A Simplified Method for Ultra-Low Density, Long-Term Primary Hippocampal Neuron Culture

All experimental procedures involving mice were approved by the Institutional Animal Care and Use Committee of the University of Arizona, and conformed to NIH guidelines.

1. Tissue Source for Hippocampal Neuron Culture

1. To generate prenatal mouse pups for hippocampal neuron culture, use time-pregnant mice (C57Bl6/J) that are bred in house¹⁷. The day with vaginal plug detection is designated as E0.5. The planned harvest time for culture is E16.5-E17.5.
   Note: This protocol describes cultures of two 24-well plates of two high density neurons co-culturing with low-density neurons (48 coverslips total). For more plates, materials and reagents can be scaled correspondingly.

2. 24-well Plates Preparation for High Density Neuron Cultures

Note: Perform the following steps (2-3) the day before the planned harvest of embryos.

1. Bring two 24 well plates into the cell culture hood, open the individual packaging.
2. Connect an 18 G syringe needle to a 1 ml plastic single-use syringe.
3. Hold the syringe, and aim the needle tip at ~1 mm left to the center of the bottom of the well. Apply moderate force (~250 g) to etch the bottom of the well (~1 mm long). A groove will be formed and the plastic materials displaced will form an elevated support above the flat bottom well surface. Etch another ~1 mm-long parallel groove at ~1 mm to the right of the center of the bottom in a similar way.
4. Repeat this step for all the wells of the two 24-well plates. The two 24-well plates used for high density culture are now ready for coating with poly-D-lysine (see step 3.8 below).

3. Coverslips Preparation for Low Density Neuron Cultures

1. Use 12-mm diameter #1 round glass.
2. Place 50 coverslips inside a 100 mm diameter sterile plastic petri dish, and submerge coverslips in 20 ml 70% ethanol solution. Place this petri dish on a rotating platform with moderate (70 rpm) speed rotation for an hour. Remove 70% ethanol and then replace with a fresh batch of 70% ethanol. Rotate and wash for another hour.
3. Discard 70% ethanol cleaning solution. Add sterile water (filtered through 0.2 µm filter unit). Rinse the coverslips rigorously by rotating the petri dish by hand. Rinse three times, each time replacing with fresh sterile water.
4. Place the petri dish with coverslips inside a sterile cell culture hood with laminar flow. Aspirate the rinsing water through the vacuum line, and add 10 ml filtered sterile water to immerse the coverslips. The coverslips should be sterile and the surface should be clean enough for poly-D-lysine coating.
5. Open two 24-well plates from their individual packing, and place them in the hood.
6. Turn on the vacuum line in the hood. Connect a 200 µl pipette tip to the vacuum line, and use a scissor to cut the tip to create a larger opening.
7. Use a fine-tip #5 tweezer to pick up coverslips from the petri dish, one at a time, and use the vacuum line to completely dry the coverslip. Then place one coverslip into each of the wells of the 24-well plate. The 50 cleaned coverslips should be enough to fill each of the wells of the two 24-well plates.
8. Dilute the poly-D-lysine stock solution (1 mg/ml) into working solution (0.1 mg/ml) with borate buffer (pH 8.5). For the four 24-well plates (including two high density plates with etched bottom), prepare 30 ml of total working solution.
9. Add 300 µl 0.1 mg/ml poly-D-lysine work solution to each well with glass coverslips. Make sure the coverslips are completely immersed in solution. If not, use the #5 tweezer tip to press down coverslips against the bottom of the well, and use the tip to guide poly-D-lysine solution to cover all the surface of the coverslips. Visually inspect all the coverslips to make sure no air was trapped between the plate and coverslips.
10. Add 300 µl 0.1 mg/ml poly-D-lysine work solution to each well of the high density culture plate with etched bottom. Rotate by hand to spread the solution to cover the entire bottom of the well.
11. Return all the four plates with poly-D-lysine to the culture incubator, and incubate O/N.

4. Washing and Pre-conditioning Plates for Culture

Note: The following steps (4-9) are carried out on the day of tissue harvest.

1. After incubation/coating of coverslips and 24-well plates O/N, bring all four plates (two with donor coverslips, two acceptor high density plates with etched plastic bottom) into the culture hood. Use the vacuum line to remove the poly-D-lysine solution.
2. Immediately following the removal of the poly-D-lysine solution, add ~1 ml sterile water to each well using a plastic transfer pipette. After all the wells are filled with water, let stand for 10 min. Remove water by vacuum, then add 300 µl Wash medium in each well to cover the bottom of the well (with or without glass coverslips). Do not let the poly-D-lysine coating dry out.
3. Return all four plates to the cell incubator. The plates are ready to use once the dispersed hippocampal neurons are prepared.

