Hippocampal Neuronal Cultures to Detect and Study New Pathogenic Antibodies Involved in Autoimmune Encephalitis

All procedures were approved by the local ethics committee of the University of Barcelona following European (2010/63/UE) regulations about the use and care of experimental animals. Written informed consent was obtained from patients, and the study was approved by the local institutional review board for the use of human samples (Hospital Clínic, HCB/2018/0192).

NOTE: There are three parts in the current protocol. The first one is for establishing the neuronal cultures, the second one is for the detection of surface antibodies using live cultures of neurons, and the third one is to determine the pathogenicity of these antibodies.

PART 1: Establishment of dissociated, fetal, rodent hippocampal neuronal cultures

1. Preparative steps

1. Poly-L-Lysine coating of the plating surfaces (3 days before dissection)
   1. Prepare the borate buffer by adding 2.38 g of boric acid and 1.27 g of borax to 500 mL of distilled water and stirring for 15 min until dissolved. Under a hood, sterilize the buffer solution by filtering (0.2 µm pore size), label, and store at room temperature (RT).
      NOTE: This solution is stable and can be used for 6 months.
      CAUTION: When preparing borate buffer, wear recommended personal protective equipment.
   2. Prepare the Poly-L-Lysine (PLL) stock solution (100 mg/mL) by adding 1 g of PLL to 10 mL of distilled water and stirring until dissolved. Prepare 500 µL aliquots of PLL stock solution. To reach a final concentration of 1 mg/mL, add 500 µL of PLL aliquot to 50 mL of borate buffer and swirl until dissolved, and sterilize the solution by filtering (0.2 µm pore size).
      NOTE: This solution is stable and can be used for 6 months.
   3. Prepare the surfaces for coating. Use 12 mm diameter coverslips for immunostaining procedures and glass-bottom dishes for studies that will include the imaging of live neurons. If using coverslips, autoclave and then place five coverslips in each dish (3.5 cm diameter).
      NOTE: The thickness of the coverslip affects the quality and intensity of the signal of the images acquired. Most microscope objectives are designed for #1.5 coverslips. Choose the appropriate coverslips according to the imaging setup and techniques.
   4. Under a hood, add 1.5 mL of PLL solution to each dish (glass-bottom dish or 3.5 cm dish with five coverslips). Make sure all the coverslips are submerged and stored at RT for 24 h.
   5. 2 days before dissection, aspirate the PLL solution and wash with sterile endotoxin-free water, making sure the coverslips are submerged. Keep water in the dishes and store at RT for 24 h.
   6. 1 day before dissection, aspirate the water and replace it with NB + B27 culture media (see below), and place the dishes in an incubator at 37 °C for 24 h.
2. Preparation of culture media and stock solution (1 day before dissection)
   1. Dulbecco's Modified Eagle Medium (DMEM): To 500 mL of DMEM High Glucose (4.5 g/L) without L-Glutamine and phenol red, add 50 mL of horse serum, 50 mL of fetal bovine serum (FBS), 10 mL of L-glutamine (200 mM), 10 mL of sodium pyruvate (100 mM), 10 mL of penicillin-streptomycin (10,000 U/mL). Filter (0.2 µm pore size) and store in 5 mL aliquots at 4 ᵒC with a tightly closed cap.
      NOTE: This solution is stable and can be used for 1 month.
   2. Neurobasal (NB) medium supplemented with B27 (NB + B27): To 50 mL of NB medium without phenol red, add 1 mL of B27 supplement.
   3. Hibernate-E medium supplemented with B27 (Hibernate + B27): To 50 mL of Hibernate-E medium, add 1 mL of B27 supplement.
   4. Extracellular Physiological Solution (EPS): To 1 L of sterile water, add the following at the specified final concentrations: NaCl (140 mM), KCl (3.5 mM), HEPES (10 mM), glucose (20 mM), and CaCl₂ (2 mM). Stir until dissolved and adjust the pH to 7.4 by adding NaOH (0.5 M) or HCl (0.5 M). Under the hood, sterilize by filtering (0.2 µm pore size) and store at 4 ᵒC.
3. Equipment and other laboratory materials (day of dissection)
   1. Set a water bath to 37 °C. Heat the following aliquots: 50 mL of HBSS, and 5 mL of DMEM.
   2. Sterilize the following tools by immersing in a beaker containing absolute ethanol (Figure 1B): forceps, curved forceps, scissors, fine-curved forceps, fine-straight forceps, surgery scissors, fine-angled forceps, and precision spring scissors.
      NOTE: To protect the tools, place a soft material (e.g., science precision wipes) at the bottom of the beaker.
   3. Fill two trays with ice and place the following items.
      1. Ice tray 1 (Figure 1C): Place surgical tools in the beaker with absolute ethanol, a beaker with HBSS for rinsing the surgical tools, a 10 cm dish with HBSS to place the embryos, a 6 cm dish with HBSS to place the heads of the embryos, and a 6 cm dish with Hibernate + B27 to place the brains of the embryos.
      2. Ice tray 2: Place a 1 mL aliquot of trypsin 2.5% and a 3.5 cm dish with Hibernate + B27 for the dissected hippocampus.
         ​NOTE: The time needed for this procedure depends on the investigator's experience and the number of embryos to be dissected. Therefore, the ice must remain frozen for the needed amount of time. Polystyrene trays are recommended for this purpose.

