Isolation, Expansion, and Nucleofection of Neural Stem Cells from Adult Murine Subventricular Zone

All experiments performed with animals were previously approved by the University of Valencia's Ethical Committee and authorized by Conselleria de Agricultura, Ganadería y Pesca, Generalitat Valenciana (Spain).

1. Primary culture of neurospheres

1. Preparation of reagents
   1. Prepare a buffer solution of Dulbecco's Phosphate Saline Buffer (DPBS) in deionized water and sterilize by autoclaving. Alternatively, prepare 0.1 M PBS pH 7.4 by adding 137 mM NaCl, 2.7 mM KCl, 10 mM Na₂HPO_4, and 1.8 mM KH₂PO₄ solution in deionized water.
   2. Fill two 12-well plates with previously cooled DPBS and keep them on ice: one will be used to preserve the whole brain before dissection, and the other will preserve dissected SVZs until tissue processing.
   3. Prepare papain-containing enzymatic mix as described below.
      1. Prepare EDTA/L-cysteine dissociation solution in Earl's Balanced Salt Solution (EBSS) to a final concentration of 0.2 mg/mL each. To achieve complete dissolution, incubate the tube in a water bath at 37 °C.
      2. Dissolve 12 U/mL of papain in EDTA/L-cysteine solution. Consider that 500 µL of enzymatic solution will be used per brain.
      3. Introduce papain solution in a water bath 37 °C for around 20 min until the enzyme fully dissolves.
      4. Filter-sterilize the enzymatic mix with a syringe and a 0.22 µm-pore filter and keep it at 4 °C until use.
         NOTE: Papain becomes activated after 30 min at 37 °C, and it can be stored for 12 h at 4 °C.
   4. Prepare the control culture medium by adding the hormone mix solution according to Table 1 and filter-sterilize it using nitrocellulose 0.22 µm-pore filters. Keep the medium at 4 °C and warm it in a water bath at 37 °C before use.
   5. Prepare the complete medium by supplementing the control medium with EGF and bFGF just before using as per Table 1. Do not filter the complete medium, as mitogens can be lost with filtering.
2. Brain extraction and SVZ dissection
   NOTE: Mice are allowed free access to food and water prior to euthanasia. C57BL/6 mice that are 8 to 16 weeks old are used. Both sexes are used without noticeable differences in the preparation.
   1. Sterilize the dissection tools by autoclaving before the surgery. Then, submerge scalpels, tweezers, scissors, and spatulas in a beaker filled with 70% ethanol. Clean every working surface thoroughly with 70% ethanol.
   2. Sacrifice the mouse by CO₂ overdose and confirm by cervical dislocation. Spray the head with 70% ethanol to minimize the possibility of brain tissue contamination.
   3. Separate the head of the mouse from the rest of the body using scissors. Cut the skin above the skull with small scissors along the rostro-caudal axis until the cranium is completely uncovered.
   4. Expose the brain by initially cutting the skull along the sagittal suture with small scissors and then, remove the bones using fine tweezers. Be careful not to cause any damage to the brain tissue beneath the skull during this step.
   5. Carefully extract the brain from the skull using a spatula and place it into a 12-well plate containing cold DPBS. Keep the plate on ice until dissection is completed.
   6. Transfer the entire brain onto the silicon pad where the dissection will take place and apply a few drops of cold DPBS over the brain (Figure 1A).
   7. Use a scalpel to remove the OB located at the rostral end of the brain and the cerebellum situated in the caudal part while retaining the central portion of the brain containing both lateral ventricles (Figure 1B).
   8. Divide the brain along the longitudinal fissure into two hemispheres and then proceed with dissection of both hemispheres separately (Figure 1C).
   9. Working with one hemisphere, reorient it so that the medial area faces upward. Open the brain along the line of the corpus callosum, separating the cortex and striatum from the hippocampus, septum, and diencephalon, thereby exposing the lateral ventricles (Figure 1D).
