A Scalable Method to Study Neuronal Survival in Primary Neuronal Culture with Single-cell and Real-Time Resolution

All procedures using animals should be supervised by the bioethical animal committee of the institute and performed in compliance with local regulations. The procedures presented herein were approved by the delegated authority and comply with the regulations in Spain and Europe.

1. Primary neuronal culture

NOTE: All the steps are performed inside the culture hood, using sterile materials and solutions to maintain sterile conditions.

1. Poly-L-lysine (PLL) coating
   1. Wash the coverslips (CVs, 15 mm) in 70% ethanol overnight (o/n) in an orbital shaker.
      NOTE: The speed of the orbital shaker should be enough to move the CV gently but not too fast to avoid damaging or scratching the CVs. The specific RPM will vary depending on the model of the shaker. CVs can be stored in ethanol for long periods (months).
   2. Remove the ethanol and wash twice with deionized water for 10 min each.
   3. Let the CVs dry out at room temperature (RT) in a 100 mm Petri dish (13 CVs/plate approximately). Ensure that the CVs do not touch each other or the edges of the plate.
      NOTE: Dry CVs can be stored at RT for long periods (months).
   4. Prepare the 5 mg/mL PLL stock solution in water, make aliquots, and store them at -20 °C. Dilute the stock solution in 1x PBS to a working concentration of 0.1 mg/mL.
      NOTE: Higher concentrations of up to 1 mg/mL PLL may be tested.
   5. Add the minimal volume of PLL to cover the surface of the CVs (see optimal volumes of PLL in Table 1). Add the PLL solution dropwise to facilitate the formation of a droplet.
      NOTE: Plating neurons on CVs is required for immunofluorescence. For time-lapse imaging (e.g., for the IN Cell), cells are plated directly over the well without CVs because the CVs may move during the time-lapse. In that case, add the minimal volume of PLL to cover the surface of each well.

Table 1: Optimal volumes of poly-L-lysine according to the surface area.

6. Incubate for a minimum of 1 h at 37 °C.
   NOTE: Incubation period could be as long as o/n.
7. Remove the PLL from the CVs. Wash the CVs twice with deionized water for 10 min each. Let the CVs dry at RT.
   NOTE: The PLL solution can be stored at 4 °C for reuse (3 times maximum).
8. Store the dried CVs inside a Petri dish at 4 °C (for up to 7-10 days) or place them inside the wells with plating medium (see composition in Table 2 and working volumes in Table 3). Add plating medium directly to the wells of 12-well plates (Table 3).
   NOTE: Neurons are very sensitive to the quality of the fetal bovine serum (FBS). Different batches of FBS should be tested to identify the batch in which the cells look healthier. Once a good lot of FBS is identified, it is recommended to obtain a large amount and store it at -80 °C. This protocol could be adapted for other types of CVs and plates (see the different working volumes in Table 3).

Table 2: Composition of the different media used.

Table 3: Surface area of the different types of plates and the working volumes.

2. Preparation of the neuronal culture
   1. Prewarm (37 °C) and equilibrate (in a 5% CO₂ atmosphere) the plating medium, neuronal medium, and electroporation medium (see composition in Table 2) in a partially opened bottle inside the cell incubator for a minimum of 2 h. Let the trypsin warm up at RT. Pre-warm the 1x Hank's Balanced Salt Solution (HBSS) at 37 °C in a water bath.
   2. Preset the electroporation parameters (Table 4).

Table 4: Electroporation settings for NEPA21.

