Generation of Functional Mouse Hippocampal Neurons

Primary culture of mouse hippocampal neurons is a very useful in vitro model for studying neuronal development, axonal and dendritic morphology, synaptic functions, and many other neuronal features. Here we describe a step-by-step process of generating primary neurons from mouse embryonic hippocampi (E17.5/E18.5). Hippocampal neurons generated with this protocol can be plated in different tissue culture dishes according to different experimental aims and can produce a reliable source of pure and differentiated neurons in less than one week. This protocol covers all the steps necessary for the preparation, culture and characterization of the neuronal culture, including the illustration of dissection instruments, surgical procedure for embryos’ isolation, culturing conditions and assessment of culture’s purity and differentiation. Evaluation of neuronal activity was performed by analysis of calcium imaging dynamics at six days in culture.

Many protocols have been developed for generating cortical and hippocampal neuron cultures by co-culturing neurons with glial feeders (Kaech and Banker, 2006) describing three-dimensional neuronal culture systems with astrocytes encapsulated in hydrogel microfibers (Kim et al., 2020), supplementing culture medium with growth factors for longterm neuron cultures (Ray et al., 1993), or generating neurons from post-natal mice (Beaudoin et al., 2012). Each of these methodologies presents advantages and disadvantages depending on the goals of the experiment. Here, we extend and describe in detail a protocol for generating hippocampal neurons that we have developed and successfully employed over the years (e.g., Dorsey et al., 2006; Tomassoni-Ardori et al., 2019). We focus primarily on providing an easy, simple and reliable procedure for obtaining primary hippocampal neurons from mice at the late embryonic stage (E17.5-E18.5). We illustrate the whole preparation process including aspects that have often been omitted in other protocols such as a description of dissection instruments and surgical procedures for embryos removal and dissection. We report analysis of neuronal differentiation and function after 6 days in culture by testing for neuronal differentiation markers and neuronal activity by cellular calcium imaging dynamics. Moreover, we show the levels of neuronal culture purity in relation to glial cells with and without treatment with the glial inhibitor cytosine arabinoside (Ara-C).
This step-by-step protocol allows the experimenter to obtain differentiated hippocampal neurons with a high degree of purity within 6 days.

Procedure
Notes:

a.
This methodology describes the critical steps necessary for generating primary hippocampal neurons derived from E17.5/E18.5 mouse embryos. Practical tips after each step are included to better guide the readers through the procedure.

b.
Depending on the final experimental aims, different tissue culture dishes/plates or glass coverslips can be used for culturing primary hippocampal neurons.

A.
Coating of tissue culture dishes/plates with poly-D-lysine

1.
Reconstitute poly-D-lysine in a sterile laminar hood by adding 50 ml of sterile tissue culture grade water to a 5 mg γirradiated and cell culture tested poly-D-lysine bottle (final working concentration of 0.1 mg/ml).

2.
Evenly coat the culture dish surface with 0.1 mg/ml poly-Dlysine solution and incubate at room temperature for 20 min in a laminar flow hood.

3.
Remove the poly-D-lysine solution by aspiration and rinse the surface thoroughly with sterile tissue-culture grade water twice.

4.
Before plating the cells, allow the surface to dry completely by leaving the dishes/plates uncovered under the laminar flow hood for about 1 h.
Note: If plating cells on glass coverslips use commercially available poly-D-lysine/laminin pre-coated 12 mm coverslips. Transfer poly-D-lysine/laminin coated coverslips directly into wells of a 24-well plate by using sterile forceps in a sterile laminar flow hood before plating neurons. Alternatively, regular 12 mm coverslips can be coated by placing them into 24-well plate wells and covering them with 0.1 mg/ml poly-Dlysine for 20 min in a laminar flow hood. After that wash twice with PBS and cover with a 10 μg/ml laminin solution for at least 4 h at 37 °C by leaving the coverslips in an incubator until cells are ready to be plated. Rinse twice with PBS just before plating the cell to prevent the culture surface from drying. The 1 mg/ml laminin stock solution is made by adding 1 ml of PBS into 1 mg laminin powder and diluting to 10 μg/ml laminin with PBS. Coating of coverslips in plates avoids the potential to scrape the coating off while transferring between dishes.

1.
Euthanize pregnant mouse according to Animal Care and Use Committees (ACUC) approved protocols (CO 2 euthanasia or isoflurane anesthesia followed by decapitation or cervical dislocation).

