Generation of 3-D Collagen-based Hydrogels to Analyze Axonal Growth and Behavior During Nervous System Development

All animal experiments were performed under the guidelines and protocols of the Ethical Committee for Animal Experimentation (CEEA) of the University of Barcelona, and the protocol for the use of rodents in this study was reviewed and approved by the CEEA of the University of Barcelona (CEEA approval #276/16 and 141/15). 

1. Purification of Rat Tail Collagen

1. Collect adult Sprague-Dawley rat tails (8-9 weeks old) after sacrificing the animal following ethical guidelines and rinse in 95% ethanol. Place 2-4 tails on ice (4 °C) and keep them covered with ice during the process.  
2. To obtain tail tendons, fix the tail at the most caudal vertebrae of the tail using a hemostat and compress the tail with a second hemostat positioned around 5-7 mm from the first. Break the tail by twisting it sharply with both hemostats. To do this, fold/unfold the vertebrae several times until it breaks.
3. Pull the vertebrae slowly with the hemostat to detach the tendons from their sheath as it comes out. At this moment, cut tendons with small scissors. Keep these tendon pieces in a sterile 100 mm Petri dish on ice.
4. Repeat the clamping and sliding for the rest of the tail until the tendons are totally extracted.
5. Repeat steps 1.2-1.4 for all the obtained tails.
6. Observe the tendons under a microscope. Discard blood vessels by cutting with small scissors and holding with straight fine forceps to improve the tendon’s purity and rinse the tendons 3 times with ultrapure water.
7. Collect 3-4 g of wet tendons. Dissolve the tendons in 150 mL of 3% glacial acetic acid at 4 °C in a 200-250 mL glass conical flask for 24-36 h, under gentle stirring.
8. Centrifuge at 20,000 x g for 1 h. In parallel, prepare the dialysis tubing membrane by cutting a piece of around 10-15 cm in length and boil it in ultrapure water containing 1 mM ethylenediaminetetraacetic acid (EDTA) for 15 min. Thereafter, gently rinse the dialysis membrane thoroughly with ultrapure water.
9. After centrifugation, collect the supernatant in the dialysis tubing membrane by decantation. The pellet contains acidic insoluble material (non-collagenous proteins) and the supernatant contains soluble collagen proteins.
10. Dialyze the supernatant against 2 L of sterile 0.1x Minimum Essential Medium Eagle (MEM), pH 4.0 in a 2-5 L glass beaker. Dialyze for 3 days. Change 0.1x MEM solution at least twice a day checking the pH at every change before using. If pH has turned basic, modify it with a few drops of 0.1 M acetic acid until pH is 4.0.
11. After dialysis, add antibiotics (1.5 mL of penicillin/streptomycin (Pen-Strep)) to the dialyzed collagen solution. Make 5 mL aliquots of the collagen stock solution and store at 4 °C.
    NOTE: From this point, all handling procedures must be performed under sterile conditions in a laminar flow hood.
12. Proceed with the gelation test of the prepared collagen stock solution following the next steps.
    1. Since the stock collagen is usually too concentrated, prepare 3 working dilutions (75%, 50%, and 25% collagen solution) by diluting the stock in 0.1x MEM, pH 4.0. The final volume recommended for each working dilution is 5 mL. In each condition, check the protein concentration using a protein colorimetric assay.
    2. Place several empty 1.5 mL conical centrifuge tubes (one for each working dilution), 10x MEM tubes, 7.5% sodium bicarbonate solution and different working dilutions of collagen on ice. Wait until these are cooled.
    3. Add 40-50 µL of 10x MEM to a cold (4 °C) centrifuge tube. Next, mix it gently with 7-8 μL of sodium bicarbonate solution.
    4. Add 310-330 μL of one of the different collagen dilutions to this tube and mix it gently with the pipette avoiding any bubble formation. Keep the collagen-MEM-sodium bicarbonate mixture on ice (4 °C) for at least 5 min.
    5. Pipette 10-25 μL of the mixture into a 35 mm Petri dish.
    6. Place the Petri dish in the CO₂ incubator set at 37 °C until the gelation of the hydrogel (± 15-20 min) is observed.
    7. Repeat steps 1.12.3-1.12.6 for the remaining collagen dilutions in different Petri dishes.
    8. Select the collagen working dilution that renders the best results.
       NOTE: The best gelled hydrogel should have a uniform translucent texture, grey color and should not be lumpy or stringy. In our experience, a dilution of 3:1 (Collagen: 0.1x MEM) generates the best experimental results.

