Isolation and Expansion of Neurospheres from Postnatal (P1−3) Mouse Neurogenic Niches

All experiments were performed in accordance with the European Community (86/609/EEC; 2010/63/EU; 2012/707/EU) and Portuguese (DL 113/2013) legislation for the protection of animals used for scientific purposes. The protocol was approved by the "iMM's institutional Animal Welfare Body – ORBEA-iMM and the National competent authority – DGAV (Direcção Geral de Alimentação e Veterinária)."

1. Basic setup and preparation of culture medium

1. On the day of dissection, prepare the appropriate amount of growth medium corresponding to serum-free medium (SFM) composed of Dulbecco's modified eagle medium [(DMEM)/F12 with L-glutamine] (Table of Materials) supplemented with 100 U/mL penicillin and 100 µg/mL streptomycin (pen/strep), 1% B27, with also 10 ng/mL EGF and 5 ng/mL bFGF. Warm the culture medium to 37 °C in a water bath.
   NOTE: The volume of growth medium depends on the number of pups, for 5 pups prepare ~100 mL (50 mL for SVZ and 50 mL for DG); however, after counting the number of cells (step 5.1) the exact volume will have to be adjusted.
2. For microdissection of SVZ and DG, prepare the calcium and magnesium-free Hanks' balanced saline solution (HBSS) dissection medium supplemented with 100 U/mL pen/strep.
   NOTE: Prepare 50−100 mL of dissection medium.
3. Set up a dissection microscope and prepare the tools needed to remove the brain (scissors and small spatula) and for SVZ and DG microdissections (Dumont small scissors, #7 forceps, #5 forceps, #5S forceps) by soaking in 70% ethanol.

2. Harvesting of postnatal (P1−3) mouse brains and SVZ/DG microdissections

1. Prepare 60 mm Petri dishes (growth area 21 cm²) with HBSS supplemented with pen/strep and 2 sample tubes (one for SVZ and one for DG) with 500 µL of supplemented HBSS each.
2. Euthanize mice pups (P1−3) according to the protocol approved by the Institutional Animal Care facility/guidelines. Perform decapitation with a single incision with sharp scissors at the base of the brainstem.
3. Holding the ventral part of the body at the base of the head and using small pointed scissors, make a midline incision in the skin over the entire length of the head, thus revealing the surface of the skull.
4. Make a longitudinal incision at the base of the skull and continue cutting along the sagittal suture using small scissors with an angle shallow as possible in order to avoid damaging the brain structures.
5. Peel the skull to the sides using curved forceps and expose the brain.
   CAUTION: Make sure the dissecting instruments are free of ethanol before touching the brain.
6. Isolate the brain from the skull using a small spatula, by sliding under the base of the brain to cut the cranial nerves and blood vessels that are connected to the base of the brain, and transfer the brain into a Petri dish containing cold supplemented HBSS solution.
7. Place the Petri dish containing the brain under a dissecting microscope at low magnification and position the brain on its dorsal surface.
8. Using fine forceps, remove the meninges from the ventral side of the brain and the olfactory bulbs, while holding the brain in position by the cerebellum. Rotate the brain onto the ventral aspect and peel off the rest of the meninges.
   NOTE: Removing the dorsal meninges is a crucial step to ensure correct brain slicing.
9. Discard the cerebellum making a cut using forceps. Place a filter paper with a pore size of 11 µm onto a tissue chopper (Table of Materials) and set the brain onto the filter paper using curved-pointed forceps. Chop the brain into 450 µm coronal sections and use a wet lamina to collect the sectioned brain into a new Petri dish filled with cold supplemented HBSS.
10. To dissect out the SVZ, use forceps to separate coronal slices in an anterior-to-posterior fashion until reaching slices with the lateral ventricles, under a dissecting microscope.
11. Cut the thin layer of tissue surrounding the lateral wall of the ventricles (which corresponds to the SVZ) with fine forceps, excluding the striatal parenchyma and the corpus callosum. Isolate the SVZ by placing the tip of the forceps in the lateral corners of the lateral ventricle: one immediately under the corpus callosum and the other into the tissue immediately adjacent to the ventral area of the lateral ventricle. Then, cut a small line of tissue surrounding the lateral ventricle.
12. Collect the dissected tissue into a sample tube with supplemented HBSS solution previously identified as SVZ.
    NOTE: Exclude the SVZ in slices where both the lateral ventricles and the hippocampal formation begin to appear.
13. Go through all slices after SVZ microdissection in an anterior-to-posterior fashion and reach the hippocampal formation. Using forceps discard the first slice with hippocampus where DG is still unrecognizable.
14. To remove the DG, first isolate the hippocampus from the slices. Refocus the microscope to visualize the borders around DG.
15. Dissect the DG portion by performing a cut between the DG and CA1 region followed by a vertical cut between the DG and CA3 region using forceps. Remove fimbria and any adjacent tissue.
    NOTE: In P1−3 animals, the DG is almost undistinguishable from Ammon's horn but displays a small tip.
16. Collect the dissected tissue into a sample tube containing supplemented HBSS solutions previously identified as DG.
    NOTE: Overall injury to the hippocampus or surrounding area will make it more difficult to isolate the DG. Using an atlas of the postnatal mouse brain is essential when the user is not familiarized with the isolation of the SVZ and DG tissue from coronal sections.