5. Preparation of Complete Culture Medium, and Trypsin Solution for Enzymatic Digestion

1. Prepare 60 ml Complete Culture medium by mixing the ingredients (see Materials). Use a 50 ml syringe connected with a 0.2 µm-pore size syringe filter, filter the Complete Culture medium into two 50 ml conical tubes. Each tube contains 30 ml Complete Culture medium.
2. In a separate 50 ml conical tube, add ~30 ml Wash medium (see Materials) and warm it up to 37 °C. This will be used to wash the tissue after enzymatic digestion.
3. Prepare the enzyme digestion medium. In a 15 ml conical tube, add 4.5 ml Wash medium and 0.5 ml 2.5% trypsin solution, and 50 µl 100X DNase I.
4. Place the Complete Culture medium, Wash medium, and enzyme digestion medium in a 37 °C water bath.

6. Preparation of Surgical Tools

1. Sterilize all the surgical tools in a sterilization pouch: two small scissors, a pair of forceps and two #5 tweezers.

7. Removal of Brains from E16.5-E17.5 Mouse Embryos, and Dissection of Hippocampi

1. Prepare two 10-cm petri dishes filled with ice-cold, sterile Hank's balanced salt solution (HBSS).
2. Euthanize the mouse dam with an intraperitoneal injection of sodium phenobarbital (100 mg/kg in a volume of ~ 0.5 ml). After injection, once the dam loses response to toe pinch, sacrifice with cervical dislocation.
3. Spray the dam's abdomen area with 70% ethanol, and make a 2-cm long incision along the midline of abdomen to expose the uterus.
4. Remove the embryos and place individual ones in a 10-cm diameter petri dish with cold dissecting HBSS buffer.
5. Hold the embryo by the neck using a forceps, and use a small scissor to make the first cut right in the midline below the cerebellum level and section across the brain tissue. Do not decapitate the embryo.
   1. Insert the right side tip of the small scissor from caudal region that is cut open, all the way through the rostral part of the embryonic brain (do not cross midline), and make a cut on the left side at the level of skull base.
   2. Insert the left side scissor tip again to make a cut on the right side. Leave a narrow band of uncut skull bone and skin tissue at the rostral end, so that the whole brain can be flipped aside for easy extraction.
   3. Use the tip of scissor to scoop out the embryonic brain into the HBSS solution. Inspect the brain under a dissecting microscope. The brain should be intact with no gross cut-off regions.
   4. Repeat these steps to collect all the embryo brains.
6. Collect all the intact brains in a new petri dish containing 20 ml cold HBSS buffer. Place the petri dish under a dissection microscope.
7. View the brain from ventral side. While holding down the brain with forceps at the caudal end, insert a sharp #5 tweezer diagonally to make a cut between the middle rostral point and the caudal-temporal region. Repeat this for the other hemisphere. After cutting, separate two hemispheres with intact hippocampus from the rest of brain stem tissues.
   1. Repeat this for each of the brains.
8. Pool all the dissected hemispheres, and remove the meninges using two pairs of #5 tweezers.
9. Identify the developing hippocampus as a curved structure that is folded inside the temporal lobe. Alternate the two pairs of tweezers to separate the hippocampus from adjoining cortical tissues. Collect and pool all the hippocampi tissue (see Figure 1).

8. Enzymatic Digestion, Separate into Single Neurons

1. Using a plastic transfer pipette, transfer all the dissected hippocampi into the enzymatic digestion solution containing 5 ml Wash medium with 0.25% trypsin + 1X DNase I in a 15 ml conical tube.
2. Place the tube inside a water bath and incubate at 37 °C for 25 min, with intermittent gentle inverting of the tube once in every 5 min.
3. After 25 min, carefully remove the enzyme solution using a hand held plastic pipette by aspiration. Add warm Wash medium to 10 ml and gently wash the tissue by slowly inverting the tube 5 times. Remove the Wash medium, and add 10 ml Wash medium again for an additional wash.
4. Remove the Wash medium, and add 3 ml of fresh warm Wash medium again. Shake the tube by hand rigorously for 15 times to dislodge the digested neurons from the tissue. The medium should become turbid once the cells are separated from the tissue into the Wash medium. Collect the cell suspension in a new 15 ml conical tube while leaving the remaining tissue in the tube.
5. Add another 2 ml Wash medium to the tissue, and gently separate the remaining tissue by triturating through a plastic pipette for ~15 times. Let sit for 5 min. Pool the cell suspension into the new 15 ml conical tube that contains collected 'shake-off' cells.
6. Count the density of neurons with a hemocytometer. Typically, 3,000-5,000 cells per microliter can be obtained depending on the number of embryos dissected. An average number of 2.5 x 10⁷ neurons are estimated if assuming six embryos are dissected.
7. Calculate the volume needed of this cell suspension to yield a density of 250,000 neurons/ml when added to the 30 ml Complete Culture medium. Add this volume to one of the two conical tubes containing 30 ml Complete Culture medium to yield the high density neuron plating medium.
8. Transfer 1.2 ml of this high density neuron solution to another 30 ml Complete Culture medium to yield a concentration of ~10,000 neurons/ml. Use this dilution to seed the low-density coverslips.