2. Preparation of Cell (COS1) Aggregates Genetically-modified to Secrete a Candidate Molecule in 3-D Collagen Hydrogels

1. Plate 2 x 10⁶ COS1 cells into a 35 mm Petri dishes and incubate with complete culture medium composed of 100 mL of D-MEM containing 10% (vol/vol) heat-inactivated fetal bovine serum, 0.5% (wt/vol) glutamine and 1% (wt/vol) Pen/Strep in a cell culture incubator, in order to reach 70-80% confluency overnight. Prepare one Petri dish for each transfection procedure.
2. The following day transfect COS1 cells with the DNA encoding the candidate molecule (Netrin-1 or Sema3E) using liposome-based transfection method following the manufacturer's instructions.
   1. To do this, mix 250 µL of serum-free medium and DNA (1-2 μg per condition) to a 1.5 mL centrifuge tube (DNA tube) and mix. Incubate at room temperature (RT) for 5 min. Prepare a second tube (liposomal tube) by adding 240 µL of the serum-free medium and 10 µL of the liposomal transfection reagent. Incubate at RT for 5 min.
   2. After incubation, add the content of the DNA-tube to the liposomal tube and mix gently. Now incubate at RT for 15 min. Replace the medium on the cultured cells with 1.5 mL of the serum-free medium and add the DNA-liposomes mixture to the Petri dish slowly dropwise. Incubate for 3 h in the CO₂ cell culture incubator.
3. After 3 h of transfection incubation, replace the medium with the complete culture medium and incubate overnight in the incubator.
4. Next day, rinse the cells with 0.1 M Dulbecco’s phosphate buffered saline (D-PBS), treat cultures with Trypsin-EDTA (800 µL per each dish for 5-15 min in the CO₂ incubator) and collect detached cells with 15 mL of complete culture medium.
5. Centrifuge the cells at 4 °C at 130 x g for 5 min. After centrifugation, remove media and preserve the pellet containing COS1 cells on ice.
6. Repeat steps 1.12.3-1.12.6 to prepare the collagen working mixture.
7. Add 100-150 μL of the collagen mixture to the pellet of the transfected cells and mix gently by pipetting up and down and spread 45-50 µL of this mixture onto a Petri dish (35 mm diameter) to form a uniform band of collagen-cells of around 1-1.5 cm in length. Place the dish in the incubator at 37 °C (5% CO₂) until the gelation (± 15-20 min) is observed.
8. Prepare a second strip containing control cells (mock transfection) in a second culture dish and plate it in the incubator (± 15-20 min) and when gelation is completed add 3-4 mL of warmed (37 °C) COS1 complete culture medium to each dish containing the gelled collagen-cells strips and keep them in the CO₂ cell culture incubator. Thereafter, cut the collagen-cells strips to generate small square pieces (400 to 500 µm in length) using a fine scalpel or a tissue chopper.
9. Transfer all the sections from the same transfection condition to a Petri dish containing 3-3.5 mL of neuronal culture media (NCM) and check under a dissection microscope the quality of the pieces. Again, keep them in the CO₂ cell culture incubator.
   NOTE: Neuronal culture medium consists of Neurobasal medium containing 1-5% (vol/vol) heat-inactivated horse serum, 2 mM glutamine, 0.5% (wt/vol) glucose, 1% (wt/vol) Pen-Strep solution and 0.044% (wt/vol) NaHCO₃. Ensure that the pH is between 7.2-7.3.

3. Generation of Embryonic Explant for Culture

1. Sterilize the surgical tools (scissors, scalpel blade handle, straight and curved forceps) by autoclaving following routine sterilization guidelines. In parallel prepare 500 mL of Hank’s balanced salt solution-glucose buffer and 4-5 Petri dishes (100 mm diameter) containing 10 mL of HBSS-G. Place these plates on ice (4 °C).
2. Sacrifice the pregnant female rat (embryonic day 16.5) outside the sterile area, following the approved ethical procedures. Cut the embryo horns with scissors from the abdominal cavity and place it into a large Petri dish containing cold HBSS-G.
3. Place the dish in the laminar flow hood and extract the embryos with straight forceps. Place them into a new dish containing cold HBSS-G. Next, remove the skin of the embryo using small forceps and carefully dissect the brain using the curved and straight forceps. Place them into a dish containing cold HBSS-G.
4. Under a dissecting microscope, cut the brain in half along the midline to separate both the hemispheres with the scalpel or fine scissors and remove the diencephalon, the meninges and blood vessels from the brain pieces with fine forceps.
5. Repeat steps 3.3-3.4 with the rest of the embryos. Do not delay the dissection for more than 2 h to preserve the tissue quality.
6. Clean all parts of the tissue chopper with 100% ethanol (especially the polytetrafluoroethylene (PTFE) cutting plate and the razor blade). Keep the tissue chopper in the laminar flux hood under UV illumination for 15 min.
7. Transfer each tissue piece (e.g., cortex with hippocampus) to the cutting plate of the tissue chopper. For hippocampus, place it perpendicular to the razor blade and obtain tissue sections of 450-500 µm in thickness.
8. Prepare several 35 mm Petri dishes with 3-4 mL of complete NCM and transfer tissue pieces from the tissue chopper plate to the Petri dishes. Many dishes may be needed as regions of interest are dissected.
9. Finish the tissue dissection in the complete NCM using fine tungsten needles. Check the quality of the obtained slices under the dissecting microscope. Ensure that the layers are clearly identifiable in the darkfield optics and dissect the region of interest (e.g., CA region of the hippocampus) with tungsten needles. Discard damaged slices. Keep hippocampal explants in complete NCM medium in the CO₂ incubator.

