An Enzyme- and Serum-free Neural Stem Cell Culture Model for EMT Investigation Suited for Drug Discovery

All animal procedures followed the 'Guide for the Care and Use of Laboratory Animals' (NIH publication, 8^th edition, 2011) and were approved by the Animal Welfare Committee of Basel (Swiss Guidelines for the Care and Use of Animals). By these guidelines the animal protocol is considered of "lowest animal severity grade".

1. Preparation of Expansion Medium

Note: Work in aseptic conditions as standard for tissue culture.

1. Take two 15 ml tubes, and add 5 ml of L-glutamine-free DMEM/F12 (1:1) from a 500 ml medium bottle into each of the two 15 ml tubes. To the first 15 ml tube, add 50 mg human apo-transferrin, 50 μl of putrescine 1 M stock (0.1 mM final concentration) and 30 μl of sodium selenium 500 µM stock (final concentration 30 nM).
   1. Filter through a 0.2 µm syringe filter into the original DMEM/F12 bottle.
2. To the second 15 ml tube, add 12.5 mg insulin. Add 6 – 9 drops of 1 M NaOH. Vortex to dissolve insulin completely. Filter through a 0.2 µm syringe filter into the original DMEM/F12 bottle.
   Note: NaOH is toxic and corrosive. Use protective gloves, coat, and safety goggles.
3. Add 5 ml penicillin (10,000 units/ml) / streptomycin (10,000 µg/ml) / amphotericin B (25 µg/ml) to the DMEM/F12 medium bottle.
4. Add 5 ml of 200 mM L-glutamine stock freshly thawed or oligopeptides containing glutamine (200 mM L-alanyl-L-glutamine dipeptide) to the DMEM/F12 medium bottle.
5. Shake well. Prepare 50 ml aliquots.
   Note: Expansion medium can be stored for at least 2 weeks at 4 °C. Aliquoted tubes show less pH changes compared to medium kept in 500 ml bottles.

2. Preparation of Passaging Medium

1. To 1 L Ca²⁺– and Mg²⁺-free HBSS media, add 990.85 mg Glucose (5 mM final concentration) and 840.10 mg NaHCO₃ (10 mM final concentration). Adjust pH to 7.3 with HCl (1 M stock). Filter sterilize with 0.2 µm filter and make 50 ml aliquots.
   Note: Passaging medium can be stored for at least 4 weeks at 4 °C.

3. Preparation of Growth Factors

1. Prepare sterile 1x PBS with 1% BSA (PBS-BSA) alone or with hydrochloric acid (HCl) at 4 mM (PBS-BSA-HCl).
   Caution: HCl is corrosive and toxic and requires special safety measures (coats, goggles, gloves, hood).
2. Dissolve 10 µg/ml recombinant human Fibroblast Growth Factor 2 (rhFGF2) in PBS-BSA (10 ng/ml final concentration), 10 µg/ml recombinant human Bone Morphogenetic Protein 4 (rhBMP4) in PBS-BSA (10 ng/ml final concentration), and 20 µg/ml recombinant human Transforming Growth Factor β 1 (rhTGFβ1) in PBS-BSA-HCl (40 ng/ml final concentration).
   1. Do not filter sterilize. Prepare aliquots.
      Note: Growth factor aliquots can be stored at -20 °C for at least 2 years.

4. Coating of Cell Culture Dishes

1. Dissolve 1,875 mg Poly-L-ornithine (PLO) in 500 ml distilled H₂O to prepare a 250x PLO stock. Filter sterilize using a 0.2 µm syringe filter and make 2 ml aliquots.
   Note: These aliquots can be stored at -20 °C for at least 2 years.
2. Dissolve 1 mg bovine fibronectin in 1 ml sterile-distilled H₂O to prepare 1,000x fibronectin stock. Do not vortex as this may clot the sticky protein. Shake gently by hand for 10 min.
   1. Incubate for 1 hr at room temperature with occasional shaking. Check for complete dissolution. Warm the solution to less than 37 °C for the complete dissolution of fibronectin. Do not filter-sterilize.
      Note: The solution may be stored at 4 °C for up to 6 months.
3. Use plasma-pretreated plastic cell culture dishes (100 mm or other sizes as required for the experiment). To assess migration, use 35 mm dishes with 500 µm grid. Incubate dishes overnight with 2 ml 1x PLO for 12 hr at 37 °C in a 5% CO₂ tissue culture incubator. Wash twice with 2 ml 1x PBS.
4. Add 2 ml 1x fibronectin (1 μg/ml final concentration) and incubate dishes for a minimum of 12 hr at 37 °C in a 5% CO₂ incubator. Remove fibronectin just before plating the cells.
   Note: Fibronectin-coated dishes may be stored for at least 2 weeks at 37 °C.