3. Brain dissection and harvesting CD-1 mouse embryos at E15 post-coitum
   1. Sacrifice the pregnant female by cervical dislocation without sedation. Sterilize the belly with 70% ethanol; cut and open the abdomen with sharp scissors.
   2. Extract the uterus and place it in a Petri dish with cold 1x HBSS. Leave it on ice inside the cell culture hood with common forceps.
   3. Extract an embryo from the uterus with the aid of forceps and scissors and decapitate it with a single cut using scissors.
   4. Hold the head and introduce forceps (e.g., Dumont fine forceps) into the eye orbit.
   5. Using the needle (30 G needle syringe) as a blade, cut the bregma and break through the posterior part of the head.
   6. Introduce sharp forceps through the incision vertically into the bregma, and break the thin layer of skin of the anterior region. Pull the forceps to follow the longitudinal fissure between the two hemispheres without damaging the cortex.
   7. Use the small forceps at a 45° angle, from the anterior to the posterior part, to remove the brain from the head.
   8. Move the brain to a 3.5 cm Petri dish with ice-cold 1x HBSS under a stereomicroscope.
4. Dissection of cortices
   NOTE: The protocol for neuronal culture is adapted from the procedure described by Dotti and Banker¹⁰.
   1. Use the Dumont forceps (11 cm) to hold the thalamus and a needle tilted at a 45° angle to separate the cortex of each hemisphere from the midbrain. Ensure that the cortex does not show traces of other tissues; use sharp forceps to remove the meninges.
      NOTE: The meninges form a thin elastic membrane enveloping the brain and are easily recognized because of their red color. Making soft movements in zig-zag will help remove the meninges. Make sure all the meninges have been removed as they are toxic for the culture.
   2. Move the clean cortex to a clean 3.5 cm Petri dish with ice-cold 1x HBSS and place it on ice.
5. Disaggregation
   1. Chop each cortex into two halves and use forceps or a sharp needle (30 G) to facilitate the effect of trypsin in the next step.
   2. Transfer the pieces to 7 mL of trypsin-EDTA buffer at room temperature in a 15 mL tube using a P1000 pipette (cut off 3-4 mm of the tip). Wait until they sink to the bottom of the tip before releasing them from the pipette tip. Mix by tilting.
   3. Incubate at 37 °C for 12 min in a water bath. Mix by tilting every 4-5 min.
      NOTE: Incubation time could vary from 12 to 15 min.
   4. Let the tissue sink by gravity and carefully remove the trypsin. Wash the tissue twice with 8 mL of 1x HBSS and agitate gently by tilting. Remove the 1x HBSS carefully using a P1000 pipette but using an uncut pipette tip.
   5. Wash the tissue once with 5 mL of plating medium. Agitate by tilting. Remove the plating medium carefully using a P1000 pipette.
   6. Add 1.5 mL of plating medium. Disaggregate the mix mechanically using a P1000 pipette by aspirating the medium from the bottom of the tube and releasing it gently over the walls for a maximum of 8-12 times.
   7. Filter through a sterile cell strainer (70 µm) placed on a 50 mL tube. Add 10 mL or other desired volume of plating medium to the tube.
   8. Introduce 5 µL of the cell suspension into the cell counting chamber, add 5 µL of trypan blue (0.4% w/v), and count the cells.
   9. Plate the cells (see section 1.6) by transferring an adequate volume of the cell suspension to have the desired number of cells for electroporation (e.g., 10⁶ cells) in a 15 mL centrifuge tube. Centrifuge the cells at 250 × g for 5 min. Proceed to section 1.6.
6. Plating cells