4. Preparation of 3-D Co-cultures in Collagen Hydrogels

1. Place several sterile 4-well culture plates in the laminar flow hood and prepare a collagen working mixture as previously indicated in steps 1.12.2-1.12.6.
2. Place 15-20 μL of the hydrogel mixture into the bottom of a well to produce a circular collagen base. Repeat this step for the rest of the wells. Do not prepare more than five plates at the same time to avoid excessive liquid evaporation from the hydrogel base.
3. Keep the dishes in the incubator until the complete gelation (± 15-20 min) is observed and take the plates out of the incubator only when the gelation is completed. Check the quality of the gelled collagen.
4. Transfer a small piece of COS1 cell aggregate with a pipette. Place it onto the hydrogel base and place a tissue piece on the same base with a pipette close to the piece of cells aggregate at one explant-size.
5. Prepare a new working collagen mixture on ice as in steps 1.12.2-1.12.6.
6. Gently pipette 15-20 μL of this new mixture and cover the explant and cell aggregate. A sandwich-like hydrogel culture will be observed. At this moment, re-orientate the explant with a fine tungsten needle (do not touch the COS1 cell aggregate!), so it faces the cell aggregate at ± 500-600 μm.
7. Return the plate to the incubator until the gelation is observed (± 10-15 min), and 0.5 mL of complete NCM supplemented with 2% B27 supplement and keep cultures for 36 to 48 h in the incubator (37 °C, 5% CO₂).

5. Fixation of Explant-cell Aggregate Co-cultures and Immunocytochemical Procedure

1. After 36-48 h of incubation, remove the medium and rinse with 0.1 M phosphate buffered saline (PBS), pH 7.3. Thereafter, fix the cultures for 1 h with 4% paraformaldehyde in 0.1 M phosphate buffer (PB), pH 7.3, at 4 °C.
2. Remove the fixative and gently rinse the cultures 3-4 times (10-15 min each) in 0.1 M PB, pH 7.3.
3. Detach the hydrogel sandwich from the bottom of the well with spatula or forceps. Transfer the collagen block with a fine paintbrush to a 6-well culture plate containing 0.1 M PBS with 0.5% non-ionic detergent.
4. Incubate free-floating hydrogels in the blocking solution (10% serum, 0.5% non-ionic surfactant, and 0.2% gelatin in 0.1 M PBS) for 2-3 h at RT with gentle agitation.
5. Rinse 3 times (10-15 min each) with 0.1 M PBS containing 0.5% non-ionic surfactant.
6. Incubate with primary antibody diluted in PBS containing 5% serum, 0.5% non-ionic surfactant, 0.2% gelatin, and 0.02% sodium azide. Incubate with the primary antibody for 36-48 h at 4 °C on a shaker.
   NOTE: Usually an antibody against class III β-tubulin (α-TUJ-1) (diluted 1:2,000) is used to define axonal growth in 3-D hydrogel cultures.
7. After incubation, rinse as in step 5.5.
8. Incubate with secondary antibody for 4 h at RT (or 6-7 h at 4 °C) on a shaker diluted in 5% serum, 0.5% non-ionic surfactant, and 0.2% gelatin. A horse anti-mouse biotinylated antibody (diluted 1:200) is used in this experiment.
9. Rinse cultures as in 5.5.
10. Incubate the cultures for 2 days at 4 °C with avidin-biotin complex (ABC) solution 1:100 diluted in PBS containing 5% serum, 0.5% non-ionic surfactant, and 0.2% gelatin. Alternatively, use horseradish peroxidase (HRP)-tagged streptavidin (diluted 1:300-400) in the same buffer.
11. Rinse cultures as described in 5.5.
12. Rinse the cultures several times with 0.1 M Tris-HCl buffer, pH 7.6 for 1 h.
13. Incubate the cultures with 0.03% of 3,3′-Diaminobenzidine tetrahydrochloride (DAB) solution in 0.1 M Tris-HCl, pH 7.6.
14. Add 5-8 µL of 1% H₂O₂ and wait 10-15 min. Monitor the development of DAB under a microscope using a 4-10x objective.
15. Stop the reaction by removing DAB solution with 0.1 M Tris-HCl buffer, pH 7.6.
16. Rinse the cultures in PBS for 30 min (several changes).
17. Mount the hydrogels onto glass slides using aqueous-based mounting media.
18. Analyze the length and distribution of the axons inside the hydrogel using Sholl analysis plug-in or with NeuriteJ plug-in for ImageJ software³⁰.