      NOTE: The SVZ is situated between the ventricle's cavity and the striatum, so the subsequent steps aim to isolate the SVZ as accurately as possible from the surrounding tissue.
   10. Remove the hippocampus, septum, and diencephalon following the ventral limit of the ventricles (Figure 1E,F), the tissue beyond the rostral and caudal ends of the SVZ (Figure 1G), and the cortex following the corpus callosum (Figure 1H,I).
   11. Tilt the tissue so that the SVZ faces sideways (Figure 1J), allowing the removal of the striatal tissue beneath the SVZ (Figure 1K) to obtain a thin piece of tissue containing the NSCs (Figure 1L).
   12. Store both dissected SVZs from the same mouse in a 12-well plate containing cold, sterile DPBS until the tissue processing begins.
       NOTE: The dissection of both SVZs from one brain should take 10 min. Longer time might affect cell viability. If an increased yield of the cultures is needed, SVZs from multiple mice of the same age, sex, genetic background, and genotype can be pooled.
3. Isolation and seeding of NSCs
   1. Cut each SVZ into 4-5 small pieces to facilitate the disaggregation of the tissue.
      NOTE: From this step on, the whole procedure must be performed in sterile conditions in a laminar flow hood.
   2. Transfer the chopped SVZs to a 15 mL centrifuge tube using a sterile plastic Pasteur pipette. Make sure there are no fragments left in the plate.
   3. Let the tissue sediment at the bottom of the tube and remove the remaining DPBS.
   4. Add 500 µL of filtered papain-containing enzymatic mix per brain (2 SVZs) and incubate the tubes in a water bath at 37 °C for 30 min.
   5. After the incubation, add 5 mL of control medium pre-warmed at 37 °C to each sample.
   6. Centrifuge the samples at 300 x g for 5 min. Then, carefully remove the supernatant using a micropipette or a vacuum pump.
   7. Add 1 mL of control medium and carefully mechanically disaggregate the pellet, pipetting up and down 10x to 20x, using a fire-polished Pasteur glass pipette until a homogeneous cell suspension is obtained.
      CAUTION: This is one of the most critical steps in the protocol, as either excessive or insufficient disaggregation will negatively affect the yield of the cultures.
   8. Add 4 mL of control medium and mix by inversion to wash the cells. Centrifuge at 300 x g for 5 min. Then, remove the supernatant.
   9. Add 1 mL of complete medium and resuspend the pellet by pipetting up and down 10x to homogenize the cell suspension.
   10. Count the cell suspension using a Neubauer chamber under the microscope after diluting 1/2 an aliquot of the cell suspension with trypan blue or utilize an automatic cell counter. Make sure that cultures are disaggregated at the single-cell level.
   11. Once the cell suspension is prepared, seed the obtained cells equally in 8 wells of a 48-well plate in a final volume of 500 µL of complete medium.
       NOTE: We usually obtain 20,000 cells from two SVZs, resulting in a final seeding density of approximately 2,500 cells/mL to achieve the maximum yield in cultures.
4. Neurospheres formation
   1. Incubate cells for 5 to 7 days in vitro (DIV) in a humidified incubator at 37 °C and 5% CO₂. Under these conditions, differentiated cells die while NSCs and progenitors proliferate in response to mitogenic stimulation, forming clonal aggregates recognized as primary neurospheres.
      NOTE: Either EGF or bFGF can be used to induce neurosphere formation. However, these two growth factors are not entirely substitutable.
   2. Check the size of the neurospheres using an inverted microscope-coupled camera, and once neurospheres reach a size range of 50 to 100 µm, count all primary neurospheres formed in the well at 10x magnification. The total number of primary neurospheres formed serves as an estimate of the number of NSCs present in the SVZ niche.
   3. After a 10-day period, collect the culture medium from each well to begin passaging. Conduct subsequent passaging after 5 to 7 DIV, although it is worth noting that primary spheres might require a slightly longer duration to reach the ideal size for subculturing (around 100 µm).