   1. Plate the desired density of the uncentrifuged cell suspension from step 1.5.7 in each well containing the plating medium.
      NOTE: In this protocol, 100,000 to 200,000 cells/well were seeded in a 12-well plate. As a reference, plate at least 3 CVs per experimental condition to account for experimental variability. A typical experimental setting may entail imaging 4-5 fields per CV (e.g., using a 20x objective, each field would be expected to be ~1.3 mm² wide).
   2. Move the plate in the cell culture hood in all the cardinal directions to distribute the cells uniformly.
7. Electroporation
   NOTE: Other transfection systems may be used. The NEPA21 system used in this protocol has the advantage of being an "open system." The protocol can be modified by the user, and it does not require specific reagents for the transfection of special cuvettes used for the electroporation. This protocol was modified from ⁵.
   1. Continue from step 1.5.9. Resuspend the cell pellet with 5 mL of electroporation medium in a 15 mL tube. Centrifuge the tube at 250 × g for 5 min.
   2. Repeat step 1.7.1. Resuspend with electroporation medium to obtain a concentration of 10⁶ cells/mL in each of the electroporation cuvettes. Mix the cells with the desired amount of DNA in the cuvette.
      NOTE: For GFP expression, 3 µg of plasmid was used per million cells. Because of the high variability, it is recommended to test the optimal concentration for each plasmid. Between 2 and 24 µg of plasmid per million cells have been tested. Diverse plasmids can be combined in the same cuvette to perform a double-transfection. In the case of co-transfection, transfect the gene of interest in excess as compared to GFP (e.g., 3:1 is an optimal ratio that should be tested) to ensure that most GFP-expressing cells co-express the gene of interest.
   3. Place the cuvette into the cuvette chamber (Figure 2). Press the Start button to execute the preset program (see electroporation conditions in Table 4). Record the values of currents and joules displayed in the Measurements frame.
      NOTE: The user may change specific conditions. For instance, decreasing the voltage may reduce the efficiency of transfection but increase neuronal survival.
   4. Take the cuvette out of the chamber. Add 500 µL of the plating medium into the cuvette immediately after the electroporation and transfer the sample from the cuvette into a 1.5 mL centrifuge tube using a pipette.
      NOTE: The faster the addition of medium, the better the survival after electroporation as it helps to avoid cell damage.
   5. Plate the desired number of electroporated cells in each well.
      NOTE: Plate from 20,000 to 100,000 cells in a 12-well plate. Depending on the conditions, only 5-10% of the electroporated cells may survive. The exact number should be tested and adapted by the user.
   6. Incubate for 2-4 h in standard conditions (37 °C, 5% CO₂, saturated humidity).
      NOTE: After the incubation period, observe the cells under the microscope. If the culture is healthy, most cells would have adhered to the surface and started sprouting visible neurite extensions.
   7. Change the plating medium to neuronal medium (volumes and composition in Table 1 and Table 2, respectively).
8. Maintaining cells
   1. To refresh the medium, remove half of the medium and add an equal volume of fresh pre-equilibrated medium after 7 days of culture.
   2. On day 10 of culture, replace half of the volume with fresh neuronal medium without the glutamine supplement (see the Table of Materials). Repeat this step every other day after day 10.
      ​NOTE: After 10 days of culture, cells produce glutamate by themselves.