2.
Pin the animal to a clean surgical area and use 70% ethanol to clean the abdomen.

3.
In order to expose the embryos, perform an incision in the middle of the abdomen using surgical scissors ( Figure 1A) starting from the pubic region and continuing up to the end of the abdominal cavity as indicated by the dashed line in Figure  2A.
Note: Use forceps ( Figure 1B) to pinch and pull up the skin layer while performing the incision with surgical scissors. This will help to avoid accidental damage to the embryos located immediately underneath.

4.
Carefully expose the whole uterus (uterine horns) containing the embryos ( Figure 2B) by using forceps and surgical scissors ( Figures 1A-1B) and immediately transfer the uterus to a 100 mm dish containing ice-cold PBS ( Figure 3A).
Note: Use forceps to grab and pull the uterus out of the abdomen while using surgical scissors with the other hand to cut any adherence and the cervix connection ( Figure 2B arrows).

5.
Continue to use forceps and surgical scissors to free the single embryos and carefully remove the placenta and yolk sack. Transfer the embryos into a new 100 mm dish containing icecold PBS to wash embryos and to remove blood and embryonal fluids ( Figure 3B).

6.
Use forceps ( Figure 1B) to hold the embryo and cut the head using surgical scissors ( Figure 3C). Transfer the heads into a new 100 mm dish containing ice-cold DMEM complete with 10% fetal bovine serum and 1% antibiotics (P/S).
Note: From this step forward, it is important to keep transferring the biological material to new dishes with clean medium containing antibiotics to lower the possibility of contamination. In the figures shown in this protocol PBS was intentionally used instead of medium containing phenol red only for the purpose of obtaining more clear images ( Figure  3A, Figure 4, Figure 5 and Figure 6).

C.
Dissection of hippocampi

1.
Move to a dissection cabinet with sterile laminar air flow and prepare a 100 mm dish containing ice-cold complete DMEM with 10% fetal bovine serum and 1% antibiotics (P/S) for dissection of the brains. Transfer one embryo head at a time into the 100 mm dish containing complete DMEM medium and gently remove the brain from the embryo head ( Figure  4A). Specifically, with one hand, use forceps ( Figure 1B) to pinch the mouth/nose area of the head (black arrow in Figure  3D). With the other hand, use the fine scissors ( Figure 1C) to enter from the back of the skull (black scissors in Figure 3D) and execute a transversal cut (along the transversal plane at the mouth/nose level indicated by the white dashed line in Figure  3D) in order to free the basal brain region. Invert the embryo head to expose the free basal area of the brain. Continue by using two forceps ( Figure 1B) to gently pull apart the skull and carefully remove the whole brain ( Figure 4A). Transfer and collect the brains into a new 100 mm dish containing clean, ice-cold complete DMEM medium. Replace the medium after each brain dissection to eliminate the blood.

2.
Transfer one brain at a time to a 35 mm dish lid filled with icecold complete DMEM medium and move under a dissection stereoscope.
Note: The lower depth of the lid of a 35 mm dish allows for better movements when using dissection instruments compared to the bottom of the 35 mm dish.

3.
Use two curved forceps ( Figure 1D): hold the brain on the side with one forceps (white dots in Figure 4A) and insert the second forceps along the midline (dashed line in Figure 4A) to separate the two hemispheres of the brain. Continue to use curved forceps to gently pull apart and flip the two halves of the brain in order to expose and visualize the hippocampal area ( Figures 4B-4C).
Note: Due to the fragility and softness of the embryonal tissues, the brain can be easily damaged while trying to dissect the hippocampal region: always use care when handling and holding embryonal brains. A few practicing sessions at the stereoscope will help to improve dissecting skills quickly.

D.
Work by using two Dumont forceps ( Figure 1E). Use one forceps to hold the whole brain and use the second forceps to gently peel the meninges off. Start by pinching the olfactory bulb area and gradually pulling back in order to remove meninges completely ( Figure 5). Hold the brain gently while performing this operation and it might be necessary to change the holding points many times according to how easily the meninges come off.

E.
It is important to properly remove the meninges to eliminate as much non-neuronal cell contribution to the neuronal culture as possible.

1.
Proceed by using Dumont forceps to finely cut along the hippocampus edges to completely free the hippocampal area ( Figures 5-6). Use curved forceps to gently pick and transfer each hippocampus to a new 35 mm tissue culture dish containing 2 ml of ice-cold serum-free DMEM medium with 1% antibiotics (P/S) and keep it on ice.
Note: The hippocampus is distinguishable by its typical Cshaped anatomical structure and by a difference in light contrast under a stereoscope compared to the cortical area to which it is connected (Figures 5-6). After removing the hippocampus it is possible to collect cortices from the same embryos for the preparation of cortical neurons if needed (not described in this protocol).