5. Standardized Dissection and Preparation of the Cortical Subventricular Zone (SVZ)

1. Obtain rat embryonic day 14.5 (E14.5, Sprague-Dawley) and mouse E13.5 (C57BL/6) embryos by timed-mating. Mate animals from 18:00 until 08:00 the next morning. Noon after the day of mating is considered E0.5.
2. Anesthetize the pregnant animal with 5% Isoflurane and 0.8 L/min oxygen flow. Check response by paw compression with sharp dissection forceps. Decapitate the animal. Avoid CO₂ asphyxiation as CO₂ affects stem cell recovery.
3. Retrieve embryos by caesarean section.
   1. Place animal in supine position and disinfect fur with 70% ethanol. Use surgical forceps and scissors to make V-shaped skin incision of about 8 cm in the lower abdomen above the uterus. Incise only skin with fur while keeping muscular walls intact.
   2. Use fresh forceps and scissors to incise the muscles and the abdominal muscular wall to enter the peritoneum.
   3. Identify the uterus in the lower posterior peritoneum. Remove the uterus by sharp separation from the surrounding tissues.
   4. Wash the uterus with embryos with sterile 1x PBS and place in ice-cold 1x PBS.
   5. Use fine-tipped scissors to open the uterine walls to release embryo by embryo under a dissection microscope (3X magnification, Figure 1B). Use a pair of forceps to remove the embryonic hulls (Figure 1B). Keep embryos in expansion medium on ice.
   6. Verify the correct developmental stage by the Atlas of the Developing Rat Nervous System⁵. The correct embryo size is shown in Figure 1B.
   7. Check correct age further by presence of beginning digit formation in the rat forelimb (FL) as illustrated in Figure 1A.
      Note: The hind limb (HL) does not yet show digit separation (Figure 1B).
4. For dissection, transfer embryos to uncoated Petri dishes filled with ice-cold 1x PBS. Perform dissection under a stereo dissection microscope (3X to 20X magnification) using autoclaved fine forceps.
   Note: The Petri dishes prevent attachment of tissue to the dish surface as may happen with plasma-coated cell culture dishes. Sharpened tungsten wire needles or similar are also helpful for certain steps of dissection.
5. Remove the head skin and skull starting at the intersection of the developing telencephalon (TEL) and diencephalon (DI) (Figure 1B) by pulling simultaneously anteriorly and posteriorly with two forceps in order to gain access to the developing neural tube (Figure 1C).
6. Identify the midbrain-hindbrain-boundary (MHB, black arrow head in Figure 1B-D), separating the mesencephalon (MES) from the rhombencephalon. Cut the rhombencephalon just at or below the MHB (Figure 1D, triangular arrow).
   ​Note: The additional subcutaneous space at the MHB facilitates skin removal without harming the neural tube.
7. Sever the connection between the telencephalon/diencephalon at the skull base with the facial skeleton (Figure 1E). This results in a block containing the telencephalon, diencephalon and mesencephalon (TEL-DI-MES) (Figures 1F-H).
8. Transfer the TEL-DI-MES block into expansion medium on ice. Keep it completely covered in expansion medium in an uncoated Petri dish. Hold the block at the mesencephalon with a pair of forceps to avoid touching the telencephalon that contains the cortical SVZ with NSCs.
9. Repeat steps 5.5 to 5.8 with all embryos (Figure 1F-H).
10. Transfer one TEL-DI-MES block into fresh ice-cold 1x PBS. Cut along the dotted line in the anterior mesencephalon (Figure 1F-H), separating the mesencephalon from the diencephalon, as shown in Figure 1I.
11. Separate the DI from the TEL by cutting along the dashed lines in Figure 1F. See isolated telencephalon in Figure 1J.
12. Split the two telencephalic hemispheres, cutting along the dotted line in Figure 1J. This results in two separated hemispheres as illustrated in Figure 1K.
    Note: The left hemisphere is magnified in Figure 1L.
13. Identify the medial ganglionic eminences (MGE, Figure 2A; *) and lateral ganglionic eminences (LGE, Figure 2A; +) visible through the future Foramen interventriculare.
    1. Using forceps or needle, dissect along a straight line at the intersection between the MGE and LGE and the anterior cortex (ant-ctx, Figure 2B). Cut a straight line through cortex, hippocampus (hip) and choroid plexus (CP). This separates the anterior cortex including the olfactory bulb (ob) from the ganglionic eminences (GE, Figures 2B, C).
14. Cut a second straight line at the intersection of the caudal ganglionic eminence (CGE, Figure 2C, D) and the posterior cortex (Figure 2C), again cutting through cortex, hippocampus and choroid plexus.
15. Flatten out the telencephalon. Completely remove the posterior pole and the olfactory bulb (Figure 2D). Identify the cortical hem containing the hippocampus and the choroid plexus (Figure 2C), as defined previously ^6,7.
    ​Note: The hippocampus has a thinner neuroepithelium than the cortex. The CP contains red vessels.
    1. Separate the cortical hem from the cortex, leaving the size of the cortex identical to the size of the ganglionic eminences (Figure 2D). This results in a block of the cortex (the target tissue) and the ganglionic eminences (GE, Figure 2D).
16. Flip the cortex-GE block with the ventricular (inner) side to the dish surface.
    Note: The outer surface of the hemisphere will face the experimenter (Figure 2E). On the outer surface are the blood vessel-containing meninges (Figure 2E).
    1. Pin the GE to the dish surface with the left hand and peel the meninges off with a pair of forceps in the right (dominant) hand (Figure 2F).
       Note: This is the technically most demanding step because the meninges strongly adhere to the cortex. The separation of meninges from the cortex is facilitated by cutting a completely straight line (not jagged) in steps 5.13.1 and 5.14.
17. Repeat steps 5.12 to 5.16 for the other hemisphere and all embryos. Cut the cortex along the LGE in a distance of half the main diameter of the LGE (Figure 2G, 2H). The dissected cortex spans an area of 1.2 mm x 2.4 mm (Figure 2H) and includes the SVZ/germinal layer with the NSCs.