3. Tissue dissociation

1. To dissociate the SVZ and DG tissue present in their respective tubes, add trypsin-EDTA 0.05% to have a final concentration of 5−10% of Trypsin-EDTA 0.05% in HBSS. Incubate for approximately 15 min at 37 °C, until the tissue is clumped together.
2. Wash the tissue from the trypsin by removing the media and adding 1 mL of new HBSS supplemented solution for 4 consecutive times.
3. Remove the HBSS and resuspend the digested tissue in 1 mL of SFM supplemented with 10 ng/mL EGF and 5 ng/mL bFGF. Mechanically dissociate the pellet by gently pipetting up and down approximately 7−10x using a P1000 pipette, until getting a homogenous cell solution.
   CAUTION: Excessive mechanical dissociation can lead to increased cell death and will negatively impact subsequent cell growth.

4. Cell-pair assay to study cell fate

1. Prior to the experiment, prepare coated 24-well plates for adherent monolayer cultures according to sections 8-10.
2. To count the number of SVZ or DG cells (obtained in section 3) to be plated, use a solution containing 0.2% Trypan blue and count the cells using a hematocytometer.
3. Dilute the dissociated cell suspension in SFM supplemented with 5 ng/mL EGF and 2.5 ng/mL bFGF (low EGF/bFGF) at a density of 11,300 cells/cm² and plate them on coated glass coverslips.
4. After 24 h, fix the cells for immunocytochemistry against NSC markers such as sex determining region Y-box 2 (Sox2) and nestin as well as with a marker of the neuronal lineage (namely doublecortin [DCX], for immature neurons) (see section 14).
   NOTE: Sox2 is a marker of NSCs that undergo mitosis. Sox2^+/+ cell-pairs resulting from a single progenitor cell division reflects stem cell expansion⁷.

5. Expansion of postnatal neural stem cells as neurospheres

1. To determine the density of the dissociated SVZ or DG cell suspension (obtained in section 3), count the cells using a hematocytometer.
2. Dilute SVZ and DG single cell suspension at a density of 2 x 10⁴ cells/mL in SFM supplemented with 10 ng/mL EGF and 5 ng/mL bFGF. Seed SVZ and DG cells in uncoated 60 mm Petri dishes with a final volume of 5 mL/Petri dish.
3. Incubate SVZ and DG cells for 6−8 days and 10−12 days, respectively to form primary neurospheres, at 37 °C with 5% CO₂.
   NOTE: Incubation days more than those mentioned can promote aggregation of neurospheres and higher levels of cell death in the center of the neurosphere.
4. When the majority of neurospheres have a diameter of 150−200 µm, perform the neurosphere passage.
   NOTE: Passaging neurospheres when they do not have an appropriate diameter compromises all the next steps.

