Isolation and Culture of Dissociated Sensory Neurons From Chick Embryos

1. Coverslip Preparation: Acid Wash and Bake

1. At least 2 days prior to dissection (step 2), begin the following steps to prepare coverslips. Perform the acid wash step to remove oils and thoroughly clean coverslips.
2. Load coverslips in porcelain holder that vertically positions coverslips and prevents them from touching each other. Submerge holder and coverslips into glass histology container filled with 2 M hydrochloric acid (HCl). Alternatively, if porcelain holder is not available, spread ~20 coverslips horizontally to cover the bottom of 100 mm glass Petri dish and submerge coverslips in 2 M HCl.
3. Place glass vessel (with coverslips submerged in acid) on rocker table in fume hood and allow gentle rocking for at least 5 hr at room temperature. Alternatively, acid wash coverslips overnight under the same conditions.
4. Wash coverslips with distilled H₂O(dH₂O) by transferring porcelain holder loaded with coverslips from 2 M HCl to a clean glass histology container filled with dH₂O. Alternatively, decant 2 M HCl from glass Petri dish with coverslips and refill with dH₂O.
5. Wash by decanting dH₂O again and refilling glass vessel with fresh dH₂O. Repeat for a total of 3 quick washes.
6. Begin longer washing regimen by returning glass vessel (with coverslips and dH₂O) onto rocker. Allow for gentle agitation for 10-30 min at room temperature. Then, decant dH₂O and add fresh dH₂O. Repeat for a total of at least ten washes. Alternatively, continue washing overnight.
7. Place washed coverslips into furnace preheated to 350 °C. If using porcelain holder, remove holder from dH₂O-filled glass container and then place holder with coverslips directly into the furnace. If using a glass Petri dish, decant dH₂O and place the glass dish with coverslips into the furnace.
8. Bake coverslips for at least five hours or overnight at 350 °C.
9. After baking is complete, remove coverslips from porcelain holder with fine forceps and place into sterile 100 mm glass Petri dish. Leave coverslips that were not originally placed in porcelain holder in the glass Petri dish.

2. Chick Dissection and Isolation of DRGs

1. Remove fertile, staged White Leghorn chick egg from incubator (held at 37-38.5 °C with gentle rocking).
   NOTE: Previous studies have used embryonic day 7-10 for isolation of DRGs^2,8,11-13,16.
2. Place selected egg in laminar flow hood. Conduct all procedures listed below under aseptic conditions in the laminar flow hood with sterile solutions and instruments. Pre-warm all solutions in a 37 °C water bath.
3. Crack egg and place contents into a plastic 100 mm Petri dish. Use standard forceps to transfer embryo to another 100 mm Petri dish.
4. Using a Pasteur pipette, add warmed 1x PBS to Petri dish to keep tissue wet. Decapitate embryo by pinching neck with fine forceps. Transfer body to another clean 100 mm Petri dish.
5. Add warm F12HS10 (Ham’s F12, 10 mM HEPES, 100 units/ml penicillin, 0.1 mg/ml streptomycin, 10% fetal bovine serum) to dish with Pasteur pipette to keep tissue wet. Use forceps to position embryo on its dorsal side (Figure 1A).
6. Place 100 mm plastic Petri dish containing embryonic tissue onto a binocular stereovision dissecting microscope in laminar flow hood. Use fine forceps to make a vertical midline incision from tail to neck by pinching the developing dermis to expose the heart, lung and other organs. Alternatively, make a series of small tears to expose internal organs.
7. Gently use 2 sets of sterile fine forceps to: a) grasp the neck and b) grasp the aorta or other tissue immediately superior to the heart and pull towards the tail, eviscerating the animal.
8. Apply warm F12HS10 with Pasteur pipette to tissue. Use forceps to remove any remaining cardiovascular, respiratory or digestive organs to expose the developing ribs, vertebral column and DRGs (Figures 1B-1D).
9. Under the dissecting microscope, use fine forceps to further remove organs and collect DRGs along both sides of lumbar vertebral column, inferior to the ribs. Collect DRGs by plucking intact DRGs from the embryo. Pinch and sever the roots that reach from the DRG towards the spinal cord and grasping the developing nerve that reaches from the DRG towards the periphery or vice versa.
10. Place intact DRGs into a 35 mm culture dish filled with warm F12HS10. Remove any excess tissue, such as the dorsal or ventral roots that may have adhered to DRGs, from the intact DRGs in 35 mm dish by pinching with fine forceps.
11. To obtain DRGs superior to the lumbar region, remove the ventral half of the thoracic and cervical vertebral column. This is done by making a cut in the developing vertebral column perpendicular to the spinal cord axis in the upper cervical region and another cut in the upper lumbar region. Then, make cuts parallel to the spinal cord axis along the right and left sides of the vertebral column. Lift the ventral half of the vertebral column from the embryo, which exposes the exposes the spinal cord (Figures 1E-1F).
12. To allow easy access to DRGs, remove the thoracic and cervical spinal cord by grasping the spinal cord with fine forceps. Use the forceps to lift the spinal cord out of the vertebral column. As an approximation, the thoracic spinal cord is at the level of ribs and the cervical area is superior to the ribs.
13. Collect intact thoracic and cervical DRGs with fine forceps and place them in warm F12HS10 with other DRGs. Remove excess tissue that may adhere to DRGs.
14. Rinse interior of a new glass Pasteur pipette with F12HS10 to help prevent DRGs from sticking to walls of pipette. Use rinsed pipette to transfer all DRGs and F12HS10 from small Petri dish to a sterile 15 ml conical tube and immediately proceed to step 3.
    NOTE: Additional embryos can be dissected to increase number of DRGs. DRGs should be kept in F12HS10 at 37 °C until ready for step 3.