6. Preparation of Explant Cultures

1. Cut the cortex into explants of less than 400 µm diameter (Figure 2I). Use a 400 µm grid (Supplementary Figure 1) positioned below the dissection Petri dish for reference (Figure 2H and 2I). Transfer all explants into a fresh Petri dish with ice-cold expansion medium.
2. Remove the fibronectin from a tissue culture dish (step 4.4). Allow it to dry. Add 1 ml of cold expansion medium to the center of the 35 mm dish with grid dimension of 10 mm x 10 mm (Figure 3). Allow the medium to form a spherical drop (Figure 3).
3. Insert up to 8 explants to the center of the drop. The explants should be ideally located on the grid with at least 3 mm distance between each other for the migration analysis (see section 8).
4. Incubate the dish for about 1 hr in a cell culture incubator (37 °C, 21% O₂, 5% CO₂) for attachment of the explants. Allow the explants to settle and attach to the fibronectin-coated surface. Do not shake the dish, since the explants may detach and move out of the optimal grid center.
5. After 1 hr incubation (37 °C, 21% O₂, 5% CO₂), fill up the dish to a total volume of 2 ml expansion medium plus the growth factors or substances of interest to test and incubate.
   Note: Neither enzymatic digestion, nor serum or centrifugation are necessary for explant preparation. This is optimal for cell integrity.

7. Preparation of NSC Single Cell Culture

1. Warm up expansion and passaging medium at 37 °C.
2. Transfer the dissected cortex pieces (from step 5.17) into a 15 ml tube with cold expansion medium (tube A). Centrifuge the cortex pieces for 5 min at 1,200 x g. Aspirate the supernatant. Add 200 µl pre-warmed expansion medium to resuspend the cortex pieces.
3. Add 700 µl of pre-warmed passaging medium to the 15 ml tube with a P1,000 tip (tube A). Gently resuspend the pellet. Place the tip at the bottom of the 15 ml tube. Gently and slowly dissociate the tissue pieces by suctioning three times with the P1,000 tip.
   Note: Passaging medium promotes cell separation and is required to avoid the use of enzymes.
4. Allow the tube to sit for 30 to 60 sec for larger (heavier) undissociated pieces to settle at the bottom. Single dissociated cells will remain in the supernatant. Transfer about 700 µl of the supernatant to a fresh 15 ml tube (tube B).
5. Repeat steps 7.3 and 7.4 to dissociate the larger pieces.
6. Repeat steps 7.3 and 7.4 a second time to dissociate the larger pieces. Transfer all supernatant to the fresh 15 ml tube (tube B).
7. Add expansion medium to a total volume of 5 ml. Count cells using a hemocytometer. Assess dead cells by trypan-blue staining.
   Note: Small NSCs tolerate the steps 7.2 to 7.6 better than larger cells. From 10 embryos expect about 4 x 10⁶ cells with 10% dead cells.
8. For expansion of NSCs, plate 1.5 x 10⁶ cells per 10 cm fibronectin-coated tissue culture dish in a volume of 8 ml expansion medium. For standard expansion, add 8 µl of 1,000x rhFGF2 stock. Add 8 µl rhFGF2 every day and change media every other day.
   Note: The cortex at this developmental stage contains a majority of NSCs and only a minority of differentiated cells. The differentiated minority cells do not respond to the rhFGF2-treatment and die during the expansion culture.
9. Passage expanded NSCs at 60% confluence.
   1. To passage NSCs, remove expansion medium. Wash cells quickly three times with 5 ml passaging medium. Wait 2 – 4 min. Without Ca²⁺ and Mg²⁺ the cells slowly detach from the surface, rounding up. Verify detachment of cells under the phase microscope. Wait another few minutes, if needed, until visual detachment from surface is observed.
      Note: Single cells will completely detach, but most cells will still adhere to the dish surface. The duration needed for detachment is dependent on the duration of fibronectin coating. A short fibronectin coating (12 hr or less) results in faster cell detachment. For longer experiments (> 1 week) fibronectin coating of more than 24 hr is recommended.
10. Use a 10 ml pipette to gently detach the cells from the surface. Collect the 5 ml passaging medium with cells and transfer it to a fresh 15 ml tube. Repeat with an additional 5 ml of passaging medium.
11. Dissociate the cells in the 15 ml tube by pipetting up and down three times, placing the 10 ml pipet at the bottom of the tube. Spin the tube for 15 min at 1,200 x g at room temperature. Remove the supernatant and resuspend the pellet in 2 ml expansion medium. Count cells using a hemocytometer. Use trypan blue to assess dead cells.
    ​Note: After expansion to 60% confluence, expect 4 – 5 x 10⁶ cells with a dead cell count at around 10%.
12. Plate 8 x 10⁵ cells per 10 cm dish for further passages.