Table 1: Culture media solutions. The recipe of the control medium is described. A pre-mixed hormone mix solution is made. Stock hormone solutions can be prepared in advance as indicated and stored at -20 °C. Before cell culture, supplement the control medium with EGF and bFGF to prepare complete medium. Storage specifications, stocks, and working concentrations are provided for each component. Please click here to download this Table.

2. Expansion of neurosphere cultures

1. Passage of neurospheres
   1. Collect the medium containing the neurospheres from the multiwell plates and transfer them to a 15 mL centrifuge tube. Ensure the recovery of as many cells as possible by rinsing the wells with a few milliliters of DPBS.
   2. Centrifuge neurospheres for 5 min at 300 x g. Then, remove the supernatant.
   3. Add 200 µL of enzymatic solution (see Table of Materials) to the pellet and gently tap the bottom of the tube to dislodge it. Incubate for 10 min at room temperature to facilitate neurospheres dissociation.
   4. Add 1 mL of control medium to halt the enzymatic reaction and mechanically dissociate neurospheres with a P1000 micropipette by pipetting up and down 10x to 20x.
   5. Add an additional volume of 4 mL of the control medium and mix by inversion to wash the cells.
   6. Centrifuge the cells for 5 min at 300 x g. Then, remove the supernatant.
   7. Add 1 mL of complete medium and resuspend the pellet to homogenize the cellular suspension by pipetting up and down 10x.
      NOTE: If the original number and size of neurospheres are too high, add more complete medium to dilute the cell suspension ensuring a reliable cell count.
   8. Count the cell suspension using a Neubauer chamber after diluting 1/2 an aliquot of the cell suspension with trypan blue or utilize an automatic cell counter.
   9. To expand the cell culture, seed cells at a density of 10,000 cells/cm² using complete medium in an appropriate culture plate or flask (see Table 2).
   10. Incubate cells for 5 to 7 DIV in a humidified incubator at 37 °C and 5% CO₂ for each passage.
       NOTE: Neurosphere cultures can be consistently expanded and propagated through repeated subculturing, not exceeding 10 passages, without altering their proliferation capacities. Either EGF or bFGF can be utilized for culturing; however, long-term expansion of the cultures can only be achieved in the presence of EGF.
2. Self-renewal assay
   1. After passaging neurosphere cultures, plate 1,000 individual cells in a 96-well plate, completing up to a final volume of 200 µL with complete medium, resulting in a final cell density of the cultures of 5 cells/µL.
      NOTE: The number of cells seeded must be as accurate as possible. If necessary, prepare intermediate dilutions to ensure a reliable cell suspension volume before plating. Moreover, it is recommended to plate 4 to 5 replicates per condition to reduce technical variability.
   2. Incubate cells at 37 °C and 5% CO₂ in a humidified incubator for 5 days or until neurospheres reach an adequate size (60 to 100 µm). Avoid moving or shaking the multiwell plate to prevent neurosphere aggregation.
   3. After neurosphere formation, count the number of neurospheres formed in each well using an inverted phase-contrast microscope equipped with an attached camera. The self-renewal assay can be conducted using EGF or bFGF separately.
3. Cryopreservation of NSC cultures
   NOTE: Neurosphere cultures can be cryopreserved for future use, minimizing the number of mice required. We recommend preserving cells up to passage 10.
   1. To cryopreserve cells, transfer 1 mL of the cell suspension in complete medium to a properly labeled cryotube.
   2. Add dimethyl sulfoxide (DMSO) to a final concentration of 10% to the cell suspension and gently mix by inverting the tubes.
      NOTE: DMSO serves as a cryoprotectant but can be toxic to cells, so this step should be performed swiftly.
   3. Place the tubes in a -80 °C freezer using a freezing container that enables a gradual temperature decrease at 1 °C/min. This gradual freezing process prevents cell damage caused by the formation of ice crystals during cryopreservation. For long-term storage, maintain the tubes in a liquid nitrogen tank at -196 °C.
4. Thawing of NSC cultures
   ​NOTE: When required, cells can be thawed, and cultures re-expanded for experiments.
   1. Remove the cryotubes containing frozen NSCs from the liquid nitrogen storage tank and immediately place them in a 37 °C water bath until completely thawed, approximately 5 min.
      CAUTION: It is crucial to minimize the bath time since DMSO is toxic and can affect cell viability.
   2. Quickly transfer thawed cells into a 15 mL centrifuge tube containing 5 mL of pre-warmed control medium.
   3. Centrifuge cells at 300 x g for 5 min and remove the supernatant. Add 1 mL of complete medium and resuspend the pellet by gently pipetting up and down 10x to homogenize the cellular suspension before seeding.
      NOTE: Thawing can affect cell viability; therefore, it is recommended to passage cells at least once before seeding for new experiments.
5. Mycoplasma testing
   ​NOTE: Mycoplasma contamination can compromise cell health, affecting growth rates. Therefore, mycoplasma testing is crucial to ensure the integrity and reliability of experiments.
   1. DNA extraction
      1. Collect a minimum of 100 µL of the cell culture for DNA extraction. Both supernatant and cell pellet can be used for DNA extraction. Store this aliquot at -20 °C until extraction.
      2. Pierce the tube lid and boil the aliquot at 99 °C for 15 min using a thermoblock.
         NOTE: Piercing of tube lids avoids tubes opening during boiling at high temperatures.
      3. Centrifuge for 5 min at 16,900 x g. Transfer the supernatant containing the DNA to a new tube. Store the tube with the DNA at -20 °C until PCR testing.
   2. PCR for Mycoplasma detection
      1. Prepare the PCR mix using the following components: 8.9 µL of nuclease-free water, 3 µL of 5x reaction buffer, 1.2 µL of 25 mM MgCl₂, 0.25 µL of 2.5 mM dNTPs, 0.15 µL of Taq polymerase and 0.25 µL of each mycoplasma-specific primer at a concentration of 10 µM (Forward, Fw: 5'GATGTTTAGCCGGGTCGAGAG3' and Reverse, Rv: 5'GATGTCAAGAGTGGGTAAGGTT3').
      2. Add 1 µL of genomic extracted DNA. Prepare the PCR mix in a laminar flow hood to prevent DNA contamination.
      3. Incubate the reaction in a thermal cycler as follows: initial denaturation at 95 °C for 5 min followed by 35 cycles of denaturation at 95 °C for 30 s, annealing at 53 °C for 1 min, extension at 72 °C for 30 s, and a final extension at 72 °C for 5 min.
      4. To identify the size of the resulting band (around 500 bp), load 15 µL of the DNA PCR product mixed with 3 µL of 6x loading dye solution onto a 2% agarose gel and run the electrophoresis for 40 to 50 min at 100 V.