3. Dissociation, Enriching, and Culturing DRG Neurons – Part 1

1. Place conical tube with intact DRGs into centrifuge and gently spin (200 x g) for 2-3 min. Confirm that DRGs have sunk to bottom of tube. If not, spin again.
2. Using a pipette, gently remove F12HS10, leaving DRG pellet undisturbed. Add 2 ml of 1x calcium and magnesium free (CMF) PBS, dislodging the pellet of DRGs. Spin again and remove CMF PBS while leaving the DRG pellet intact. Repeat this wash step for a total of three washes to remove fetal bovine serum in F12HS10, which would interfere with trypsin digestion in next step.
3. Add 2 ml of warmed trypsin to 15 ml tube containing 2 ml CMF PBS and DRGs. Place tube in 37 °C water bath for 10-15 min to digest DRGs into dissociated cells.
4. During incubation, make F12H + supplement containing media (Ham’s F12, 10 mM HEPES 100 units/ml penicillin, 0.1 mg/ml streptomycin, 10 ng/ml NT3, 10 ng/ml NGF, 200 mM l-glutamine and N2) and warm in 37 °C water bath. Typically, 10 ml of total media is sufficient for 4-5 35 mm dishes of primary sensory neurons. Also begin thawing laminin-1 for coating coverslips (step 4).
5. Gently triturate DRGs in trypsin solution with p1000 pipette and visually inspect for absence of intact DRGs. Continue trituration until intact DRGs are no longer seen by eye and/or allow for longer trypsin incubation in 37 °C water bath until intact DRGs are dissociated. To protect DRG neurons from damage, avoid air bubbles during trituration and limit trypsin digestion to <30 min.
6. Centrifuge tube containing dissociated DRGs and trypsin at 200 x g for 3-5 min to form pellet. Spin longer if needed until pellet is formed.
7. Carefully aspirate trypsin without disrupting pellet. Add 2 ml of F12HS10 to pellet. Gently triturate DRGs with a p1000 pipette tip.
8. Transfer all contents from 15 ml conical tube into 100 mm culture dish. Rinse the tube with 8 ml F12HS10 and transfer it to the culture dish, 2 ml at a time for a total of 10 ml. This rinsing step helps collect cells that may have remained adhered to the walls of the tube.
9. Place 100 mm culture dish (with 10 ml F12HS10 and dissociated DRG cells) into a humidified 37 °C incubator for 3 hr.
   NOTE: CO₂ is not required in the incubator because HEPES buffers media pH in the absence of CO₂. During incubation, glial cells bind to surface of tissue culture plastic and neurons tend to loosely adhere to glial bed.

4. Coating Acid Washed and Baked Coverslips with Laminin-1

1. Thaw sterile aliquots of frozen laminin-1 on ice.
2. Under sterile conditions, add 1 ml of sterile 1x PBS to 50 μl aliquot of laminin-1.
3. Use a spectrophotometer to obtain an absorbance value of laminin-1 at 280 nm. Use this value in the following equation to determine the concentration of laminin-1.

ABS ₂₈₀ = (concentration mg/ml) * (extinction coefficient of protein)
The extinction coefficient for laminin-1 is 0.86.

4. Under sterile conditions, dilute laminin-1 with 1X PBS to obtain the concentrations 1 mg/ml and 20 mg/ml. Use these two applied concentrations of laminin-1 to achieve a ten-fold difference of laminin-1 bound to the coverslip (30 ng/cm² and 300 ng/cm², respectively) after incubation^7,8,10.
5. In a laminar flow hood, use fine forceps to place one acid-washed and baked coverslip into the bottom of one 35 mm culture dish. Repeat as necessary to obtain the number of dishes needed.
6. Subject the coverslips to UV light for 20-30 min to further ensure sterilization. Keep dish lid off during this step.
7. Add 400 μl of prepared laminin-1 to each glass coverslip. Use pipette tip to spread laminin-1 to ensure full coverage of coverslip. Surface tension will allow laminin-1 to remain on coverslip and not spread onto bottom of culture dish, which is required.
8. Place lid onto dish and allow laminin-1 to incubate for 2-3 hr at room temperature (3 hr is optimal).
9. After laminin-1 incubation, rinse coverslips using aseptic technique with sterile PBS in laminar flow hood. Remove solution on coverslip and add 400 μl of PBS. Wait 5-10 min. Repeat for a total of 3 rinses.
10. Leave PBS on coverslip after last rinse. Do not break surface tension during rinsing and do not allow coverslips to dry out. Leave lid on until ready to plate dissociated neurons onto the coverlip.