8. Migration Assessment

1. For migration assessment, isolate explants according to Section 5. Seed the explants in 35 mm grid dishes. One hour after plating, add FGF2 (rhFGF2). Place the dishes in a 37 °C incubator for 2 days.
   ​Note: For optimal explant attachment, avoid moving the dishes.
2. Remove medium by 1,000 µl tip. Add 2 ml of expansion medium by 1,000 µl tip starting at the inner circle (Figure 3). This reduces surface tension on explants.
3. Add growth factors alone or in combination as required for experiment. Use FGF2/BMP4/TGFβ1 as positive control. Note: In this experiment, 2 µl FGF2 per 35 mm dish, 2 µl BMP4 and 4 µl TGFβ1 were added alone and in combination (Figure 5).
   1. Add additional factors and/or testable substances. Keep dishes again untouched for 2 days at 37 °C.
4. Take photographs of explants on an inverted microscope.
   Note: Living explants yield better phase contrast images than fixed cells. Both may be used.
5. For migration assessment, use a graphics software that allows pixel measurements (e.g., Fiji⁸).
6. Measure the dish grid of 500 µm distance in pixels and use it as internal reference.
7. Define the center of the explant as migration start point. Measure the distance between the center of the explant and the outermost group of 10 cells.
   Note: All cells migrate centrifugally away from the explant. They spontaneously form a sheet at the periphery under the circumstances tested (Figure 5).

9. Invasion Assessment

1. Prepare fibronectin-coated tissue culture 24-well plates according to Protocol Section 4.
2. Place 300 µl of warm expansion medium into the top well of cell invasion chambers. Incubate the chambers for 30 min at 37 °C and 5% CO₂.
3. Remove the expansion medium from the top well and place the Boyden chamber into the coated 24-well plate.
4. Add 500 µl of warm expansion medium with 0.5 µl of rhFGF2 and 0.5 µl rhBMP4 to the bottom well.
5. Passage NSCs and count as described in Section 7. Passages 1 to 4 are suitable.
6. Plate 5 x 10⁵ cells in a volume of 300 µl of warm expansion medium with 0.3 µl of rhFGF2 and 0.3 µl of rhBMP4 in the top well of the Boyden chamber.
7. Culture the seeded NSCs for 24 hr.
8. Add additional factors and/or testable substances to the chamber and culture the cells for another 48 hr.
   Note: If the cells are invasive, they will pass from the top well through the basement membrane layer to the bottom well. Invasive cells will cling to the bottom of the Boyden chamber.
9. Follow the protocol provided by the manufacturer. Use cotton swabs (included in the invasion assay kit) to remove the non-invasive cells in the top well by cleaning twice.
10. Remove the medium from the wells and add 500 µl expansion medium with a plasma membrane stain for living cells and with the nuclear dye DAPI to the wells.
11. Incubate for 10 min at 37 °C and 5% CO₂.
    Note: The dyes will integrate into the membrane and the nucleus of the invasive cells, respectively, for optimal visualization⁸. Option: Omit plasma membrane dye if multicolor immunocytochemistry is planned.
12. Remove the medium with the dyes and wash twice with expansion medium. Visualize and count invasive cells with an inverted fluorescence microscope as demonstrated previously⁸.
    Note: Some invasive cells may have detached from the bottom of the chamber and have dropped to the well surface below where they stick to the coated surface. For complete analysis include the cells at the well surface.
13. Remove the medium and fix the cells in ice-cold fresh 4% PFA for 10 min. Wash 3x with 1x PBS. Perform immunocytochemistry on invasive cells, as previously described^8-10.