6. Passaging of neurospheres

NOTE: The following protocol can be applied to expand both SVZ and DG neurospheres.

1. To passage neurospheres, collect the SFM with growth factors containing neurospheres from the 60 mm Petri dish(es) and centrifuge for 5 min at 300 x g.
2. Discard the supernatant and resuspend the neurosphere pellet using a chemical dissociation kit (mouse) according to the manufacturer's instructions (Table of Materials).
   NOTE: Observe the incubation times precisely as they are crucial for performance.
3. Centrifuge for 5 min at 300 x g, remove the supernatant and add 1 mL of SFM supplemented with 10 ng/mL EGF and 5 ng/mL bFGF.
4. Triturate up and down approximately 10x with a P1000 pipette to dissociate neurospheres.
5. Count the number of cells using a solution containing 0.2% Trypan blue and a hematocytometer.
6. Reseed cells at a density of 2 x 10⁴ cells/mL in uncoated 60 mm Petri dishes.
7. Incubate SVZ and DG cells for 6−8 days and 10−12 days, respectively to obtain secondary neurospheres, at 37 °C with 5% CO₂.
   NOTE: The self-renewal capacity of SVZ- and DG-derived NSPCs can be accessed by following protocol sections 5 and 6. For that, seed SVZ and DG cells at a density of 1.0 x 10⁴ cells/mL (in uncoated 24-well plates) in growth SFM medium containing 5 ng/mL EGF and 2.5 ng/mL bFGF (low EGF/bFGF). Count the number of resulting primary and secondary neurospheres.

7. Storage of neurospheres

1. Collect the medium containing neurospheres (obtained from steps 5.3 and 6.7) from the 60 mm Petri dishes.
2. Centrifuge for 5 min at 300 x g and discard the supernatant.
3. Wash the cells 2x with 1 mL of HBSS (5 min at 300 x g).
4. Centrifuge for 5 min at 300 x g, discard the supernatant and store the pellet of neurospheres at -20 °C for molecular biology analysis.

8. PDL coating plate procedure

1. To prepare solution 1 (0.1 M borate buffer), weigh 3.92 g of boric acid and dilute in 400 mL of high purity water. Adjust the pH to 8.2 and make up to 500 mL with high purity water.
2. To prepare solution 2 (0.167 M borate buffer), weigh 10.3 g of boric acid and dilute in 900 mL of high purity water. Adjust the pH to 8.2 and make up to 1,000 mL with high purity water.
3. To reconstitute poli-D-lysine (PDL) (1 mg/mL in 0.1 M borate buffer), dilute 100 mg of PDL in 100 mL of solution 1.
4. Make aliquots of 10 mL to use immediately or freeze and store at -20 °C.
5. Under the laminar flow, add 1 coverslip per well and sterilize under UV light for 15 min.
6. Use the reconstituted PDL or thaw frozen reconstituted PDL.
7. Prepare the final solution of 100 µg/mL PDL in 0.167 M borate buffer by adding 10 mL of reconstituted PDL to 90 mL of solution 2.
8. Add the final solution to wells for a minimum of 2 h to overnight at 37 °C.
   NOTE: For 24-well plates, add a volume of 500 µL in each well.
9. Remove the solution and wash 3x with high purity water.
10. Let the coverslips dry in the laminar flow hood.
11. Leave the multi-well culture plates at 4 °C.

9. PDL/Laminin coating plate procedure

1. On day 1, coat the plates with PDL as described in section 8.
2. On day 2, remove the PDL solution and wash 3x with high purity water. Let dry.
3. Prepare 5 µg/mL laminin in cold SFM devoid of growth factors.
4. Add dissolved laminin to the coverslips and incubate at 37 °C overnight.
   NOTE: For 24-well plates, add a volume of 500 µL in each well.
5. Remove laminin using a pipette.
   NOTE: Do not wash the coverslips from laminin.
6. Use immediately or store at -20 °C.