Table 2: Plates and flasks used for culturing. The dimensions and volumes of the most commonly used seeding plates and flasks are provided. The table includes the diameter, growth area, and medium volume used for each vessel, along with an example of the number of NSCs to be seeded under expansion conditions (10,000 cells/cm²). Please click here to download this Table.

3. Nucleofection of neurosphere cultures

1. Preparation of reagents
   1. DNA preparation
      1. Prepare 25 µL of competent E. coli DH5α cells in a 1.5 mL tube for transformation.
      2. Add 1 µL of the plasmid of interest (at 100 to 200 ng/µL) to the cells and subject them to a thermal shock at 42 °C for 45 s. Here, a plasmid containing a short hairpin (sh)RNA for silencing Snrpn expression and a control plasmid shSCRAMBLE are used.
         NOTE: Keep competent cells on ice throughout the procedure to maintain transformation effectiveness.
      3. Plate the transformed cells on LB agar plates with the appropriate antibiotic and incubate them at 37 °C for 18 h. Keep the Petri dishes inverted to prevent droplet formation due to condensation, which might disturb bacterial colonies.
         NOTE: Add appropriate antibiotics to the LB culture medium to exclusively allow the growth of transformed bacteria.
      4. Using a sterile pipette tip, pick individual bacterial colonies and transfer them to an Erlenmeyer flask previously filled with 250 mL of LB to initiate a liquid culture. Avoid picking small satellite colonies positioned around the larger target colonies, as they might not have incorporated the desired plasmid.
      5. Incubate the flask while vigorously shaking on an orbital shaker at 37 °C overnight.
      6. Utilize an endotoxin-free maxiprep kit and extract pure plasmid DNA from the bacterial cultures following the manufacturer's instructions. Reconstitute the DNA in the appropriate volume of endotoxin-free water to obtain a plasmid concentration of 1 to 2 µg/µL.
         NOTE: The maxiprep yield depends on the culture growth. Usually, resuspending the plasmid DNA pellet in a volume of 200 µL will result in the desired DNA concentration.
      7. Store the tube with the plasmid DNA at -20 °C until use. Prepare a 1.5 mL tube with a final volume of 5 µL containing 2 to 6 µg of the plasmid DNA intended for nucleofection.
   2. Material preparation
      1. Prepare the nucleofection solution. If using the recommended kit (see Table of Materials), prepare 95 µL of the nucleofection solution in a 1.5 mL tube for each desired nucleofection. Pipettes and cuvettes are usually included in the kit. Prepare one of each for every nucleofection condition.
      2. Prepare a T25 flask filled with 4 mL of pre-warmed complete medium per condition and keep it in the incubator until use.
2. Nucleofection of NSCs
   1. Use 1 to 2 x 10⁶ individual cells per condition for assessment. Centrifuge the cells for 5 min at 300 x g, then remove the supernatant.
   2. Add 1 mL of DPBS and resuspend the pellet by pipetting up and down 10x to ensure a homogeneous cellular suspension.
   3. Repeat centrifugation (step 2.1.6) for a second wash and finally resuspend the pellet in 95 µL of nucleofection solution.
      NOTE: Washing steps are crucial to remove antibiotics from the control medium and ensure effective nucleofection.
   4. Combine the 95 µL of cells with a premixed solution containing 5 µL of each plasmid (2 µg total) selected for nucleofection.
   5. Using a P200 micropipette, transfer this solution to a cuvette and place it in the nucleofector device.
   6. Nucleofect the cells with the desired plasmids using the optimized NSC program of the nucleofector system.
