Isolation and Culture of Dissociated Sensory Neurons From Chick Embryos

Summary

Cell culture models provide detailed control over environmental conditions and thus provide a powerful platform to elucidate numerous aspects of neuronal cell biology. We describe a rapid, inexpensive, and reliable method to isolate, dissociate, and culture sensory neurons from chick embryos. Details of substrata preparation and immunocytochemistry are also provided.

Abstract

Neurons are multifaceted cells that carry information essential for a variety of functions including sensation, motor movement, learning, and memory. Studying neurons in vivo can be challenging due to their complexity, their varied and dynamic environments, and technical limitations. For these reasons, studying neurons in vitro can prove beneficial to unravel the complex mysteries of neurons. The well-defined nature of cell culture models provides detailed control over environmental conditions and variables. Here we describe how to isolate, dissociate, and culture primary neurons from chick embryos. This technique is rapid, inexpensive, and generates robustly growing sensory neurons. The procedure consistently produces cultures that are highly enriched for neurons and has very few non-neuronal cells (less than 5%). Primary neurons do not adhere well to untreated glass or tissue culture plastic, therefore detailed procedures to create two distinct, well-defined laminin-containing substrata for neuronal plating are described. Cultured neurons are highly amenable to multiple cellular and molecular techniques, including co-immunoprecipitation, live cell imagining, RNAi, and immunocytochemistry. Procedures for double immunocytochemistry on these cultured neurons have been optimized and described here.

Introduction

Neurons are complex cells that carry information essential for a variety of functions including sensation, vision, motor movement, learning, and memory. Unique from other cell types, neurons extend arm-like processes, called axons, to form essential neural highways for communication. During development specialized compartments located at the tips of growing axons, called growth cones, navigate through a concert of extracellular cues to lead the axon to its appropriate destination. The intricate molecular mechanisms that underlie growth cone navigation are not fully understood. To better understand these mechanisms, investigators have used cell culture models to study neurons in a defined and simplified in vitro environment. Studying neurons in culture¹ has led to significant advances in our understanding of neuronal cell biology including: neuronal differentiation², cytoskeletal dynamics, endocytosis and trafficking, dendrite regulation^3,4, axonal regeneration⁵, and clinical conditions such as neuropathies⁶. In addition, cultured neurons are highly amenable to a wide range of research techniques including immunocytochemistry, cell surface co-immunoprecipitation, Western blot, transfection, RNAi, and live imaging such as timelapse analysis of growth cone motility. Thus, culturing primary neurons is a powerful approach to elucidate numerous aspects of the cell biology of neurons.

The cell culture model provides investigators with detailed control over environmental conditions and variables. For example, the substrata on which neurons are plated (and grow upon) can be easily manipulated. Here, we provide detailed instructions for generating two distinct substrata, one with a low laminin-1 concentration and the other with saturating concentrations of laminin-1. Surprisingly, different concentrations of the same molecule can have dramatic effects on the internal state of neurons as well as their cell surface composition. For example, intracellular levels of cAMP and surface levels of integrins are significantly different in neurons plated on these two substrata^7,8. Additional studies have shown that other molecules, including fibronectin and chondroitin sulfate proteoglycans, impact the expression of cell surface molecules and neuronal motility^7-11. In addition, soluble molecules such as neurotrophins and neurotropins also impact cell membrane composition and neuronal motility^12-16 and can be easily and accurately manipulated in a cell culture model.

Here, we describe methods to isolate and culture dissociated sensory neurons from chick embryos. This procedure has been used to make significant breakthroughs in neurobiology, including axon outgrowth^{5,7,8,10,11,16-21} and was modified from a procedure designed to isolate ganglion cells²². There are several advantages to this approach. First, many features of chick dorsal root ganglion (DRG) development are well characterized including the time frame for birth, axon extension, and protein expression profiles^2,23-28, thus providing an instructive basis upon which to build informative in vitro experiments. Second, dissociated neuronal cultures allow the investigator to more directly study neurons compared to alternative approaches using intact DRG explants (which contain neurons and non-neuronal cells) and/or mixed cultures containing both dissociated neurons and non-neuronal cells. Third, the procedure described here is straightforward, inexpensive and amenable to undergraduates. Therefore, this technique can be used for research as well as for teaching purposes. Furthermore, minor variations of this protocol should allow fast, high yield purification of neurons from sources other than DRGs. For example, this procedure could be modified to provide neuronally enriched cultures from other tissues such as embryonic forebrain or spinal cord.