5. Dissociation, Enriching, and Culturing DRG Neurons – Part 2

1. After incubation, use a 10 ml sterile pipette to gently rinse the bottom of the 100 mm dish containing dissociated DRG cells. Hold dish at ~45° angle, pull ~7 ml of F12HS10 with pipette aid, then gently expel 7 ml onto approximately 1/3 of dish. Repeat 5x, gently dislodging neurons.
2. Rotate dish and repeat rinsing procedure, for another six rinses, on another 1/3 of the dish. Repeat procedure again for remaining 1/3 of dish. In this way, gently rinse the entire bottom of the dish with F12HS10 to remove neurons.
3. Using same pipette, transfer F12HS10 containing neurons from 100 mm dish to 15 conical tube. Centrifuge for 5-10 min at 200 g to pellet cells.
4. Remove F12HS10, leaving pellet undisturbed. Resuspend pellet with 2 ml of warmed F12H + supplements.
5. Use a hemocytometer to determine cell density.
6. Dilute cells as necessary with F12H + supplements to obtain desired plating concentration (120,000 cells/ml for coverslips coated with 1 μg/ml laminin-1 and 40,000 cells/ml for coverslips coated with 20 μg/ml laminin-1^8,16).
7. Remove PBS from laminin-coated coverslips and immediately place 400 μl of cells onto coverslips. Carefully transfer dish to a humidified, 37 °C cell culture incubator. Allow to incubate for at least 1 hr and up to 3 hr to allow neurons to adhere to laminin-1.
8. Under sterile conditions, gently flood 35 mm dishes containing neurons with 1.6 ml of F12H + supplements.
   NOTE: At this time, surface tension on coverslip can be broken. Neurons have adequately adhered to laminin-1 on coverslip.
9. Return cells to incubator and incubate overnight. Perform experiments such as immunocytochemistry the following day.

6. Immunocytochemistry

1. Prior to immunocytochemistry, warm fixative solution (4% paraformaldehyde, 30% sucrose 2X PBS) in 37 °C water bath.
   NOTE: Before fixing cells, treat with various reagents, such as netrin-1, Mn²⁺, sonic hedgehog, semaphorin, and ephrin^16,29-31.
2. Slowly remove 1 ml of the 2 ml of F12H + supplements from culture dish. Gently add 1 ml of warmed fixative solution. Allow to incubate at room temperature for 10-15 min.
3. Carefully remove fixative solution and add 2 ml PBS. Apply solution towards edge of dish, avoiding possible dislodging of fixed cells due to application of PBS. Incubate at room temperature for 5 min.
4. Repeat PBS wash for a total of 5 washes. Do not allow cells to dry during any step of immunocytochemistry.
5. Remove PBS. Apply 1 ml of blocking solution (0.1% Triton-X, 1X PBS, 5% normal goat serum). Incubate for 1 hr at room temperature. Alternatively, to stain surface molecules, do not use Triton-X in blocking solution⁸.
6. Dilute primary antibodies into blocking solution per manufacture’s recommendation. Use 1:500 for antibodies against NCAM and activated β1 integrins¹⁶. Remove blocking solution, add 1 ml of primary antibody solution and incubate for 1 hr at room temperature or overnight at 4 °C.
7. Remove primary antibody solution and add 2 ml of 1X PBS. Incubate for 5-10 min at room temperature. Repeat for a total of 4 PBS washes. Washes can be extended, if needed.
8. Dilute secondary antibody 1:1,000 in blocking solution. Remove PBS, add 1 ml secondary antibody solution and incubate for 1 hr at room temperature.
   NOTE: Secondary antibodies are light sensitive. Keep solutions and cells in the dark as much as possible from this point forward.
9. Remove secondary antibody solution and rinse 5x with 1x PBS, as done previously.
10. Apply ~60-80 μl of Fluoromount G onto a microscope slide. Using fine forceps, gently lift coverslip out of dish and lower coverslip into Fluoromount G onto microscope slide. Avoid air bubbles.
11. Store slides horizontally at 4 °C in the dark. After 12-24 hr, Fluormount G will polymerize and coverslip will be firmly attached to microscope slide. Image cells after this polymerization step is complete.