10. Poly-L-ornithine (PLO) /laminin coating procedure

1. Under the laminar flow, add one coverslip per well and sterilize under UV light for 15 min.
2. Add 0.01% PLO solution to each well for 20 min at room temperature (RT).
   NOTE: For 24-well plates, add a volume of 500 µL in each well.
3. Remove the solution and wash 3x with sterilized 1x PBS. Let dry.
4. Prepare 5 µg/mL laminin in sterile 1x PBS.
5. Incubate for 2 h at 37 °C.
6. Remove laminin.
   NOTE: Do not wash the coverslips from laminin.
7. Use immediately.
   NOTE: Make sure that the coverslip is fully covered by the PLO solution by gently tapping the coverslip with a pipette tip. When shaken, the multi-well plates should make no sound.

11. Evaluation of neuritogenesis by generating a differentiated monolayer of cell

1. Collect media containing neurospheres from 60 mm Petri dishes (obtained from section 5) and centrifuge for 5 min at 300 x g at RT.
2. Discard the supernatant and dissociate the pellet of neurospheres in 1 mL of dissociation PBS (i.e., PBS without Mg²⁺/Ca²⁺ and with EDTA [2.7 mM KCl, 1.5 mM KH₂PO₄, 137 mM NaCl, 8.1 mM Na₂HPO₄ and 0.5 mM EDTA 4Na, at pH = 7.40]) by incubating for 15 min followed by mechanical dissociation. Alternatively, dissociate neurospheres using a chemical dissociation kit (mouse) (Table of Materials).
3. Centrifuge for 5 min at 300 x g at RT and discard the supernatant.
4. Resuspend the cell pellet in 1 mL of SFM devoid of growth factors.
5. Determine cell density using a hematocytometer.
6. Dilute the dissociated cell suspension in SFM devoid of growth factors at a density of 3,766 cells/cm² and plate cells on coated glass coverslips in 24-well plates.
7. After 1−3 days, fix cells for immunocytochemistry against a protein of the cytoskeleton (see section 14).

12. Differentiation of neurosphere cultures

NOTE: Neurospheres obtained from cell expansion, either from primary or passaged neurospheres (obtained in sections 5 or 6) can be differentiated into cells from different neural lineages.

1. When neurospheres have a diameter of 150−200 µm, collect 25 µL of neurosphere suspension medium and plate on coated glass coverslips, in 24-well plates.
   NOTE: To collect more neurospheres, gently rotate the Petri dish to concentrate the neurospheres in the center. Then, pipette from the center.
2. Put the plates in an incubator at 37 °C for 15 min so that neurospheres adhere to the substrate. Afterwards, add 500 µL of SFM devoid of growth factors (differentiative conditions).
3. After 24 h, replace the medium with fresh SFM devoid of growth factors.
4. Differentiate for different time points (e.g., 2 and 7 days in vitro, DIV2 and DIV7, respectively) with 5% CO₂ and 95% atmospheric air at 37 °C.
   NOTE: Cell survival, proliferation and differentiation can be analysed using different cell assays.

13. Cell biology assays

1. Cell survival assay
   1. Expose plated neurospheres to 3 µg/mL propidium iodide (PI) for 30 min before cell fixation in the incubator at 37 °C.
      NOTE: PI is an autofluorescent agent that is only able to enter cells with compromised membrane integrity⁸. Other methods to analyse cell survival can be used such as caspase 3 staining or the terminal deoxynucleotidyl transferase dUTP nick-end labeling (TUNEL) assay.
2. Cell proliferation assay
   1. Expose plated neurospheres to 10 µM 5-bromo-2'-deoxyuridine (BrdU) for 4 h before fixation in the incubator at 37 °C.
      NOTE: BrdU is a synthetic thymidine analogue that can be incorporated during DNA synthesis in proliferating cells⁹.
3. Cell differentiation assay
   1. Expose 7-day-old plated neurospheres to 10 µM BrdU in the first 24 h, in the incubator at 37 °C.
   2. Renew the SFM devoid of growth factors (differentiative conditions) and allow cells to develop in the absence of BrdU for the following 6 days until fixation.
      NOTE: These pulse-chase experiments, by co-labelling with markers of mature neural cells, allow the evaluation of progenitor cells that differentiate into mature cells during the protocol.