9. Plating of Neurons and Long Term Co-culture

1. Bring the two 24-well plates with etched bottoms for high density neurons into the cell culture hood. Remove the 300 µl precondition medium by vacuum. Plate 0.6 ml high density cell suspensions at 250,000 neurons/ml.
2. Bring the other two 24-well plates with coverslips into the culture hood, and remove the 300 µl precondition medium by vacuum. Plate 0.6 ml low density cell suspensions at 10,000 neurons/ml on the coverslips inside the two 24-well plates.
3. Return all four 24-well plates to the incubator. Let sit for 2 hr.
4. Flip the low density coverslips with adhering neurons to the high density culture. Make sure the neurons are facing each other (i.e., not separated by the glass coverslip). Then return the two plates with co-culture to the incubator.

10. Co-culture Sustaining

1. Feed co-cultures by adding 300 µl fresh Feed medium (see Materials) at day in vitro (DIV) 5. There is no need to remove 50% of original Complete Culture medium, as reported by many other studies. The feeding medium also contains 15 µM cytosine arabinoside (Ara-C, to yield a final concentration of 5 µM) to minimize the chance of glia contamination and proliferation.
2. Following this initial feeding, add 300 µl fresh Feed medium without Ara-C to the well every week. Should the culture be kept for more than one month, replace half of the medium with fresh Feed medium every week beyond one month time point (with the first medium half-replacement at DIV33). Morphologically, both high density and low density neurons look healthy up to three months (the longest time observed by the experimenters).

11. Illustration of an Experimental Manipulation-low Density Neuron Transfection

Note: When transfection in low density neurons is desired, a simple calcium phosphate protocol can be adopted. We have found that low density cultures have better transfection efficiency with calcium phosphate protocol before DIV12, but they can be transfected with much lower efficiency at DIV21 or older, during which time dendritic spines are prominent. The feasibility of transfection of the cultured low density neuron is illustrated by transfecting neurons with a pEGFP-C3 plasmid (see below).

1. At DIV21, remove 600 µl conditioned medium from each well of the co-culture, and transfer it to a new 24-well plate. Then add 600 µl fresh Feed medium to the co-culture to bring it back to the original volume.
   1. Transfer the coverslips into the new 24-well plate with 600 µl conditioned medium. Make sure the low density neurons are facing upwards. Place the high density plates and the new plates with coverslips back in the incubator.
2. Prepare two 1.5 ml sterile tubes. In one tube, add 600 µl 2X HEPES buffered saline (HBS) solution. Calculate the volume of 12 µg pEGFP-C3 DNA based on plasmid concentration. In the other tube, add 74 µl CaCl₂ (2 M stock) and the volume of water that after adding plasmid will make 600 µl. Mix well by pipetting and then add plasmid DNA. Slowly mix well. Let both tubes sit for 5 min at RT.
3. While hand-holding the 2X HBS tube and moderately agitating the solution on a mixer, add all of the 600 µl the DNA-CaCl₂ solution drop by drop to the 600 µl 2X HBS tube. It is critical that the precipitate formed is in the shape of 'fine sand' granules. Mixing too strong and too fast is not desirable, since it will more likely produce large 'snow flake'-like precipitates, which has dramatically lower transfection efficiency based on our observations.
4. Let the precipitate continue to form in the dark at RT for 30 min.
5. Add 100 µl precipitate to each well that contains low density neuron on the coverslips, drop by drop and evenly. Return the plate to incubator, and incubate for 2 hr. Assuming most of the CaCl₂ is in free form, this results in a concentration of ~17.6 mM Ca²⁺, which may be toxic to neurons following prolonged incubation.
6. Prepare a 50 mL conical tube with ~50 ml Wash medium, warm it up to 37 °C.
7. At the end of 2 h, bring the low density neuron with DNA-calcium phosphate precipitates into the hood. Pick up each individual coverslip with a sharp #5 tweezer, and dip it three times in the 50 ml Wash medium to wash it briefly. Then return it to the high density neurons in its original co-culture plates, with low density neurons facing downwards.
8. Continue the culture until the low density culture neurons on coverslips are harvested for studies.