2. Dissection and seeding (Figure 1)

1. Hippocampal isolation
   NOTE: Pregnant rats with embryos at E18 are euthanized immediately before the protocol starts in accordance with the local ethics committee of the University of Barcelona, following European (2010/63/UE) regulations about the use and care of experimental animals. In this protocol, carbon dioxide (CO₂) inhalation was used as the method of euthanasia.
   1. Dissect the rat at the level of the abdominal peritoneum using forceps and scissors. Extract the uterus with the E18 embryos and place it in a 10 cm dish that has been chilled on ice.
      NOTE: It is important to avoid the rat hairs attaching to the embryos. It is recommended to use copious amounts of 70% ethanol to sterilize the abdomen. From this step onward, work inside the hood on ice tray 1.
   2. Open the embryonic sacs and transfer the embryos to the 10 cm dish with HBSS, making sure the embryos are fully immersed.
   3. Remove the embryo head with scissors and place it in the 6 cm dish with HBSS. Repeat this process with all the embryos.
      NOTE: To avoid contamination, all tissue except for the embryo heads should be discarded in a container with a screw cap.
   4. Hold the embryo head with fine-curved forceps and pierce the orbits with a pair of fine-straight forceps, entering at a 45° angle, and then release the fine-curved forceps.
      NOTE: The forceps must not go through the brain, so it is important to maintain the angle when entering.
   5. Dissect the skin and skull with the surgery scissors, starting from the occipital bone to the frontal bone. Remove the brain with a pair of fine-curved forceps and place it in the 6 cm dish with Hibernate + B27. Repeat this process until all the brains are recovered.
   6. Sagittally separate the telencephalons with the fine-straight forceps.
      NOTE: From this step onward, the dissection of the hippocampus is carried out under a stereomicroscope. It is recommended to use articulated lamps so that the light illuminates the dissection surface from the sides. It is also recommended to use a black background as this provides more contrast, allowing to better distinguish the hippocampus.
   7. Prepare a 10 cm dish by placing drops of Hibernate + B27 at 1 cm distances (e.g., in a circle). Place one telencephalon per drop of Hibernate + B27 and visualize through the dissecting scope (1.25x objective magnification is recommended).
   8. Carefully peel the meninges and remove the thalamus to better visualize the hippocampus.
   9. Dissect the hippocampus with the precision spring scissors and place it in the 3.5 cm dish with Hibernate + B27 in ice tray 2. Repeat for every telencephalon until all the hippocampi are collected.
   10. Carefully collect all the hippocampi with a Pasteur pipette and transfer them to a 50 mL tube.
       NOTE: Take the minimum volume of Hibernate + B27 when collecting the hippocampi so as not to dilute the trypsin.
2. Cell dissociation
   1. Enzymatic dissociation of the hippocampus
      1. In the 50 mL tube containing the hippocampi, add 1 mL of 2.5% trypsin and bring it to a 5 mL volume with HBSS. Incubate for 15 min in the water bath at 37 °C.
      2. To dilute the trypsin, add 10 mL of pre-heated HBSS and incubate for 5 min in the water bath at 37 °C.
      3. Transfer the hippocampi that now appear as a mucus mass to a 50 mL tube with a 1,000 µL micropipette, add 6 mL of pre-heated HBSS, and incubate for 5 min in the water bath at 37 °C.
   2. Mechanical dissociation of the hippocampus
      1. Transfer the mass to a 2 mL tube with conical bottom, taking the minimal volume. Add 1 mL of pre-heated DMEM media. Homogenize the pellet with a 1,000 µL micropipette by gently aspirating up and down.