   7. After completing the electroporation, carefully introduce warm complete medium to the cuvette using the Pasteur pipettes provided in the kit. Subsequently, transfer the contents of the cuvette to a previously prepared T25 flask containing pre-warmed complete medium. After electroporation, there might be cell death, DNA precipitation, and the formation of a viscous aggregate. Avoid transferring this to the T25 flask, as it might reduce cell survival.
      NOTE: This step is crucial and should be performed as quickly as possible to ensure a high yield of nucleofection and subsequent culture survival.
   8. Incubate nucleofected cells at 37 °C and 5% CO₂ in a humidified incubator for 3 to 5 days.
3. Selection of nucleofected cells
   1. Strategy A: Flow cytometry selection by fluorescence-activated cell sorting (FACS)
      1. After 3 to 5 days of nucleofection, passage the nucleofected cultures following step 2.1 and collect all individualized cells obtained.
      2. Filter samples through a 40 µm cell strainer to remove cell aggregates and analyze them with the cell sorter flow cytometer.
         NOTE: It is crucial to sort the same number of cells in each condition to obtain comparative results.
      3. Using the cytometer's software, gate alive cells based on two measures: forward scattered light (FSC-A) that corresponds to cell size and side scattered light (SSC-A) that corresponds to intracellular complexity. Select the population of living cells characterized by high FSC-A and high SSC-A.
         NOTE: Optionally, add 0.1 µg/mL of 4′,6-diamidino-2-phenylindole (DAPI) to cell suspension before sorting. This will allow the discarding of dead cells and apoptotic bodies from the gating.
      4. Discard cell doublets and triplets by plotting FSC-A versus FSC-H parameters. Choose the events included in the diagonal of the plot, representing singlets.
         NOTE: When cells pass through the laser, the cytometer detects voltage changes over time. FSC-A represents the signal area, and FSC-H signifies peak height. Singlets show a proportional relationship between FSC-A and FSC-H (located on the diagonal of the plot), while cell aggregates display a greater FSC-A relative to FSC-H due to a longer signal.
      5. Select nucleofected cells based on the fluorescence intensity of the reporter gene encoded by the plasmid used. For instance, when using GFP⁺ cells can be selected based on FITC-A fluorescence levels versus SSC-A plot.
      6. Isolate the desired number of target cells for each condition and collect them directly in a collection tube filled with complete medium.
      7. Seed nucleofected NSCs at a density of 10,000 cells/cm². Incubate the sorted cell cultures at 37 °C and 5% CO₂ in a humidified incubator.
   2. Strategy B: Selection using antibiotic resistance encoded in the plasmid
      1. After 48 h of nucleofection, add the appropriate antibiotic concentration to select resistant cells.
         NOTE: Before starting, it is recommended to test multiple concentrations of the antibiotic on non-nucleofected cells. The optimal antibiotic concentration is the lowest amount at which all non-resistant cells are eliminated. For example, we determined an optimal concentration of 4 µg/mL antibiotic blasticidin for NSC cultures.
      2. Incubate the cells at 37 °C and 5% CO₂ in a humidified incubator for 48 h.
      3. After 48 h of antibiotic treatment, subculture the nucleofected cells according to step 2.1. If the neurospheres remain small, remove the antibiotic from the culture medium by centrifugation at 300 x g for 5 min. Resuspend the cells and culture them in a flask using only complete medium. Incubate the cells at 37 °C and 5% CO₂ in a humidified incubator until they can be expanded following step 2.1.