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Figure 2.

Fluorescence photomicrographs of adult-formed neurons in Area X, the hippocampus, and high vocal center (HVC) from an adult male zebra finch. Row 1: BrdU⁺ cell nuclei in Area X, the hippocampus, and HVC visualized using a FITC filter. Row 2: The same fields of view showing BrdU⁺/Hu⁺ neurons, revealed with a dual FITC-rhodamine filter to show colocalization of the two markers. Row 3: The same fields of view showing Hu⁺ neuronal label in the cytoplasm, visualized with a rhodamine filter. Scale bars, 10 µm. The fluorophore used for visualizing BrdU was Alexa 488 (Molecular Probes); the neuron-specific protein Hu was labeled with Cy-3 (Jackson ImmunoResearch Laboratories). From Pytte et al. 2007; reprinted with permission from Wiley Periodicals, Inc.