         NOTE: It is critical to avoid generating bubbles when aspirating as bubbles may lyse the cells.
      2. Repeat the up and down aspiration with a pre-pulled glass pipette (10x-20x), with its tip in contact with the conical bottom of the tube. Avoid generating bubbles. At the end of this step, the mixture must be translucent.
      3. Once homogenously mixed such that the mixture is translucent, transfer the mixture to a tube containing 4 mL of DMEM media at 37 °C and homogenize with a glass pipette by pipetting up and down.
3. Cell seeding
   1. Count the cells. The number of neurons in the solution can be counted according to standard laboratory procedures.
      NOTE: In imaging experiments for antibody detection and calcium activity, a concentration of 50,000 cells per 3.5 cm dish is optimal.
   2. Keeping the cells suspended, withdraw the calculated volume, and plate in 3.5 cm dishes containing PLL-coated coverslips or in the PLL-coated glass-bottom dishes.
   3. Evenly distribute the cells on the dishes by softly shaking in crossed movements (back and forth, then side to side; repeat as needed) and place the dishes in the CO₂ (5%) incubator.
      NOTE: It is important to use crossed movements to avoid the cells settling in the periphery of the dish.
   4. Every week, add approximately 1 mL of NB + B27, so the culture does not dry.
      NOTE: After 2 weeks (14 days in vitro [div]), the neurons are mature and express all factors to be used for experimentation. The total average number of neurons obtained using this protocol is approximately 2.5 x 10⁶ neurons coming from an average of 12 E18 embryos per pregnant rat.

   
PART 2: Using hippocampal neuronal cultures for antibody detection in neuronal cell surface proteins

​NOTE: This part of the protocol demonstrates how hippocampal cultures are used to identify anti-NMDAR antibodies in the serum and/or CSF of patients with anti-NMDAR encephalitis. Fluorescent immunostaining is used to visualize the reactivity in the live neurons, but other visualization methods could also be used. An appropriate control for this experiment would be serum or CSF from a healthy individual. When using human samples, keep in mind that approval from the institutional ethical committee may be required.

3. Live fluorescent immunostaining

1. Use 14 div cells that have been grown on coverslips (50,000 cells per 3.5 cm dish containing five coverslips).
2. Inside the hood, rinse the coverslips with NB pre-heated to 37 °C.
3. Add the sample that in this case contains anti-NMDAR antibodies (used as a primary antibody) diluted in NB media at 1:200 for serum or 1:2 for CSF and incubate 1 h at 37 °C (inside the CO₂ (5%) incubator).
4. Rinse carefully with PBS 3x at RT.
5. Add fixation solution (4% formaldehyde in PBS) and incubate at RT for 5 min.
   CAUTION: When handling 4% formaldehyde, work inside the hood and wear recommended personal protective equipment.
6. Wash 3x with PBS for 5 min each.
7. Add secondary antibody Goat anti-Human AF488 at 1:1000 dilution and incubate at RT for 1 h.
   NOTE: During the incubation, protect from light exposure by covering with aluminum foil.
8. Wash 3x with PBS for 5 min each. Rinse with distilled water
9. Mount the coverslips with a liquid mounting medium (e.g., antifading mounting media with DAPI; approximately 7 µL), aspirate any remaining liquid and rinse with distilled water. The neurons are now ready for fluorescence imaging.
   CAUTION: When handling mounting medium, wear the recommended personal protective equipment.

