- 2x curved forceps
- 1x 10 cm dish
- 1x razor blade
- 4x 1 mL micropipette tips -- 2 mL serological pipettes (or wide bore pipette tips)
A. 1x No. 10 scalpel
B. 2x Dumont No. 5 forceps
C. 1x 22.5° micro knife
D. 1x small flat spatula
E. 2x sterile syringes
F. 2x 27G ½ inch needles
G. 3x 6 cm dish
H. 7x 15 mL conical tubes
I. 1 x microscissors*
*microscissors are required for isolation of mouse embryonic hippocampus
Media and Solutions
1. Coating of Tissue Culture Dishes
Prepare one day prior to isolation under sterile conditions.
- Start with Poly-D-Lysine (BD cat no. 354210) 20 mg vial; reconstitute with 4 mL UltraPure Distilled water to 5mg/ml stock solution. Divide to 1 mL aliquots and store at -20°C.
- Dilute stock to working solution of 0.05 mg/ml. Add 1 mL of working solution to 35 mm dishes and incubate for 2 hours.
- Wash 3x with UltraPure Distilled water. Let air dry for ~4 hours.
- Parafilm wrap and store at 4°C for up to 2 weeks.
2. Preparing Dissection Solution
Adapted from: “Preparation of Dissociated Mouse Cortical Neuron Cultures.” Lutz G. W. Hilgenberg, Martin A. Smith. J Vis Exp. 2007; (10): 562. Published online 2007 December 19, doi: 10.3791/562. PMCID: PMC2557074.
- Make Solution A by adding the following materials to 100 mL UltraPure Distilled water. Bring final volume to 500 mL. Filter and store at 4°C.
Raw Material Amount Final Concentration Sodium Chloride (NaCl) 80.0 g 137 mM Potassium Chloride (KCl) 4.0 g 5.4 mM Sodium Phosphate Dibasic Anhydrous (Na2HPO4) 0.24 g 0.17 mM Potassium Phosphate Monobasic Anhydrous (KH2PO4) 0.3 g 0.22 mM
- Make solution B by diluting 2.5ml of 1M HEPES Buffer Solution in 247.5ml of UltraPure Distilled water to a final working concentration of 9.9mM. Filter and store at 4°C.
- Complete Dissection Solution (DS) by adding the following materials to 400 mL UltraPure Distilled water. Adjust pH to 7.4 with NaOH. Bring final volume to 500 mL. Filter and store at 4°C.
|Raw Material||Amount||Final Concentration|
|Solution A||25 mL||--|
|Solution B||4.0 g14 mL||--|
|D-Glucose||3.0 g||33.3 mM|
|Sucrose||7.5 g||43.8 mM|
3. Preparing Complete Neurobasal Media
Using the Gibco B-27 Plus Neuronal Culture System add 10 mL of B-27 Plus Serum-Free Supplement (50X)one 500 mL bottle of Neurobasal Plus Media.
4. Preparing Trypsin Inhibitor / BSA Stock Solutions
Dissolve 1 g BSA (SIGMA, A-7030) and 1 g Trypsin Inhibitor (Worthington, LS003087) in 20 mL prepared Dissection Solution. Adjust pH to 7.4 with 1 N NaOH. Sterile filter through 0.2 μm syringe filter. Divide into 800 μL aliquots and store at -20°C.
1. Sacrifice an appropriately timed pregnant female (16.5 days) according to your facilities’ IACUC regulations. Open abdominal cavity and remove uterus, containing embryos. Place in 10 cm dish containing cold dissection solution.
2. Carefully remove placenta using two small curved forceps with gentle opposite pulling motions. Then remove embryo from remaining sac and place in fresh cold dissection solution. Note: If isolating tissue from multiple embryos, proceed to isolate all required embryos from the uterus and the embryonic sac and keep submerged in cold dissection solution on ice.
3. Carefully decapitate animal with clean sterile scalpel and place in 6 cm dish containing 4-5 mL dissection solution.
4. Under dissecting microscope, use two No.5 Dumont forceps and carefully remove thin skin layer by pinching skin at center and peeling away. To cut and remove forming skull pieces, anchor head by piercing orbital cavities with forceps and use a 22.5° angled micro knife to make a very shallow incision from anterior to posterior.
5. Without removing the anchoring forceps, carefully place the second set of forceps at an angle to grab the skull flap without piercing the tissue. Pull gently to remove soft bone fragments. Try to avoid pulling off small pieces with jerky movements.
6. Using small flat spatula, angled beneath the brain, lift carefully, remove brain from skull and place in a clean 6 cm dish with fresh dissection solution.
7. Separate two hemispheres from the midbrain. Using one sterile syringe with needle attached, anchor tissue in the midbrain and use other needle to gently separate the two hemispheres along the middle of the sagittal plane. Then use the needles to separate the midbrain from each of the two hemispheres.
8. With the outer surface of the hemisphere facing upwards, remove meninges. Use one needle to anchor tissue and forceps to carefully peel away meninges from the outer surface. Without releasing meninges from forceps, turn tissue over, and carefully peel away remaining meninges. If done properly, entire meninges can be removed in one piece.
9. Proceed to isolate cortex from remaining inner midbrain region for each hemisphere. Using one needle as an anchor, carefully remove the midbrain from the inner side of the cortex with short cuts using the other needle. If isolating hippocampus,anchor tissue and use microdissection scissors to isolate from cortices.
10. Once desired regions have been isolated, the cortices need to be cut into approximately 5-6 smaller pieces for digestion. To cut cortices, use an x-position with the needles, where one needle serves as an anchor while the other needle is used to cut tissue. The hippocampus can remain whole. Note: If isolating tissue from multiple brains, isolated tissue can be stored in dissection solution on ice for up to 30 minutes.