Immunocytochemistry protocols have been optimized for these dissociated neuronal cultures and are described in detail here. The procedure for double immunocytochemistry against neural cell adhesion molecule (NCAM) and β1 integrins is provided. Data generated from these immunocytochemical methods have been used to examine the spatial patterning and intensity of several molecules in cultured neurons^8,16.

Protocol

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.

Results

The protocol described here enables investigators to culture an enriched population of dissociated embryonic sensory neurons with very few (e.g., <5%) non-neuronal cells^7,8,10,16. Numerous DRGs can be obtained from the lumbosacral, thoracic and cervical regions. Depending upon the needs of the investigator, DRGs from these distinct anatomical regions can be easily isolated. For example, Figure 1 shows images of the chick embryo through various stages of dissection with the lumbar D...

Discussion

Here we present detailed protocols for isolating and culturing dissociated sensory neurons from a chick embryo. This procedure generates an enriched population of robustly growing neurons in vitro^7,8,10,16. Numerous cellular and molecular techniques can be applied to these cultured neurons, including immunocytochemistry, which is described here. This protocol was recently used to quantitatively assess the intensity of immunolabeled activated integrins in sensory growth cones^8,16. In these p...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Sterile small culture dishes (35 mm)	Corning	430165
Sterile large culture dishes (100 mm)	Falcon	353003
Sterile large Petri dishes (100 mm)	VWR	89000-302
Glass Petri dish (100 mm)	VWR	D108962
Fine forceps	Fine Science Tools	11251-10
Standard forceps	Fine Science Tools	1100-12
Coverslips (22 x 22 mm)	Fisher	12 518 105K	0.13-0.17 mm thickness is optimal for microscopy
Porcelin coverslip holder	Thomas scientific	8542E40
Fetal Bovine serum	Gibco	17502-048	use 50 ml aliquots per 500 ml of Ham's F12 media
HCl	VWR	VW3204-1	use at 2 M, can be used twice before discarding
NaCl	Sigma	S9888	use 5 M NaCl solution for laminin-1 stock solution
Laminin-1	Invitrogen	23017-015	Add 72 µl of sterile 5 M NaCl and aliquot into 50 µl
15 ml Conical tube	cell treat	229411
PBS CMF 10x	Gibco	14200	used at 1x, dilute with sterile Millipore filtered water
PBS 10x	Gibco	14080	used at 1x, dilute with sterile Millipore filtered water
Ham's F12	Lonza	12-615F
Fetal Bovine Serum Albumin (BSA)	Gibco	10438	use in F12HS20 at 10%
HEPES	Sigma	H3375	make sterile 1 M solution in distilled water, dilute in F12 media to obtain final concentration of 10 mM
Penicillin/streptomyocin	Sigma	P0781	use 5 ml in 500 ml of F12 media
NT3	Millipore	GF031	Add 1 ml of sterile Millipore water, aliquot under sterile conditions, store at -20 °C, use at final concentration of 10 ng/ml in F12H media, keep in -20 °C non-defrosting freezer
NGF	RnD systems	256-GF	Add 1 ml of sterile 1x PBS with 0.1% BSA, aliquot under sterile conditions, store at -20 °C, used at final concentration of 10 ng/ml in F12H media
L-Glutamine	Sigma	G7513	aliquot under sterile conditions, store at -20 °C, use at final concentration of 200 mM in F12H media
Trypsin	Sigma	T4049	aliquot under sterile conditions, store at -20 °C
Bovine Serum Albumin (BSA)	Gibco	15260
Other items needed: general dissection instruments, including glass Pasteur pipettes, fertile white leghorn chicken eggs, check egg incubator (humidified, 37 °C), cell incubation chamber (humidified, 37 °C), laminar flow hood, binocular stereovision dissecting scope
Immunocytochemistry Reagents Table
Sucrose	Sigma	S9378	used in fixative solution (4% paraformaldehye, 30% sucrose, 2x PBS)
Triton-X 100	Sigma	T-8787
Paraformaldehyde	Sigma	P6148	used in fixative solution (4% paraformaldehye, 30% sucrose, 2x PBS)
Fluoromount G	Southern Biotech	0100-01
Normal goat serum	Life technologies	PCN500
Microscope slides	VWR	16004-430
Primary Antibody Table
Antibody against NCAM	Millipore	AB5032	polyclonal
Antibody against activated β1 ingegrin	Millipore	MAB19294	monoclonal
Secondary Antibody Table
Goat anti-mouse IgG Alexa 488	Life technologies	A11001
Goat anti-rabbit IgG Alexa 548	Life technologies	A11036