PART 3: Demonstration of antibody pathogenic effects using calcium imaging

NOTE: This part of the protocol demonstrates how to determine if the antibodies in patients have a functional effect on the cultures, which would suggest pathogenicity. In order to increase the significance of the effects, a pool of CSF from eight patients was used. Calcium activity of the live cultures was recorded upon chemical stimulation (NMDA + Glycine) using the GECI fluorescence calcium indicator (GCaMP5G). These studies require an inverted fluorescence microscope with a cell chamber that can maintain the required physiological conditions of the neurons.

4. Calcium imaging

1. Use 14-18 div cells that have been grown on coverslips (50,000 cells per 3.5 cm dish with five coverslips)
2. 1 week prior to imaging, add the viral vector pAAV2-CAG-GCaMP5G at 2.5 x 10¹⁰ GC/mL to the cells and incubate for 5-7 days.
   NOTE: This commercially available, pre-packaged AAV serotype 2 vector over-expresses genetically encoded calcium indicator GCaMP5G under a CAG promoter.
3. 1 day prior to imaging, add the patient sample (in this example, a pool of CSF diluted 1:25 in NB + B27) and incubate for 24 h. Always filtrate human samples (0.2 µm pore size) prior to use to avoid contamination.
   NOTE: For these studies, control CSF from healthy subjects needs to be run in parallel.
4. On the day of imaging, prepare the imaging setup and set the microscope cell chamber to 37 °C, fill the lane with distilled water, and infuse with 5% CO₂ to maintain cell physiological conditions.
5. In the hood, wash the cells from Step 3 with 3 mL of EPS pre-heated to 37 °C
6. Cover the cells with 2.45 mL of EPS and transfer them to the microscope cell chamber. Add NBQX (10 µM) to block the AMPA and KA receptors to visualize exclusively the NMDA receptor response.
7. In the inverted fluorescence microscope equipped with a mercury lamp and a FITC filter cube, acquire a 2 min movie with frames recorded every 100 ms (spontaneous activity).
   NOTE: A 20x NA 0.75 air objective was used, and images (512 x 512 pixels, 16-bit grayscale) were taken every 100 ms with a sCMOS camera.
8. Acquire the second movie of 4 min with frames recorded every 100 ms. Shortly after starting the acquisition, add stimulation solution (NMDA [100 µM] + Glycine [1 µM]) to the dish.
   NOTE: The addition of media into the dish will generate turbulence that could disrupt the imaging conditions (focus, fluorescence intensity, and/or unspecific background signal). Do not add more than 50 µL of solution into the filled 2.5 mL dish to decrease the probability of such alterations.
9. Using image processing software (ImageJ was used here), extract the fluorescence signal over time, obtained upon stimulation, from the cultures incubated with the patient and control CSF samples.
   NOTE: The GCaMP probes, when binding to free calcium ions, undergo a conformational change causing them to become brighter. Therefore, the increase in fluorescence intensity correlates with a calcium influx due to neuronal depolarization.
10. Determine the regions of interest (ROI) to analyze. Manually segment the somas of neurons and add the ROIs to the ROI manager (Analyze > Tools > ROI Manager > Add). Save the ROI from the ROI Manager menu (More > Save).
11. Set the measurements (Analyze > Set measurements) and select the Mean gray value. Extract the mean fluorescence intensity profile from the cell somas by clicking More > Multi Measure, and then save the table generated as a .xls spreadsheet.
12. Perform statistical analyses to compare the fluorescence between groups (patients' CSF vs. control's CSF).