F.
Trypsinization of the hippocampi

1.
Mince the hippocampal tissues in the same 35 mm tissue culture dish containing 2 ml of ice-cold serum-free DMEM medium used for the collection by cutting them into small pieces using two Dumont forceps (ideally 5-6 pieces from each hippocampus of approximately 2-3 mm 3 ) ( Figure 6C).
Note: It is important to collect and keep the hippocampi in serum-free DMEM medium (Steps C5 and D1) to avoid inhibition of Trypsin-EDTA activity during the next step (Step D2).

2.
Move to a tissue culture hood with sterile laminar air flow and use a pipette to transfer the medium containing the minced hippocampi to a 50 ml sterile conical tube. Gently add 2 ml of 0.25% Trypsin-EDTA to the hippocampi and place the tube in a water bath at 37 °C for 25 min (final trypsin concentration of 0.125%). Note: During the 25 min digestion in trypsin, the minced hippocampal pieces will naturally sediment by gravity to the bottom of the 50 ml conical tube.

3.
Prepare a 15 ml sterile conical tube containing 2 ml of warm (37 °C) DMEM complete with 10% fetal bovine serum and 1% antibiotics (P/S). At the end of the digestion, carefully transfer the 50 ml digestion tube back to a tissue culture hood without disturbing or resuspending the hippocampal sediment and, with a pipette with a 1 ml tip, transfer only the hippocampal sediment to the new 15 ml conical tube containing the 2 ml of complete medium.
Note: Use the 1 ml tip to directly contact the hippocampal sediment at the bottom of the 50 ml tube. Gently aspirate and transfer only the hippocampi at the bottom of the tube without excess liquid from the digestion. The fetal bovine serum contained in the new 2 ml medium will inactivate the trypsin and stop its enzymatic activity.

G.
Trituration of the hippocampal tissue

1.
Use a Bunsen burner to fire-polish a sterile glass Pasteur pipette and generate a smaller opening and a smoothened tip for dissociating the hippocampal tissue (Figure 7). Use the fire-polished pipette shown in Figure 7B to slowly and gently triturate the hippocampal pieces. Pipet up and down six to eight times.
Note: Trituration of the hippocampi is a critical step to achieve good cell yield and cell viability. After fire-polishing, be sure to let the glass pipette cool down before triturating the hippocampi. Hippocampal chunks should be completely broken up after pipetting up and down six to eight times.

2.
Use the Bunsen burner again to prepare a new fire-pulled glass Pasteur pipette with a longer and tapered tip ( Figure 7C). Use this pipette to slowly perform a single up and down pipetting of the hippocampal cell slurry in order to release single cells.
After obtaining a single cell suspension proceed immediately to Procedure F. Keep the single-cell suspension in a tissue culture hood with sterile laminar air flow at room temperature while counting the cells.
Note: Performing multiple pipetting with the fire-pulled Pasteur pipette with a longer and tapered tip ( Figure 7C) will damage the cells and affect cell viability.

H.
Cell counting, cell seeding and cell culture

1.
Collect 50 μl of the cell suspension and add 5 μl of Trypan blue dye to assess cell viability. Count cells using a cell counter.
Note: Cell viability ranges between 70-90% of live cells. As an example, the cell viability and cell count of a neuronal preparation from 8 embryos is about 4 × 10 6 live cells from a total of 5 × 10 6 cells (cell viability = 80%).

2.
Centrifuge the remaining hippocampal cell suspension at 350 × g for 5 min. Discard the supernatant and resuspend the cell pellet by pipetting gently up and down in warm Neurobasal medium with B27 supplement. Adjust the volume of medium according to the tissue culture plates that will be used. Use a seeding density of approximately 260,000 cells/square centimeter. Seed the cells in the tissue culture dish/plate previously coated with poly-D-lysine.
Note: Cell yield is approximately 400,000-600,000 cells/ embryo. Prepare complete medium for neuronal culture by adding 10 ml of B27 supplement and 5 ml of P/S antibiotics (100x concentrate) into 500 ml of Neurobasal medium. Some protocols (e.g., Ruhl et al., 2019) suggest to pre-plate neurons for one hour in DMEM medium with 10% FBS to facilitate neuron adhesion to the plate. However, we have found that plating directly in Neurobasal medium with B27 supplement is sufficient for good neuron adhesion.