11. Using syringe with no needle 11 attachment, or wide bore micropipette tip, aspirate tissue pieces and place in clean dish. Whichever method used, take special care as to not disrupt or tear tissue pieces.
12. Add 5 mL sterile 10X TrypLE Select and place dish in 37°C incubator for 25-30 minutes.
13. During incubation time prepare wash solutions and media if have not done so already. Keep wash solutions on ice. Ensure complete media is pre-warmed to 37°C prior to use.
|Solutions||High Wash (Hi)||Low Wash (Li)|
|Trypsin Inhibitor/ BSA Stock||600 μL||160 μL|
|Dissection Solution||2.4 mL||7.84 mL|
|Aliquot Volume||1.5 mL||2.6 mL|
|Aliquot # Per Wash||2||3|
14. Note: All subsequent steps are performed in a laminar flow hood using either a 2ml serological pipette or wide bore pipette tip for aspiration. Remove dish containing tissue pieces from 37°C and aspirate tissue pieces with as little TrypLE solution as possible. Place in 15 mL conical tube containing 10 mL cold dissection solution and let tissue settle for 2 minutes.
15. Aspirate tissue pieces with as little solution as possible, and transfer to 1st High Trypsin inhibitor wash (Hi), swirl and let tissue settle for 2 minutes. Repeat for 2nd Hi wash.
16. Aspirate tissue pieces with as little solution 16 as possible, and transfer to 1st Low Trypsin inhibitor wash (Li), swirl and let tissue settle for 2 minutes. Repeat for 2nd and 3rd Li washes.
17. After the last Li wash, aspirate tissue pieces and place in 6 cm dish containing 5 mL prewarmed complete media.
18. With a clean razor blade cut the tips off of three 1 mL micropipette tips to give 3 different bore sizes of tips for trituration.
19. Starting with the widest bore size tip and subsequently moving to the next smaller size, slowly triturate tissue pieces by pipetting up and down 10-20 times. Switch to uncut tip for the final trituration. All trituration should take less than 5 minutes.
20. Transfer media containing cells to clean 15 mL conical. Rinse dish with 5 mL additional complete media and add to conical tube for final volume of 10 mL.
21. Allow larger tissue pieces to settle for approximately 2 minutes and then transfer 9.5 mL of cell suspension, taking care not to disturb settled tissue at bottom of tube, to a new, clean 15 mL conical.
22. Perform Trypan Blue exclusion assay to count cells and assess viability. Plate at 200K-300K per plate for best transfection results.
- 1X D-PBS containing Ca2+ and Mg2+: (14040-141)
- Goat Serum: (16210-064)
- Mouse Anti-MAP2 antibody: (13-1500)
- Rabbit Anti-GFAP antibody
- Alexa Fluor 488 goat anti-mouse: (A-11029)
- Alexa Fluor 594 goat anti-rabbit: (A-11037)
- 4’, 6-diamidino-2-phenylindole, dihydrochloride (DAPI): (D1306)
- 1% Triton X-100: (HFH10)
- ProLong Gold Antifade Reagent: (P36930)
- EM Grade Paraformaldehyde (PFA): (Electron Microscopy Services)
1. Preparing 20% PFA Stock Solution
- Add 1X PBS to 20g of PFA, and bring volume up to 100 mL.
- Add 0.25 mL of 10 N NaOH and heat the solution at 60°C using a magnetic stirrer until the solution is completely dissolved.
- Filter the solution through a 0.2 μm filter and cool on ice. Ensure the pH remains around 7.5-8.0.
- Aliquot 2 mL into 15 mL tubes, freeze on dry ice, and store at –20°C.
2. Preparing 4% PFA For Fixing
- Remove culture medium and gently rinse cells twice with D-PBS without dislodging them.
- Fix cells with freshly prepared 4% PFA at room temperature for 15 minutes.
- Rinse cells three times with D-PBS without dislodging them.
- Permeabilize the cells with 0.3% Triton X-100 (diluted in D-PBS) for 5 minutes at room temperature.
- Rinse cells three times with D-PBS.
- Incubate cells in 5% goat serum diluted in D-PBS for 60 minutes at room temperature.
- Remove 5% goat serum solution and incubate the cells overnight at 4°C with the primary antibody (Mouse anti-MAP2 at 10 μg/mL and/or Rabbit anti-GFAP) diluted in 5% goat serum.
- Wash cells three times for 5 minutes with D-PBS (if using a slide, use a staining dish with a magnetic stirrer).
- Incubate cells for 60 minutes at room temperature with fluorescence-labeled secondary antibody (Alexa Fluor 488 goat anti-mouse (H+L) at 10 μg/mL and/or Alexa Fluor 594 goat anti-rabbit (H+L) at 10 μg/mL) diluted in 5% goat serum.
- Wash cells three times with D-PBS. In final wash, counter-stain cells for 10 minutes with DAPI solution (3 ng/mL).
- Rinse cells with D-PBS, and if desired, mount using 3 drops of ProLong Gold antifade reagent per slide and seal with cover slip. Slides may be stored away from direct light at 4°C.
- Observe cells under microscope using filters for FITC, Cy5, and DAPI.
Transfect cells according to the following diagram. Volumes are given on a per-plate basis for a 35 mm glass-inserted plate (MatTek, P35G-0-14-C).
*mRNA can be generated using Ambion mMessage mMachine T7 Ultra Kit (AM1345)
LT176 rev. 06-21-2013