14. Immunostaining of neurosphere cultures

1. Cell fixation
   1. Prepare 4% paraformaldehyde (PFA) in 1x PBS and store at 4 °C or -20 °C.
   2. Remove the SFM devoid of growth factors from wells and add, to each well, 500 µL of 4% PFA at 4 °C for 20 min at RT.
   3. Wash 3x with 1x PBS, for 5 min each time, the coverslips containing differentiated neurospheres.
   4. Store coverslips until use in 500 µL of 1x PBS at 4 °C.
      NOTE: If the experiment does not have BrdU, skip to step 14.3.
2. Denaturation method (for BrdU experiments only)
   1. Prepare 1 M HCl at 37 °C.
   2. Rinse coverslips 3x in 1x PBS.
   3. Permeabilize cells for 30 min in PBS containing 1% nonionic surfactant (e.g., Triton X-100).
   4. Denature dsDNA with 1 M HCl pre-heated to 37 °C for 30−40 min at 37 °C (~300 µL/well).
   5. Wash wells 4x with 1x PBS.
3. Permeabilization and blocking
   1. Rinse coverslips in 1x PBS for 5 min.
   2. Incubate for 1.5 h with 0.5% nonionic surfactant and 3% bovine serum albumin (BSA) in 1x PBS (~300 µL/well).
      NOTE: For NeuN, use 6% BSA in 1x PBS.
4. Incubation and mounting
   1. On day 1, without washing, incubate cells with primary antibodies (Table of Materials) in 0.1% nonionic surfactant and 0.3% BSA in 1x PBS in the incubation chamber (for 24-well plates use 20 µL/well). Leave coverslips incubating overnight at 4 °C light protected if antibodies are conjugated to a fluorophore.
   2. On day 2, return coverslips to their respective wells and rinse 3x in 1x PBS for 5 min.
   3. Counterstain with appropriate fluorescence conjugated secondary antibodies (dilution 1:200) and with 12 µg/mL Hoechst 33342 in 1x PBS for 2 h at RT and light protected in the incubation chamber (20 µL/coverslip).
   4. Wash coverslips 3x in 1x PBS for 5 min.
   5. Mount coverslips onto microscope slides using 5 µL/coverslip of fluorescence mounting medium.
   6. Let coverslips air dry at RT, protected from light, for 1 day.
5. Microscopy
   1. View and acquire images using a fluorescence microscope.
   2. For each condition, use three replicates. Perform cell counts in five independent microscopic fields in each coverslip with a 40x objective (~100 cells per field).

15. Preparation of EGF and bFGF stock solutions

1. EGF stock solution
   1. To reconstitute lyophilized EGF, dilute the product in high purity water to reach a final concentration of 20 µg/mL.
   2. Aliquot and store in microtubes at -5 to -20 °C.
2. bFGF stock solution
   NOTE: bFGF must be reconstituted with a solution of 10 mM Tris, pH 7.6.
   1. Centrifuge the vial briefly before opening to bring the content to the bottom.
   2. Prepare 50 mL of 10 mM Tris, pH 7.6. For that, weigh 60.57 mg of Tris ((HOCH₂)₃CNH₂) and dilute it in 40 mL of high purity water. Adjust the pH to 7.6 and make up to 50 mL with high purity water.
   3. Prepare 10 mL of 0.1% BSA in 10 mM Tris, pH 7.6. For that, weigh 10 mg of BSA and dilute it in 10 mL of 10 mM Tris.
   4. Filter solutions prepared in steps 15.2.2 and 15.2.3 with a 0.22 µm filter under a laminar flow hood.
   5. Reconstitute 10 µg of bFGF in 1,000 µL of 0.1% BSA in 10 mM Tris with pH 7.6 to reach a final concentration of 10 µg/mL. Aliquot into microtubes at -20 °C for a maximum of 6 months.