3.
After 18 to 24 h from the initial plating, add cytosine arabinoside (Ara-C) to the culture to stop the proliferation of dividing cells (non-neuronal cells). Remove half of the volume of the culture medium (Neurobasal medium containing B27 supplement) and replace it with fresh medium containing 2 μM Ara-C (1 μM Ara-C final concentration).
Note: Failing to add Ara-C to a primary neuron culture will considerably affect the purity of the culture (Figures 8B and  8F). Death of some mitotic cells can be observed after adding Ara-C to the neuron culture.

4.
Culture the primary hippocampal neurons by replacing only half of the medium every 48 to 72 h. It is not necessary to continue adding Ara-C when replacing medium after the initial treatment. After 6 days in culture, the primary neurons will be well developed and differentiated as assessed by both morphology but especially expression of typical neuronal markers such as Tubulin-β-III, NeuN, Rbfox1, neurotrophin receptors and several other neuronal proteins (Tomassoni-Ardori et al., 2019) ( Figure 8 and Figure 9). Notes:

a.
Replacing only half of the culture medium maintains the growth factors naturally secreted and present in the neuronal conditioned medium.

b.
Although absolute prevention of microbial contamination (bacteria, mycoplasma, fungi) during the preparation of primary cell cultures is impossible, these steps can greatly improve the outcome: i) work in a sterile environment; ii) use proper aseptic technique including wearing sterile gloves and using sterile glassware and disposable pipettes and tips; iii) keep all the equipment clean (dissecting microscope, incubators, centrifuges and water baths).

I.
Western blot analysis for the detection of markers of differentiated neurons ( Figure 8B)

1.
After 6 days of in vitro culture, remove the culture medium from one well of a 24-well plate containing neurons.

2.
Rinse cells with PBS once.

3.
Lyse cells by adding 150 μl of 2x Laemmli sample buffer directly into the well.

4.
Transfer the cell lysate to a 1.5 ml tube and sonicate the sample for 10 min to shred genomic DNA and eliminate viscosity.

5.
Heat the sample at 95°C for 5 min to promote protein denaturation.

6.
Load 10 μl of sample per lane into a 4-12% NuPAGE pre-cast gel and run the gel at 120 V.

7.
Transfer gel to a PVDF membrane.

11.
Incubate membrane with HRP-conjugated secondary antibodies at room temperature for 1 h while gently shaking. Note: Although in this protocol we show that differentiated hippocampal neurons can be obtained by 6 days in culture and we recommend the use of Ara-C for improving neuronal culture purity (Figure 8 and Figure 9), in some cases coculturing neurons with glial cells can further support neuronal maturation and long-term survival (Kaech and Banker, 2006;Kim et al., 2020).

I.
Functional phenotypic characterization of hippocampal neurons by calcium imaging (Figure 10 and Video 1)

A.
A further test of functionally differentiated neurons includes imaging of the calcium dynamic occurring in vitro following spontaneous or glutamate-induced depolarization (Grienberger and Konnerth, 2012

2.
After 6 days in vitro, remove the culture medium from the 24-well plate with the coverslips containing the hippocampal neurons. Incubate each coverslip with 250 μl of Fluo-4 loading buffer at 37 °C for 30 min following by a 30 min wash in ECS buffer at 37 °C in order to allow the de-esterification and activation of Fluo-4.

3.
Adjust the flow in the recording chamber to 0.5 ml × min and set the temperature to 37 °C.

4.
Place the coverslip in the recording chamber and select a suitable field.

5.
Set focus, set laser intensity (480 nm) at a lower possible value and integration time (pixel dwell time) such that the acquisition time is shorter than 500 ms before starting the time series acquisition at the confocal at 2 images × second.

6.
Record neuronal spontaneous activity and then apply glutamate at 1 mM for 10 s to depolarize and visualize all the cells in the field.

7.
For image analysis import the file in ImageJ (      A. Dissected embryo brain. Dashed white line indicates where to separate the two brain hemispheres using one curved forceps while holding the brain on the side by using the second curved forceps (white dots) (Step C3). B. Separated brain hemispheres. White arrow heads indicate the location of the hippocampal region on each hemisphere (Step C3). C. Higher power image showing the hippocampal region on one brain hemisphere indicated by a white dashed line.

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