2. Construction of the hypoxic chamber

1. Use any hermetic container with a screw lid or an equivalent sealing system for this purpose (Figure 3A). Drill two holes in the wall of the plastic container and introduce a plastic tube in each one. Seal them with silicone so that the container is hermetically sealed (Figure 3).
   ​NOTE: Here, a cylindrical plastic container (size 1 liter) with a screw lid was used to construct the chamber. Ensure that the container is big enough to accommodate the CVs required for the experiment. To treat more CVs in the same space, plastic platforms can be stacked on top of each other (Figure 3A).
2. To prepare the bubbling chamber (see 3.2), drill a hole into the cap of a flask, using any standard mechanical drill with a drill slightly smaller than the tube, and introduce a plastic tube through the hole into the flask. Seal the hole with parafilm to maintain the hermetic seal (Figure 3B).
3. Sterilize the container and the flask with 70% ethanol and irradiate with ultraviolet light for one cycle before using them.

3. Oxygen-glucose deprivation (OGD)

NOTE: The protocol for OGD is adapted from Tasca et al.⁷

1. Prepare the OGD solution with phosphate solution (Table 2) that has been previously supplemented with antibiotics.
   NOTE: For each 35 mm plate, use 5 mL; for a 100 mm plate, use 10 mL.
2. Add the OGD solution inside the bubbling chamber (Figure 3B) and place it inside a water bath (already prewarmed at 37 °C). Connect the flask with the nitrogen source (N₂, 100%).
   NOTE: Please note that because the OGD solution contains HEPES as the buffer, it does not require CO₂. If the OGD solution is based on a bicarbonate buffering system, use 5% CO₂ and 95% N₂ to saturate the hypoxic solution and chamber.
3. Saturate the OGD solution with gently bubbling nitrogen. Leave the solution in a mildly bubbling state for 20-30 min.
4. Immediately transfer the flask to the cell culture hood and add 5 mL of the OGD solution without oxygen in 35 mm plates or 10 mL in 100 mm culture plates.
5. Move the CVs with the cultured cells to the plates with OGD. Without covering the plates, move them into the hypoxia chamber and close them firmly but carefully.
   NOTE: Steps 3.4 and 3.5 must be completed as quickly as possible. Maintain the control CVs (Neuronal Medium in normoxia) in the well plates inside the incubator. Alternatively, expose the control neurons to the normoxic OGD medium supplemented with glucose (10 mM) to examine the effects of hypoxia and glucose deprivation more selectively.
6. Connect the nitrogen system to the hypoxia chamber and check that the tubes are open. Saturate the chamber with N₂ for a few minutes at a pressure of 2-3 bars (e.g., 2-3 min; 1 bar = 750 mmHg) (Figure 3C).
   NOTE: Do not forget to bubble the OGD solution again for 15 min between multiple time points.
7. Reduce the nitrogen pressure to 1.3-1.5 bars and close both tubes of the hypoxia chamber (Figure 3C). Leave the chamber in the incubator for the required time (e.g., 60 min) at 37 °C, 5% CO₂.
   NOTE: OGD duration may require adjustment depending on the experimental condition and the research goals. A short incubation time (e.g., 30 min) would simulate mild hypoxia. Two hours of OGD are usually very harsh toward the neurons. It is recommended that an OGD duration of 45 to 90 min be tested before starting the study. Of course, the duration of the OGD will impact the total survival and the relative onset of neuronal death after recovery from OGD. If the goal is to analyze neuronal death in real time, aim to achieve OGD conditions that would provoke cell death a few hours after recovery from OGD for a more effective analysis.
8. Open the chamber and move the CVs to their original multiwell plate for recovery. After this step, consider two alternatives: 1) for small-scale experiments, proceed to section 4 for immunofluorescence and image analysis; 2) for high content screening, proceed to section 5 for the analysis in the cell analyzer (or equivalent platform).
   ​NOTE: Live imaging provides a more flexible approach because it allows real-time monitoring of the different experimental points.

4. Analysis by standard immunofluorescence and ImageJ

NOTE: Recovery time may vary from 1 to 24 h depending on the conditions.

1. Wash the neurons with 1x PBS for 10 min.
   NOTE: This step is optional as there are several ways to proceed and analyze the images.
2. Fix the neurons with 4% paraformaldehyde for 10 min at RT. Wash them three times with 1x PBS for 10 min each wash.
   NOTE: As paraformaldehyde is toxic, work under a chemical hood.
3. Permeabilize the neurons by adding 1x PBS containing 0.1% Triton X-100 for 10 min. Wash the cells quickly with 1x PBS. Repeat the washing step twice for 10 min each wash.
4. Add 350 µL of blocking solution (1x PBS containing 2% bovine serum albumin) to the CVs and leave them for at least 30 min. Spin the primary antibody (AB) stock (5 × g for 30 s) and mix it with the blocking solution at the required dilution, e.g., anti-GFP at 1:600.
   1. Transfer the coverslips to parafilm in a humidified chamber (a box with water-soaked tissue to maintain humidity and avoid evaporation). Depending on the size of the CVs, carefully add 60 or 80 µL of the primary antibody dilution to each CV, and ensure that the cells are in contact with the antibody. Incubate the CVs o/n at 4 °C.
      NOTE: A droplet should be formed on the CV surface to cover the entire CV surface. Avoid disturbing the cells while pipetting the antibody solution on the CVs. Incubation time could be reduced to 2 h at RT.
5. Wash the CVs with 1x PBS for 10 min over the parafilm. Repeat the washing step twice.
6. Dilute the secondary AB, e.g., Alexa 488 (1:500), and 4′,6-diamidino-2-phenylindole (DAPI, 1:2,000) in blocking solution. Depending on the size of the CVs, add a 60-80 µL droplet of the secondary AB solution on the cells. Incubate the CVs for 1 h in the dark at RT. Wash the secondary AB and DAPI from the neurons with 1x PBS for 10 min and repeat this step twice.
7. Mount the CVs in mounting medium (see the Table of Materials) for immunofluorescence analysis and store them in the dark at 4 °C after drying.
8. Image capturing and analysis.
   NOTE: An inverted microscope was used to take the images at 20x (Figure 4A, see the Table of Materials).
   1. Use the open-source ImageJ software to analyze the images and obtain the mask for the GFP channel (Figure 4C).
      NOTE: There are several ways of imaging and analyzing the neurons, depending on equipment availability and the experimental format. The following workflow is provided as a basic example wherein the objective is to identify the soma of individual GFP-expressing neurons present in the image. The specific values presented here, such as the threshold value and particle size, should be tested by the user based on their specific experimental conditions. In addition, different filters and processing steps may be added. For instance, the ImageJ filters Subtract Background and Despeckle often help in cleaning the image at the beginning of the processing. Optimization of the workflow for specific experimental conditions requires previous experience in image analysis and the knowledge of basic concepts of quantitative image analysis. As an in-depth guide on image analysis is out of the scope of the current protocol, we recommend the following resources: ImageJ user guide, https://imagej.nih.gov/ij/docs/guide/; for examples and tutorials, https://imagej.nih.gov/ij/docs/examples/index.htm; MBF ImageJ bundle, https://imagej.net/mbf/index.htm.
   2. For segmentation, select the Intensity segmentation with the necessary parameters to isolate the soma, which is the brightest region of the neurons.: Menu | Image | Adjust | Threshold | Set min threshold (e.g., 100) | click on Apply.
   3. Repeat the threshold from 4.8.2. to all the images to see the mask (i.e., the segmented image) for the somas of the neurons (Figure 4E).
   4. To obtain the number of surviving cells in each image and export the results to a spreadsheet to perform calculations, select Menu | Analyze | Analyze particles and enter the necessary parameter values (e.g., Size: 50-infinity pixel; Circularity: 0.5-1.00). Consider that at t = 0 h, the number of cells counted are the total number of cells identified, and calculate the percentage of cells that survived at the different time points.
      NOTE: A lower size threshold is useful for filtering out smaller objects than the soma (e.g., 60-80 µm²).
   5. Use the Record Macro function of ImageJ to automatize and repeat the analysis: Menu | Plugins | Macros | Record | click on Create to generate the macro.

5. Real-time analysis with IN Cell Analyzer 2200

1. Insert the 12-well plate in the cell analyzer, connect the CO₂ system, and set the temperature at 37 °C.
   ​NOTE: This step can be easily scaled up in this platform, performing a similar experiment in neurons plated in a 96 wells plate.
2. Select the 20x objective, randomly select 5 fields per well (covering 2.33% of the well in this example), and focus them automatically for each fluorescence channel (fluorescein isothiocyanate [FITC] in this case). Select the parameters of acquisition (30 min time-lapse for 16 h). Press Start and wait.
   NOTE: The number of imaging fields must be determined by the investigator and should be sufficient to account for the variability inside a single well.
3. Use IN Cell Developertoolbox v1.9 to set up the appropriate workflow for analysis.
4. Analyze the neuron images (FITC channel) to generate a mask for the soma of the neurons (Figure 4).
   NOTE: The objective is to identify the soma of individual neurons present in the image. This analysis follows steps that are equivalent to those describes in section 4 for ImageJ. Please refer to section 4 for the workflow and general considerations. The intensity threshold is determined to select the soma, which is the brightest region of the neurons.
   1. For segmentation, select the Intensity segmentation with the necessary parameters (in this case, a minimum threshold of 20005.40).
   2. For postprocessing, select the Sieve option, keeping targets with an area greater than a predetermined value (here, 189 pixels (~80 µm²)).
   3. Select the Sum parameter to count and sum all the neurons identified in the mask (surviving neurons) for each field and for all the fields of a well at each time point of the time course.
   4. Export the results to a spreadsheet. At t = 0 h, consider that the number of cells counted are the total cells identified, and calculate the percentage of cells that survived at the different time points. Calculate the time at which 50% of the cells are dead to facilitate the comparison between conditions.
      NOTE: The parameters for the segmentation and postprocessing steps must be adjusted by the user for different fluorescent channels depending on the structures to be identified.